Aerobic Utilization of Hydrocarbons, Oils, and Lipids [1st ed.] 978-3-319-50417-9, 978-3-319-50418-6

Citation preview

Handbook of Hydrocarbon and Lipid Microbiology

Series Editors: Kenneth N. Timmis (Editor-in-Chief) Matthias Boll · Otto Geiger · Howard Goldfine · Tino Krell Sang Yup Lee · Terry J. McGenity · Fernando Rojo Diana Z. Sousa · Alfons J. M. Stams · Robert Steffan · Heinz Wilkes

Fernando Rojo  Editor

Aerobic Utilization of Hydrocarbons, Oils, and Lipids

Handbook of Hydrocarbon and Lipid Microbiology Series Editors Kenneth N. Timmis (Editor-in-Chief) Emeritus Professor Institute of Microbiology Technical University Braunschweig Braunschweig, Germany Matthias Boll Institute of Biology/Microbiology University of Freiburg Freiburg, Germany Otto Geiger Centro de Ciencias Genómicas Universidad Nacional Autónoma de México Cuernavaca, Morelos, Mexico Howard Goldfine Dept. Microbiology University of Pennsylvania Philadelphia, PA, USA Tino Krell Department of Environmental Protection Estacion Experimental del Zaidin Granada, Granada, Spain Sang Yup Lee Dept.Chem.Engineer.&BioProcess Korea Adv.Inst.Science&Techn. Taejon, Korea (Republic of) Terry J. McGenity School of Biological Sciences University of Essex Colchester, UK

Fernando Rojo Centro Nacional de Biotecnología CSIC Madrid, Spain Diana Z. Sousa Laboratory of Microbiology Wageningen University Wageningen, The Netherlands Alfons J. M. Stams Laboratory of Microbiology Wageningen University Wageningen, The Netherlands Robert Steffan Cape Coral, FL, USA Heinz Wilkes ICBM Carl von Ossietzky University Oldenburg, Niedersachsen, Germany

This handbook is the unique and definitive resource of current knowledge on the diverse and multifaceted aspects of microbial interactions with hydrocarbons and lipids, the microbial players, the physiological mechanisms and adaptive strategies underlying microbial life and activities at hydrophobic material:aqueous liquid interfaces, and the multitude of health, environmental and biotechnological consequences of these activities. Scientific Advisory Board Victor de Lorenzo, Eduardo Diaz, Otto Geiger, Ian Head, Sang Yup Lee, Terry McGenity, Colin Murrell, Balbina Nogales, Roger Prince, Juan Luis Ramos, Wilfred Röling, Eliora Ron, Burkhard Tümmler, Jan Roelof van der Meer, Willy Verstraete, Friedrich Widdel, Heinz Wilkes and Michail Yakimov. More information about this series at http://www.springer.com/series/13884

Fernando Rojo Editor

Aerobic Utilization of Hydrocarbons, Oils, and Lipids With 149 Figures and 29 Tables

Editor Fernando Rojo Centro Nacional de Biotecnología CSIC Madrid, Spain

ISBN 978-3-319-50417-9 ISBN 978-3-319-50418-6 (eBook) ISBN 978-3-319-50419-3 (print and electronic bundle) https://doi.org/10.1007/978-3-319-50418-6 Library of Congress Control Number: 2018962938 © Springer Nature Switzerland AG 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

Both marine and terrestrial environments suffer frequent spills of hydrocarbons caused by the extraction and transportation of crude oil or its refined derivatives. Significant quantities of hydrocarbons, oils, and lipids also enter the environment via natural processes, including escape from oil reservoirs and production by living organisms. Many microbes have acquired the ability to degrade and assimilate these compounds and play a key role in their recycling. This book provides a broad view of those microbes that perform these actions under aerobic conditions (the anaerobic metabolism of these compounds is covered in a separate volume). Examples are provided of the breakdown of aliphatic and aromatic hydrocarbons, their halogenated and sulfated derivatives, lipids, fatty acids, and steroids. Different chapters focus on the enzymes involved, the metabolic pathways followed, the regulatory mechanisms that control the expression of the pathway genes, the physiology and genetic features of the microbes concerned, and the global environmental consequences of their activity. Most of the data available refer to bacteria, but some chapters discuss yeasts as well. The metabolic versatility of these microbes, their importance in the recycling of these organic compounds, and their role in environmental homeostasis are emphasized throughout. Such a broad topic cannot be covered on a single volume and naturally requires the several volumes that make up this series. I would like to express my gratitude to all the authors who contributed chapters to this book, to Series Editor Prof. Ken Timmis for his help and advice, and to the staff at Springer who helped bring this volume to light. December 2018

Fernando Rojo

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Contents

Part I 1

2

Biochemistry of Aerobic Degradation of Hydrocarbons . . . .

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Widdel and F. Musat

3

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization: An Introduction . . . . . . . . . . . . . . . . . . . F. Widdel and F. Musat

33

3

Physiology and Biochemistry of the Aerobic Methanotrophs . . . . Valentina N. Khmelenina, J. Colin Murrell, Thomas J. Smith, and Yuri A. Trotsenko

4

Biochemistry and Molecular Biology of Methane Monooxygenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tim Nichol, J. Colin Murrell, and Thomas J. Smith

73

99

5

Enzymes for Aerobic Degradation of Alkanes in Bacteria . . . . . . . Renata Moreno and Fernando Rojo

117

6

Enzymes for Aerobic Degradation of Alkanes in Yeasts Ryouichi Fukuda and Akinori Ohta

........

143

7

Aerobic Degradation of Aromatic Hydrocarbons . . . . . . . . . . . . . . D. Pérez-Pantoja, B. González, and Dietmar H. Pieper

157

8

Biosynthesis and Insertion of Heme . . . . . . . . . . . . . . . . . . . . . . . . Katrin Müller, Toni Mingers, V. Haskamp, Dieter Jahn, and Martina Jahn

201

Part II 9

Biochemistry of Aerobic Degradation of Lipids

.........

Membrane Lipid Degradation and Lipid Cycles in Microbes . . . . Diana X. Sahonero-Canavesi, Isabel M. López-Lara, and Otto Geiger

229 231

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Contents

10

Classification of Lipolytic Enzymes from Bacteria . . . . . . . . . . . . . Filip Kovacic, Nikolina Babic, Ulrich Krauss, and Karl-Erich Jaeger

255

11

Pathways for the Degradation of Fatty Acids in Bacteria Lorena Jimenez-Diaz, Antonio Caballero, and Ana Segura

.......

291

12

Bacterial Metabolism of Steroids . . . . . . . . . . . . . . . . . . . . . . . . . . Beatriz Galán, Julia García-Fernández, Carmen Felpeto-Santero, Lorena Fernández-Cabezón, and José L. García

315

13

Aerobic Degradation of Chloroaromatics . . . . . . . . . . . . . . . . . . . . Dietmar H. Pieper, B. González, B. Cámara, D. Pérez-Pantoja, and W. Reineke

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14

Structure-Function Relationships and Engineering of Haloalkane Dehalogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Piia Kokkonen, Tana Koudelakova, Radka Chaloupkova, Lukas Daniel, Zbynek Prokop, and Jiri Damborsky

367

15

Aerobic Degradation of Gasoline Ether Oxygenates Michael Hyman

...........

389

16

Bacterial Metabolism of C1 Sulfur Compounds . . . . . . . . . . . . . . . Rich Boden and Lee P. Hutt

421

17

Aerobic Bacterial Catabolism of Dimethylsulfoniopropionate . . . . Rich Boden and Lee P. Hutt

465

18

Chemolithoheterotrophy: Means to Higher Growth Yields from This Widespread Metabolic Trait . . . . . . . . . . . . . . . . . . . . . Rich Boden and Lee P. Hutt

Part III Genetics and Functional Genomics of Aerobic Degradation of Hydrocarbons and Lipids . . . . . . . . . . . . . . . . . . . . . 19

20

493

519

Genetic Features and Regulation of n-Alkane Metabolism in Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Renata Moreno and Fernando Rojo

521

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ryouichi Fukuda and Akinori Ohta

543

21

Genetics and Ecology of Isoprene Degradation . . . . . . . . . . . . . . . Andrew T. Crombie, Nasmille L. Mejia-Florez, Terry J. McGenity, and J. Colin Murrell

22

Current View of the Mechanisms Controlling the Transcription of the TOL Plasmid Aromatic Degradation Pathways . . . . . . . . . . Patricia Domínguez-Cuevas and Silvia Marqués

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Contents

23

Genetics and Biochemistry of Biphenyl and PCB Biodegradation . . . Loreine Agulló, Dietmar H. Pieper, and Michael Seeger

24

An Update on the Genomic View of Mycobacterial High-Molecular-Weight Polycyclic Aromatic Hydrocarbon Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ohgew Kweon, Seong-Jae Kim, and Carl E. Cerniglia

25

Degradation of Aromatic Compounds in Pseudomonas: A Systems Biology View . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . J. Nogales, José L. García, and E. Díaz

26

Phylogenomics of Aerobic Bacterial Degradation of Aromatics . . . D. Pérez-Pantoja, R. Donoso, H. Junca, B. González, and Dietmar H. Pieper

27

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

28

Regulation of Fatty Acids Degradation in Bacteria . . . . . . . . . . . . Lorena Jimenez-Diaz, Antonio Caballero, and Ana Segura

29

Genetics and Molecular Features of Bacterial Dimethylsulfoniopropionate (DMSP) and Dimethyl Sulfide (DMS) Transformations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . J. M. González, A. W. B. Johnston, M. Vila-Costa, and A. Buchan

30

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

Part IV Global Activities, Global Consequences of Aerobic Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31

ix

595

623

639 689

737 751

773

785

795

Global Aerobic Degradation of Hydrocarbons in Aquatic Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sara Kleindienst and Samantha B. Joye

797

Global Consequences of the Microbial Production and Consumption of Inorganic and Organic Sulfur Compounds . . . . . Donovan P. Kelly and Ann P. Wood

815

Potential for Microbial Interventions to Reduce Global Warming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Donovan P. Kelly and Ann P. Wood

827

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

841

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About the Series Editor-in-Chief

Kenneth N. Timmis Emeritus Professor Institute of Microbiology Technical University Braunschweig Braunschweig, Germany Kenneth Timmis studied microbiology and obtained his Ph.D. at Bristol University. He undertook postdoctoral 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 and subsequently 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), and concomitantly Professor of Microbiology in the Institute of Microbiology of the Technical University Braunschweig. He is currently Emeritus Professor in this institute. The Editor-in-Chief has worked for more than 30 years in the area of environmental microbiology and biotechnology, has published over 400 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, and on the ecology of hydrocarbon-degrading microbial communities,

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About the Series Editor-in-Chief

discovered the new group of marine oil-degrading hydrocarbonoclastic bacteria, initiated genome sequencing projects on bacteria that are paradigms of microbes that degrade organic compounds (Pseudomonas putida and Alcanivorax borkumensis), and pioneered the topic of experimental evolution of novel catabolic activities. He is Fellow of the Royal Society, Member of the European Molecular Biology Organisation, Fellow of the American Academy of Microbiology, Member of the European Academy of Microbiology, and Recipient of the Erwin Schrödinger Prize. He is the founder and Editor-in-Chief of the journals Environmental Microbiology, Environmental Microbiology Reports, and Microbial Biotechnology.

About the Volume Editor

Fernando Rojo Research Professor and Director of the National Center of Biotechnology Madrid, Spain Fernando Rojo studied biochemistry and molecular biology and obtained his Ph.D. degree at the Autonomous University of Madrid, Spain. He undertook postdoctoral training at the Department of Medical Biochemistry of the University of Geneva, Switzerland, and at the Center of Molecular Biology “Severo Ochoa” (CBMSO-CSIC), Madrid, Spain. His work has been devoted to the analysis and modification of bacterial pathways for the degradation of hydrocarbons and toxic compounds and to the elucidation of molecular mechanisms of gene expression in bacteria. During the last 20 years, his research group has been focused on the analysis of the global regulation networks that allow coordinating metabolism in pseudomonads. He has published over 130 papers in international journals. He is currently Research Professor of CSIC and Director of the National Centre of Biotechnology (CNB-CSIC).

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Contributors

Loreine Agulló Laboratorio de Microbiología Molecular y Biotecnología Ambiental, Department of Chemistry and Center for Nanotechnology and Systems Biology and Centro de Biotecnología, Universidad Técnica Federico Santa María, Valparaíso, Chile Nikolina Babic Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Jülich, Germany Rich Boden School of Biological and Marine Sciences, Sustainable Earth Institute, Faculty of Science and Engineering, University of Plymouth, Plymouth, UK A. Buchan Department of Continental Ecology-Limnology, CSIC, Centre d’Estudis Avançats de Blanes, Blanes, Spain Department of Microbiology, University of Tennessee, Knoxville, TN, USA Antonio Caballero Abengoa Research, Sevilla, Spain Bacmine, Tres Cantos, Spain B. Cámara Laboratorio de Microbiología Molecular y Biotecnología Ambiental, Departamento de Química and Centro de Biotecnología, Universidad Técnica Federico Santa María, Valparaíso, Chile Carl E. Cerniglia Division of Microbiology, National Center for Toxicological Research, Food and Drug Administration, Jefferson, AR, USA Radka Chaloupkova Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic Andrew T. Crombie School of Biological Sciences, University of East Anglia, Norwich, UK

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Contributors

Jiri Damborsky Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic Lukas Daniel Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic E. Díaz Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain Patricia Domínguez-Cuevas Department of Biology, University of Copenhagen, Copenhagen, Denmark R. Donoso Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile Carmen Felpeto-Santero Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain Lorena Fernández-Cabezón Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain Ryouichi Fukuda Department of Biotechnology, The University of Tokyo, Tokyo, Japan Beatriz Galán Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain José L. García Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain Julia García-Fernández Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain Otto Geiger Centro de Ciencias Genómicas, Universidad Nacional Autónoma de México, Cuernavaca, Morelos, Mexico B. González Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile

Contributors

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J. M. González Department of Microbiology, University of La Laguna, Tenerife, Spain V. Haskamp Institute of Microbiology, Braunschweig University of Technology, Braunschweig, Germany Lee P. Hutt School of Biological and Marine Sciences, Faculty of Science and Engineering, University of Plymouth, Plymouth, UK Michael Hyman Department of Plant and Microbial Biology, North Carolina State University, Raleigh, NC, USA Karl-Erich Jaeger Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Jülich, Germany Institute of Bio- and Geosciences IBG-1: Biotechnology, Forschungszentrum Jülich GmbH, Jülich, Germany Dieter Jahn Institute of Microbiology, Braunschweig University of Technology, Braunschweig Integrated Center of Systems Biology BRICS, Braunschweig, Germany Martina Jahn Institute of Microbiology, Braunschweig University of Technology, Braunschweig, Germany Lorena Jimenez-Diaz Abengoa Research, Sevilla, Spain A. W. B. Johnston School of Biological Sciences, University of East Anglia, Norwich, UK Samantha B. Joye Department of Marine Sciences, University of Georgia, Athens, GA, USA H. Junca Research Group Microbial Ecology: Metabolism, Genomics and Evolution, Microbiomas Foundation, Chia, Colombia Donovan P. Kelly School of Life Sciences, University of Warwick, Coventry, UK Valentina N. Khmelenina G.K. Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino/Moscow, Russia Seong-Jae Kim Division of Microbiology, National Center for Toxicological Research, Food and Drug Administration, Jefferson, AR, USA Sara Kleindienst Center for Applied Geosciences, Eberhard Karls University, Tübingen, Tübingen, Germany Piia Kokkonen Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic

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Contributors

Tana Koudelakova Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic Filip Kovacic Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Jülich, Germany Ulrich Krauss Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Jülich, Germany Ohgew Kweon Division of Microbiology, National Center for Toxicological Research, Food and Drug Administration, Jefferson, AR, USA Isabel M. López-Lara Centro de Ciencias Genómicas, Universidad Nacional Autónoma de México, Cuernavaca, Morelos, Mexico Silvia Marqués Department of Environmental Protection, CSIC, Estación Experimental del Zaidín, Granada, Spain Terry J. McGenity School of Biological Sciences, University of Essex, Colchester, UK Nasmille L. Mejia-Florez School of Environmental Sciences, University of East Anglia, Norwich, UK Toni Mingers Institute of Microbiology, Braunschweig University of Technology, Braunschweig, Germany Renata Moreno Centro Nacional de Biotecnología, CSIC, Madrid, Spain Katrin Müller Institute of Microbiology, Braunschweig University of Technology, Braunschweig, Germany J. Colin Murrell School of Environmental Sciences, University of East Anglia, Norwich Research Park, Norwich, UK F. Musat UFZ – Helmholtz Centre for Environmental Research, Leipzig, Germany Tim Nichol Biomolecular Sciences Research Centre, Sheffield Hallam University, Sheffield, UK J. Nogales Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain Akinori Ohta Department of Biological Chemistry, College of Bioscience and Biotechnology, Chubu University, Kasugai, Aichi, Japan D. Pérez-Pantoja Departamento de Bioquímica y Biología Molecular, Facultad de Ciencias Biológicas, Universidad de Concepción, Concepción, Chile

Contributors

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Dietmar H. Pieper Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany Zbynek Prokop Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic J.-L. Ramos Department of Environmental Protection, Consejo Superior de Investigaciones Científicas, Estación Experimental del Zaidín, Granada, Spain W. Reineke Chemical Microbiology, Bergische Universität Wuppertal, Wuppertal, Germany Fernando Rojo Centro Nacional de Biotecnología, CSIC, Madrid, Spain Diana X. Sahonero-Canavesi Centro de Ciencias Genómicas, Universidad Nacional Autónoma de México, Cuernavaca, Morelos, Mexico Michael Seeger Laboratorio de Microbiología Molecular y Biotecnología Ambiental, Department of Chemistry and Center for Nanotechnology and Systems Biology and Centro de Biotecnología, Universidad Técnica Federico Santa María, Valparaíso, Chile Ana Segura Abengoa Research, Sevilla, Spain Department of Environmental Protection, Consejo Superior de Investigaciones Científicas, Estación Experimental del Zaidín, Granada, Spain V. Shingler Department of Molecular Biology, Umeå University, Umeå, Sweden Thomas J. Smith Biomolecular Sciences Research Centre, Sheffield Hallam University, Sheffield, UK Yuri A. Trotsenko G.K. Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino/Moscow, Russia P. van Dillewijn Department of Environmental Protection, Consejo Superior de Investigaciones Científicas, Estación Experimental del Zaidín, Granada, Spain M. Vila-Costa Department of Marine Sciences, University of Georgia, Athens, GA, USA F. Widdel Max Planck Institute for Marine Microbiology, Bremen, Germany R.-M. Wittich Department of Environmental Protection, Consejo Superior de Investigaciones Científicas, Estación Experimental del Zaidín, Granada, Spain Ann P. Wood Department of Biochemistry, King’s College London, London, UK

Part I Biochemistry of Aerobic Degradation of Hydrocarbons

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An Introduction F. Widdel and F. Musat

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Functionalization of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Aerobic Activation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Dioxygen as a Reactant . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Some Common Principles of Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Heme-Iron Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Non-Heme Di-Iron Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Particulate Methane Monooxygenase (Particulate Di-Metal or Tri-Metal Monooxygenase) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Non-Heme Mono-Iron Oxygenases (Ring Hydoxylating Dioxygenases) . . . . . . . . . . . . 3.7 Flavin-Containing Monooxygenases, and Monooxygenases with Unknown Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Anaerobic Activation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Anaerobic Activation of Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Glycyl Radical Enzymes for Anaerobic Activation of Non-Methane Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Ethylbenzene Dehydrogenase, a Molybdo-Enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Unknown and Hypothesized Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

4 5 6 6 9 12 14 16 16 18 19 20 22 24 24 25 27

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 F. Widdel (*) Max Planck Institute for Marine Microbiology, Bremen, Germany e-mail: [email protected] F. Musat UFZ – Helmholtz Centre for Environmental Research, Leipzig, Germany # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_50

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F. Widdel and F. Musat

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 σ-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

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An. . .

5

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 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 ðC
H ! C
XÞ. Homolytic C–H-bond cleavage, viz. formation of a carbon radical ðC
H ! C • þ H • Þ, is most common. The energy for homolytic C–Hbond cleavage at sp3 carbon atoms of various hydrocarbons (in the gas phase ca. between 360 and 400 kJ mol
1), which determines the activation energy, depends on the carbon compound (for compilation of energies of homolytic C–H-bond cleavage see Chap. 2, ▶ “Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization: An Introduction” in this volume; Widdel et al. 2007). Homolytic cleavage is facilitated if the carbon radical is stabilized (delocalized) by an adjacent π-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 ð> Cþ
Þ ion gains enough stability via adjacent carbon atoms and a π-electron system. In this case, an HO
-ion may be added subsequently (see Sect. 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
 ( Ni3þ
CH3 ; see Sect. 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 mol
1). 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 σ-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

6

F. Widdel and F. Musat

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 ðAR
H ! AR
OHÞ. 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 ðAR
H þ Xþ ! AR
X þ Hþ Þ, 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.

3

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 ðO2 =2H2 O : E ¼ þ1:23 V, E0 ¼ þ0:818 VÞ 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 ð“CH3
CH3 þ O2 ! CH2 OH
CH2 OH”Þ would have a ΔG of
290 and a ΔH 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 (3Σ g O2), the paramagnetic di-radical (Fig. 1). A concerted combination of these electrons with those of organic compounds in their common singlet state is extremely unlikely (“spin-forbidden”;

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An. . .

7

Fig. 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 σ-bond between the nuclei are usually designated as 2px orbitals. This leaves 2py and 2pz to form the π-bonds. However, conventions may differ, and the atomic orbitals along the connecting axis may be also termed 2pz, leaving 2px and 2py for the π-bonds

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 (1Δg 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 ð0Þ. 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). 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, 1

Chemical generation of singlet dioxygen is difficult to achieve. An example is the oxidation of  hydrogen peroxide with hypochlorous acid H2 O2 þ HClO ! H2 O þ1 Δg O2 þ Cl
þ Hþ .

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F. Widdel and F. Musat

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 (O2 =O2 •
, E ¼
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 ðR • þ O2 •
þ Hþ ! R
O
OHÞ. 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 to an organic peroxide that disintegrates into the two carbon compounds (Lorimer 1981). (3) Superoxide can be further reduced to
inorganic peroxide O2 2 ; (Fig. 1); the free form of the latter is usually the fully protonated one, H2 O2 ðH2 O2 =HO2
, pKa ¼ 11:8Þ. Hydrogen peroxide formation has a high redox potential ðO2 •
þ 2Hþ =H2 O2 ; E0 ¼ þ0:89 VÞ. 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–Obond. 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 Fe2þ as reductant, is usually formulated with the hydroxyl radical as product ðFe2þ þ H2 O2 þ Hþ ! Fe3þ þ HO • þ H2 OÞ. However, the prevalent form may be an iron(IV)-oxo (ferrylIV) species in equi2þ 2þ librium with the hydroxyl radical ( ½FeIII
OH þ • OH Ð ½FeIV ¼ O þ H2 O; 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

Another redox potential often indicated, E ¼
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 ¼ 7 does not involve a proton, because superoxide is deprotonated ðO2 •
=HO2 • , pKa ¼ 4:6Þ.

2

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An. . .

9

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 pterinbound 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 non-hydrocarbon (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 substrate-binding site of an alkane monooxygenase. 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; Hence, the formal oxidation state of the carbon changes by þII. 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 ð
CH3 ,
IIIÞ is converted to a hydroxymethyl group ð
CH2 OH,
IÞ. In an aromatic hydrocarbon, the (formally localized) “vinylen” ð─CH ¼ CH─, 2 
I ¼
IIÞ can be dioxygenated yielding a hydrodiol ð
CHOH
CHOH
, 2  0 ¼ 0Þ and a non-aromatic ring, or can be monooxygenated yielding an “enol” ð
CH ¼ COH
,
I þ I ¼ 0Þ with maintenance of the 3

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F. Widdel and F. Musat

Table 1 Overview of aerobic hydrocarbon activation in living organisms Hydrocarbon(s) Methane

Short-chain non-methane alkanes (C2 to ca. C10) Long-chain non-methane alkanes, ðca: > C10 Þ

Organisms Only bacteria (aerobic methanotrophs)

Bacteria

Bacteria

Yeasts (mainly Candida sp.) Animals

Alkenes

Bacteria

Aromatic hydrocarbons

Bacteria

Some filamentous fungi (e.g., Cunninghamella) Mammals

Enzyme type for activation [Fe2]Monooxygenase, soluble (sMMO) [Cu3]Monooxygenase, particulate (pMMO) [Fe2]Monooxygenase

[Heme]Monooxygenase (P450-type) [Fe2]Monooxygenase [Flavin]Monooxygenase [Heme]Monooxygenase (P450-type) [Heme]Monooxygenase (P450-type) [Fe2]Monooxygenase [Fe]-Diooxygenase

[Fe2]Monooxygenase [Flavin]Monooxygenase [Heme]Monooxygenase (P450-type) [Heme]Monooxygenase (P450-type)

Remarks In few species of aerobic methanotrophs In essentially all species of aerobic methanotrophs

In bacteria probably most common One case reported; mechanism unknown Yeasts may use alkanes as real growth substrate By-reaction (“detoxification”) without further oxidation. Epoxide formation Very common for aromatic hydrocarbon utilization; formation of cis-hydrodiols Mono-hydroxylation of ring or alkyl side chain Epoxidation of styrene at the side chain Mono-hydoxylation of ring, or formation of epoxides yielding trans-hydrodiols. Role as sole growth substrate uncertain Formation of epoxides; may yield trans-hydrodiols. By-reaction (“detoxification”) without further oxidation.

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)

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Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An. . .

11




Fig. 2 Principle of hydrocarbon oxygenation. O2 is always reduced to a bound peroxide state, ½O2 2 ; 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

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 Hþ þ 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 redoxactive 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.

12

F. Widdel and F. Musat

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 2008; 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 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 ðFeIV ¼ OÞ hosted in an oxidized porphyrin (lacking one π-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 omegahydroxylation 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 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 Named after the National Institute of Health where this hydrogen shift was first detected (Guroff et al. 1967).

4

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An. . .

13

Fig. 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 π–cation radical ðPor • þ Þ; the system is isoelectronic with Por
FeV ¼ O. The favored mechanism is hydroxyl group formation via H-atom abstraction ðR  H  O ¼ FeIV Por • þ Þ , 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 0 would be Por . Heme formation by chelation of Fe2þ , (which needs removal of 2 Hþ from 0




tetrapyrrole-nitrogen) leads to ½Por2 Fe2þ 

. Binding of O2 followed by reduction with h i one


, and “internal” electron from Fe2þ and one external electron leads to Por2 Fe3þ ðO
OÞ2


0

protonation to ½Por2 Fe3þ ðO
OHÞ
 . Addition of another Hþ and elimination of H2O yields
2
5þ
þ . “Internal” donation of a π–electron from the porphyrin ring to the iron leads Por Fe ¼ O2
þ to Por • þ=2
Fe4þ ¼ O2
. If the axial cysteine thiolate ligand is included, the complex is

 0 RS Por • þ=2 Fe4þ ¼ O2
. Dissociation of the porphyrin carboxyl groups would yield

2
2
RS Por • þ=4
Fe4þ ¼ O

without an expoxide as intermediate (Fig. 4b; 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 et al. 1982; Perbellini et al. 1980).

14

F. Widdel and F. Musat

Fig. 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

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 (Käppeli 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 P450 enzymes in hydrocarbon activation. There are soluble (Leahy et al. 2003; Lippard 2005) and membrane-bound di-iron monooxygenases.

1

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons: An. . .

15

Fig. 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

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, methyl- substituted 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 alcohol or methylbenzyl alcohols) encoded by the TOL plasmid in the same organism also involves a diiron center (Austin et al. 2003).

16

F. Widdel and F. Musat

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 (Bédard 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 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 membranebound ammonia monooxygenase (AMO5) of ammonia-oxidizing bacteria (Bédard 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 non-aromatic cis-hydrodiols is the most common activation mechanism in the degradation of aromatic hydrocarbons. Numerous ring-hydroxylating dioxygenases have been described and classified (see ▶ Chaps. 7, “Aerobic Degradation of Aromatic Hydrocarbons” by Pérez-Pantoja et al., and ▶ 13, “Aerobic Degradation of Chloroaromatics” by D. H. Pieper, in this volume; Kweon et al. 2008; Reineke 2001). 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 5

Abbreviation AMO must not be confused with the same one sometimes used for alkene monooxygenase.

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Fig. 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)

Fig. 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 ðAR þ O ¼ FeV
OH! • AR
O
FeIV
OH ! HO
AR
O
FeIII Þ or formation of an arene expoxide as intermediate

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F. Widdel and F. Musat

single hydroxyl group into the hydrocarbons indene (fused aromatic six- and fivemembered 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 (Haddock 2010; chapter ▶ “Aerobic Degradation of Aromatic Hydrocarbons”, 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 flavin4a-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 (Haddock 2010; 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 ðC6 H5
CH ¼ CH2 Þ 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). A surprising finding was the recovery of a flavin-monooxygenase from an alkane-degrading 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 long-chain 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 longchain 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

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Fig. 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 non-hydrocarbons, 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
þ O2 ! FADH • þO2 •
, 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 π-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 σ-bonds is matter of discussion

and mechanism of the reaction need 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 co-reactants 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

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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) Methane

Non-methane alkanes

Organisms Archaea (assumed syntrophism with SO4 2
-reducing bacteria) Bacteria (NO3
reducing) Bacteria

Enzyme type for activation Ni-porphinoid enzyme (closely related to CH3–coenzyme M reductase) Enzyme unknown Glycyl radical enzyme Unknown enzyme

Alkenes

Bacteria

Unknown enzyme

Aromatic hydrocarbons Benzene, naphthalene, phenanthrene

Bacteria

Unknown enzymes

Methylbenzenes

Bacteria

Glycyl radical enzyme

Ethylbenzene

Bacteria, SO4 2
reducing Bacteria, NO3
reducing

Glycyl radical enzyme

Ethylbenzene, n-propylbenzene

Mo-cofactorcontaining dehydrogenase

Remarks Assumed reversal of the final step in methanogenesis Different from the final step in methanogenesis Leads to a substituted succinate Apparent activation at carbon-3 Introduction of an HO-group

Activation results in addition of a COO
group (methylation also discussed) Leads to a substituted succinate Leads to a substituted succinate Leads to (S)-1phenylethanol or (S)-1phenylpropanol

discovery and 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 (Taupp et al. 2010). 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 sulfate-reducing bacteria of the Deltaproteobacteria. These associations are

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Fig. 9 Hypothesized anaerobic activation of methane by a thiyl radical vs. activation by a highvalent nickel (drawn according to discussions by Krüger 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 ðCoB
S • Þ 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 þ ; not depicted) to the coenzyme M thiolate to form methyl-coenzyme

usually interpreted as a 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 SO4 2
þ þ H ! HCO3 þ H2 S þ H2 O. 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 nickel-porphinoid 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 (Krüger et al. 2003), or (b) a reaction of cofactor F430 in its Ni(III) state as a strong electrophile to form

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F. Widdel and F. Musat

Fig. 10 Relationships of the assumed anaerobic methane-activating enzyme (reverse methylcoenzyme M reductases, rMcr) in anaerobic methanotrophs (bold) to regular methyl-coenzyme M reductase (Mcr) in methanogenic archaea (drawn according to Hallam et al. 2003; Krüger et al. 2003; Lösekann 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 (Krüger 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 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). A different mode of methane activation may take place in a highly enriched methane-oxidizing 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 radicalcatalyzed 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 NonMethane 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

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Fig. 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

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 reaction (Fig. 11) has been studied in detail with the toluene-activating benzylsuccinate synthase from denitrifying Betaproteobacteria (Boll et al. 2018; 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 carbon-2 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).

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Fig. 12 Relationships of anaerobic hydrocarbon-activating enzymes (bold) to other glycyl radical enzymes. (Modified from Musat et al. 2009)

4.3

Ethylbenzene Dehydrogenase, a Molybdo-Enzyme

Ethylbenzene in denitrifying bacteria is activated by dehydrogenation (anaerobic hydroxylation) to 1-phenylethanol (Boll et al. 2018; Johnson et al. 2001; Kniemeyer and Heider 2001) rather than by addition to fumarate, as in sulfatereducing bacteria (see above). The three-dimensional structure of the enzyme, ethylbenzene dehydrogenase, has been resolved and a detailed mechanism has been suggested (Boll et al. 2018; 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

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25

Fig. 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 Boll et al. 2018). 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

were based on labeling studies and the identification of aromatic hydrocarbonderived carboxylic acids with CO2-derived 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 naphthalenedegrading sulfate-reducing bacteria, methylation was essentially excluded (Musat and Widdel 2009; Musat et al. 2009). 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 ¼ CH2 þ H2 O ! R
CH2
CH2 OH; Schink 1985). Water addition would be particularly favored by involvement of a tertiary carbon atom, as occurring with monoterpenes (Harder 2010). In principle, also anaerobic hydroxylation at the carbon atom next to the double bond (the allylic carbon) appears possible ð
CH2
CH ¼ CH
þ H2 O !
CHOH
CH ¼ CH
þ 2 ½HÞ, 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

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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; Funhoff et al. 2006). 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. 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 radical-catalyzed 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 hydrocarbon-activating enzymes from ancestors that activate other substrates are of basic interest.

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Hallam SJ, Girguis PR, Preston CM, Richardson PM, DeLong EF (2003) Identification of methyl coenzyme M reductase A (mcrA) genes associated with methane-oxidizing Archaea. Appl Environ Microbiol 69:5483–5491 Halliwell B, Gutteridge JM (1984) Oxygen toxicity, oxygen radicals, transition metals and disease. Biochem J 219:1–14 Han W-G, Noodleman L (2008) Structural model for the peroxo intermediate P and the reaction pathway from P ! Q of methane monooxygenase using broken-symmetry density functional calculations. Inorg Chem 47:2975–2986 Harder J (2010) Anaerobic Degradation of Isoprene-Derived Compounds. In: Timmis KN (ed) Handbook of Hydrocarbon and Lipid Microbiology. Springer, Berlin, Heidelberg Harmer J, Finazzo C, Piskorski R, Ebner S, Duin EC, Goenrich M, Thauer RK, Reiher M, Schweiger A, Hinderberger D, Jaun B (2008) A nickel hydride complex in the active site of methyl-coenzyme M reductase: implications for the catalytic cycle. J Am Chem Soc 130:10907–10920 Hedegaard J, Gunsalus IC (1965) Mixed function oxidation. J Biol Chem 240:4038–4043 Heider J (2007) Adding handles to unhandy substrates: anaerobic hydrocarbon activation mechanisms. Curr Opin Chem Biol 11:188–194 Isin EM, Guengerich EP (2008) Substrate binding to cytochrome P450. Anal Bioanal Chem 392:1019–1030 Johnson HA, Pelletier DA, Spormann AM (2001) Isolation and characterization of anaerobic ethylbenzene dehydrogenase, a novel Mo-Fe-S enzyme. J Bacteriol 183:4536–4542 Käppeli O (1986) Cytochromes P-450 of yeasts. Microbiol Rev 50:244–258 Karlsson A, Parales JV, Parales RE, Gibson DT, Eklund H, Ramaswamy S (2003) Crystal structure of naphthalene dioxygenase: side-on binding of dioxygen to iron. Science 299:1039–1042 Kniemeyer O, Heider J (2001) Ethylbenzene dehydrogenase, a novel hydrocarbon-oxidizing molybdenum/iron-sulfur/heme enzyme. J Biol Chem 276:21381–21386 Kniemeyer O, Fischer T, Wilkes H, Glöckner F-O, 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 sulphate-reducing bacteria. Nature 449:898–902 Kovaleva EG, Neibergall MB, Chakrabarty S, Lipscomb JD (2007) Finding intermediates in the O2 activation pathways of non-heme iron oxgenases. Acc Chem Res 40:475–483 Krüger M, Meyerdierks A, Glöckner 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 Kweon O, Kim S-J, Baek S, Chae J-C, Adjei MD, Baek D-H, Kim Y-C, Cerniglia CE (2008) A new classification system for bacteria Rieske non-heme iron aromatic ring-hydroxylating oxygenases. BMC Biochem. doi:10.1186/1471-2091-9/11 Leahy JG, Batchelor PJ, Morcomb SM (2003) Evolution of the soluble diiron monooxygenases. FEMS Microbiol Rev 27:449–479 Li L, Liu X, Yang W, Xu F, Wang W, Feng L, Barlam M, Wang L, Rao Z (2008) Crystal structure of long-chain alkane monooxygenase (LadA) in complex with coenzyme FMN: unveiling the long-chain alkane hydroxylase. J Mol Biol 376:453–465 Liebermann RL, Rosenzweig AC (2005) Crystal structure of membrane-bound metalloenzyme that catalyzes the biological oxidation of methane. Nature 434:177–182 Lippard SJ (2005) Hydroxylation of C-H bonds at carboxylate-bridged diiron centres. Philos Trans R Soc Lond A 363:861–877 Lipscomb JD (1994) Biochemistry of the soluble methane monooxygenase. Annu Rev Microbiol 48:371–399 Lorimer GH (1981) The carboxylation and oxygenation of ribulose-1,5-bisphosphate: the primary events in photosynthesis and photorespiration. Annu Rev Plant Physiol 32:349–383

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Lösekann 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 73:3348–3362 Maeng JH, Sakai Y, Tani Y, Kato N (1996) Isolation and characterization of a novel oxygenase that catalyzes the first step of n-alkane oxidation in Acinetobacter sp. strain M-1. J Bacteriol 178:3695–3700 Martinho M, Choi DW, Dispirito AA, Antholine WE, Semrau JD, Münck E (2007) Mössbauer studies of the membrane-associated methane monooxygenase from Methylococcus capsulatus Bath: evidence for a diiron center. J Am Chem Soc 129:15783–15785 Morohashi K, Sadano H, Okada Y, Omura T (1982) Position specificity in n-hexane hydroxylation by two forms of cytochrome P-450 in rat liver microsomes. J Biochem 93:413–419 Munro AW, Girvan HM, McLean KJ (2007) Variations on a theme – novel mechanisms, redox partners and catalytic functions in the cytochrome P450 superfamily. Nat Prod Rep 24:585–609 Murrell JC, Gilbert B, McDonald IR (2000) Molecular biology and regulation of methane monooxygenase. Arch Microbiol 173:325–332 Musat F, Widdel F (2009) Anaerobic degradation of benzene by a marine sulfate-reducing enrichment culture, and cell hybridization of the dominant phylotype. Environ Microbiol 10:10–19 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 sulfate-reducing bacteria. Environ Microbiol 11:209–219 Oku Y, Ohtaki A, Kamitori S, Nakamura N, Yohda M, Ohno H, Kawarabayasi Y (2004) Structure and direct electrochemistry of cytochrome P450 from the thermoacidophilic crenarchaeon, Sulfolobus tokodaii strain 7. J Inorg Biochem 98:1194–1199 Perbellini L, Brugnone F, Pavan I (1980) Identification of the metabolites of n-hexane, cyclohexane, and their isomers in men’s urine. Toxicol Appl Pharmacol 53:220–229 Poulos TL, Finzel BC, Howard AJ (1987) High-resolution crystal structure of cytochrome P450cam. J Mol Biol 195:687–700 Powley MW, Carlson GP (2000) Cytochrome P450 involved with benzene metabolism in hepatic and pulmonary microsomes. J Biochem Mol Toxicol 14:303–309 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 Raghoebarsing AA, Pol A, van de Pas-Schoonen KT, Smolders AJ, Ettwig KF, Rijpstra WI, Schouten S, Damsté JS, Op den Camp HJ, Jetten MS, Strous M (2006) A microbial consortium couples anaerobic methane oxidation to denitrification. Nature 440:918–921 Reineke W (2001) Aerobic and anaerobic biodegradation potentials of microorganisms. In: Beek B (ed) The handbook of environmental chemistry, vol. 2, part K, Biodegradation and persistence. Springer, Berlin, pp. 1–161. Resnick SM, Lee K, Gibson DT (1996) Diverse reactions catalyzed by naphthalene dioxygenase from Pseudomonas sp strain NCIB 9816. J Ind Microbiol 17:438–457 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 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 Sawyer DT (1981) How super is superoxide? Acc Chem Res 14:393–400 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 Scheller U, Zimmer T, Kärgel E, Schunck W-H (1996) Characterization of the n-alkane and fatty acid hydroxylating cytochrome P450 forms 52A3 and 52A4. Arch Biochem Biophys 328:245–254

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Schink B (1985) Degradation of unsaturated hydrocarbons by methanogenic enrichment cultures. FEMS Microbiol Lett 31:69–77 Schlichting I, Berendzen J, Chu K, Stock AM, Maves SA, Benson DE, Sweet RM, Ringe D, Petsko GA, Sligar S (2000) The catalytic pathway of cytochrome P450cam at atomic resolution. Science 287:1615–1622 Seiffert GB, Ullmann GM, Messerschmidt A, Schink B, Kroneck PMH (2007) Structure of the nonredox-active tungsten/[4Fe:4S] enzyme acetylene hydratase. Proc Natl Acad Sci USA 104:3073–3077 Shanklin J, Cahoon EB (1998) Desaturation and related modifications of fatty acids. Ann Rev Plant Physiol Plant Mol Biol 49:611–641 Shima S, Thauer RK (2005) Methyl-coenzyme M reductase and the anaerobic oxidation of methane in methanotrophic Archaea. Curr Opin Microbiol 8:643–648 Smith BD, Sanders JL, Porubsky PR, Lushington GH, Stout CD, Scott EE (2007) Structure of the human lung cytochrome P450 2A13. J Biol Chem 282:17306–17313 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 Spormann AM, Widdel F (2001) Metabolism of alkylbenzenes, alkanes and other hydrocarbons in anaerobic bacteria. Biodegradation 11:85–105 Szaleniec M, Hagel C, Menke M, Nowak P, Witko M, Heider J (2007) Kinetics and mechanism of oxygen-independent hydrocarbon hydroxylation by ethylbenzene dehydrogenase. Biochemistry 46:7637–7646 Taupp M, Constan L, Hallam S (2010) The Biochemistry of Anaerobic Methane Oxidation. In: Timmis KN (ed) Handbook of Hydrocarbon and Lipid Microbiology. Springer, Berlin, Heidelberg Thauer RK, Shima S (2008) Methane as a fuel for anaerobic microorganisms. Ann N Y Acad Sci 1125:158–170 Throne-Holst M, Wentzel A, Ellingson TE, Kotlar H-K, Zotchev SB (2007) Identification of novel genes involved in long-chain n-alkane degradation by Acitnetobacter sp. strain DSM 17874. Appl Environ Microbiol 73:3327–3332 Ullrich R, Hofrichter M (2007) Enyzmatic hydroxylation of aromatic compounds. Cell Mol Life Sci 64:271–293 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 Urlacher VB, Schmid RD (2006) Recent advances in oxygenase-catalyzed biotransformations. Curr Opin Chem Biol 10:156–161 van Beilen JB, Funhoff EG (2005) Expanding the alkane oxygenase toolbox: new enzymes and applications. Curr Opin Biotechnol 16:308–314 van Beilen JB, Funhoff EG (2007) Alkane hydroxylases involved in microbial alkane degradation. Appl Microbiol Biotechnol 74:13–21 van Beilen JB, Neuenschwander M, Smits THM, Roth C, Balada SB, Witholt B (2002) Rubredoxins involved in alkane oxidation. J Bacteriol 184:1722–1732 van Beilen JB, Funhoff EG, van Loon A, Just A, Kaysser L, Bouza M, Holtackers R, Röthlisberger 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 Berkel WJH, Kamerbeek NM, Fraaije MW (2006) Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J Biotechnol 124:670–689 Whittington DA, Lippard SJ (2001) Crystal structures of the soluble methane monooxygenase hydroxylase from Methylococcus capsulatus (Bath) demonstrating geometrical variability at the dinuclear iron active site. J Am Chem Soc 123:827–38 Widdel F, Rabus R (2001) Anaerobic biodegradation of saturated and aromatic hydrocarbons. Curr Opin Biotechnol 12:259–276

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Widdel F, Musat F, Knittel K, Galusko A (2007) Anaerobic degradation of hydrocarbons with sulphate as electron acceptor. In: Barton LL, Hamilton WA (eds) Sulphate-reducing bacteria. Cambridge University Press, Cambridge, pp 265–303 Wilkes H, Rabus R, Fischer T, Armstroff A, Behrend 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 Wolter M, Zadrazil F, Martens R, Bahadir M (1997) Degradation of eight highly condensed polycyclic aromatic hydrocarbons by Pleurotus sp. Florida in solid wheat straw substrate. Appl Microbiol Biotechnol 48:398–404 Yoshizawa K, Yumura T (2003) A non-radical mechanism for methane hydroxylation at the diiron active site of soluble methane monooxygenase. Chem Eur J 9:2347–2358 Zahn JA, Arciero DM, Hooper AB, DiSpirito AA (1996) Evidence for an iron center in the ammonia monooxygenase from Nitrosomonas europaea. FEBS Lett 397:35–38

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization: An Introduction F. Widdel and F. Musat

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Some Basic Thermodynamic Aspects of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Energetics of Hydrocarbon Utilization by Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Catabolic Net Reactions of Hydrocarbons from the Energetic Perspective . . . . . . . . . . . 3.2 Hydrocarbon Activation from the Energetic Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Quantitative Aspects of Cell Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 ATP and Growth Yields . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Requirement for Minerals (N, P, Fe) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

34 34 40 40 45 50 50 57 59 60 71

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 ΔG 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.

F. Widdel (*) Max Planck Institute for Marine Microbiology, Bremen, Germany e-mail: [email protected] F. Musat UFZ – Helmholtz Centre for Environmental Research, Leipzig, Germany e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_2

33

34

1

F. Widdel and F. Musat

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 nonaromatic 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 compounds or elements (Appendix Table 5). In the case of gaseous hydrocarbons, a high volumetric energy density is 1

A fermentable hydrocarbon is, for instance, the unsaturated acetylene. Also some other unsaturated hydrocarbons are, at least theoretically, fermentable.

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

35

ol i s m

Cell synthesis

An ab

NH4+ HPO2– 4

Nonproductive

FeII, FeIII Hydrocarbon

Activated product

Energy

Energy ta Ca

Aox

bol ism

Aox

Oxidized form of electron acceptor

Ared Reduced form of electron acceptor

Ared

CO2

Energy dissipation

Fig. 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

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 ¼ 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 ΔG -values. If a reaction is feasible under the given conditions (exergonic reaction), the ΔG-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 to the surroundings (exothermic reactions),

2

H, 1.008; C, 12.011.

36

F. Widdel and F. Musat

which is expressed by their heat or enthalpy3 changes (ΔH-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 (ΔGf , sometimes also termed Gf ) or enthalpies of   formation (ΔHf , 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

(1)

(with a, b, c, d being the stoichiometric factors), the standard free energy change (viz., for all compounds at standard conditions) is the difference

      ΔG ¼ c ΔGf C þ d ΔGf D
a ΔGf A þ b ΔGf B

(2)

Calculation of the free energy change ΔG for nonstandard 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, ΔGStandard, according to ΔG ¼ ΔGStandard þ R T ln

acC  adD aaA  abB

(3)

T in this equation must be the temperature for which the underlying ΔGStandard value has been given (viz., usually 298.15 K), and ΔG 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

3

Heat change of reaction under constant pressure. T ¼ 298:15 K ð25 CÞ; standard activity of solutes, a ¼ 1; standard (partial) pressure of gases ¼ 101 kPa (standard fugacity ¼ 1). 5 ΔGStandard values at temperatures other than can be calculated via the integrated “Delta @ 298.15  KΔH ΔG version” of the Gibbs-Helmholtz equation @T T p ¼ T 2 : Assuming that temperature dependence of ΔH 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 4

ΔGStandard ¼ T

T  ΔG 298:15

 þ

1


 T  ΔH 298:15

The same result is obtained from ΔG ¼ ΔH
TΔS S by assuming that ΔH and ΔS are essentially constant within the range of physiologically relevant temperatures.

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

37

their pressures in atm, an otherwise obsolete unit.6 With such simplification, as well as with R ¼ 8:315  10
3 kJ K
1 mol
1, T ¼ 298:15 K ð25 CÞ, the common use of kJ as energy unit, and ln x ¼ 2:303 lg x, (3) converts to 

ΔG ¼ ΔG þ 5:71 lg

½Cc ½Dd ½Aa ½Bb

ðat 298:15 KÞ

(4)

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 ¼ 1, or ½Hydrocarbon ¼ 1. For instance, if a gaseous hydrocarbon at standard pressure dissolves in water and reaches the dissolution equilibrium (ΔG of transfer ¼ 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. If inorganic (fully oxidized) Carbon is involved, also acid-base dissociation has to be Considered (Appendix Fig. 7). The free energy data (Appendix Table 6) reveal some basic and sometimes “counterintuitive” thermodynamic properties of hydrocarbons. Many hydrocarbons  are metastable (thermodynamically unstable; ΔGf 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  ΔG ¼
43:3 kJðmol C2 H6 Þ
1

(5)

In the presence of CO2 or bicarbonate, even methane is metastable:

6

CH4 þ CO2 ! 2 CGraphite þ 2 H2 O  ΔG ¼
29:2 kJðmol CH4 Þ
1

(6)

CH4 þ CO2 ! 2 CGraphite þ 2 H2 O  ΔG ¼
29:2 kJðmol CH4 Þ
1

(7)

The apparent correctness of the old unit atm is due to the fact that it is numerically equivalent with standard fugacity ¼ 1. Activities and fugacities are by definition without units, and the formally ½A correct approximated substitution would aA ¼ ½A Actual , etc. Here, the use of the modern unit Pa or Standard kPa for [A], etc. is coherent.

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Nevertheless, formation of elemental carbon by reactions (5, 6, and 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 CO3 2 , are the thermodynamically stable forms of carbon. Other forms or intermediate oxidation states (H2 C ¼ 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 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 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  ΔG ¼
49:8 kJðmol C16 H34 Þ
1  ΔH ¼ þ454:4 kJðmol C16 H34 Þ
1

(8)

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 reveals the transition from gaseous to liquid  hydrocarbons (n-butane/n-pentane), which is mirrored by a discontinuity of the ΔHf

7

The extremely low hypothetical equilibrium concentrations of these species can be calculated. Linearity in the series of the higher alkanes may be a “pre-assumption” and basis for calculation of   ΔGf or ΔHf 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

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

39

0.4

Co2 (g) 0.2 HCO3– 0.0 CO32–

C

E –0.2 [V]

H

–0.4

H

2O

2

CH4 (g)

–0.6

–0.8 0

2

4

6

10

8

12

14

pH

Fig. 2 Stability diagram (Pourbaix diagram) of carbon species. The equilibrium (borderline) redox 

potential E (in V) as a function of pH was calculated from ΔGf values (in J) according to E ¼ X X ΔGof, red ΔGof, ox
∏aox þ 0:0592 lg with n ¼ number of electrons; F ¼ 96, 485 C n F n ∏ared
1 þ mol ; a ¼ activity . Πaox includes the H -activity, the negative logarithm of which is the pH. Activities or fugacities: CO2 , CH4 , 1:0; HCO3
and CO3 2
, 10
2 (black) or 1.0 (gray). The borderlines between CO2 ðgÞ, HCO3
and CO3 2
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 2Hþ =H2 ) is indicated for comparison

values. This is because liquid pentane has “given off” the heat of condensation to the  surroundings (liquid n-pentane, the real standard state: ΔH f ¼
173 kJ mol
1 ;  hypothetical gaseous standard state: ΔHf ¼
146 kJ mol
1 ). The discontinuity  of ΔGf 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:   ΔGf ¼
9:21 kJ mol
1; hypothetical gaseous standard state: ΔGf ¼
8:11 kJ mol
1).

40

F. Widdel and F. Musat

Fig. 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

+100

2

4

6

8

C-Atoms 10 12

14

16

18

ΔGf°

0

ΔGf° or ΔHf° [kJ mol–1]

–100 Gaesous

Liquid

Solid

–200 ΔHf°

–300

–400

–500

–600

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 Δ 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 ΔG. 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.

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

41

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 Cc Hh þ ð4c þ hÞ O2 ! 4c CO2 þ 2h H2 O 10 Cc Hh þ ð8c þ 2hÞ NO3
þ ð8c þ 2hÞ Hþ ! 10c CO2 þ ð4c þ hÞ N2 þ ð4c þ 6hÞ H2 O Cc Hh þ ð4c þ hÞ FeðOHÞ3 þ ð3c þ hÞ CO2 ! ð4c þ hÞ FeCO3 þ ð6c þ 2hÞ H2 Oa


8 Cc Hh þ ð4c þ hÞ SO4 2 þ ð8c þ 2hÞ Hþ ! 8c CO2 þ ð4c þ hÞ H2 S þ 4h H2 O 8 Cc Hh þ ð8c
2hÞ H2 O ! ð4c
hÞ CO2 þ ð4c þ hÞ CH4 Anabolism 4 Cc Hh þ h CO2 þ ð4c
hÞ H2 O ! ð4c þ hÞhCH2 Oi dass ¼ 0:133=ð4c þ hÞ mol g
1 Y ass ¼ ð4c þ hÞ=0:133 g mol
1 17 Cc Hh þ ð4h
cÞ CO2 þ ð14c
5hÞ H2 O ! ð4c þ hÞ C4 H7 O3 dass ¼ 0:165=ð4c þ hÞ mol g
1 Y ass ¼ ð4c þ hÞ=0:165 g mol
1 17 Cc Hh þ ð4h
cÞ CO2 þ ð4c þ hÞ NH3 þ ð10c
6hÞ H2 O ! ð4c þ hÞ C4 H8 O2 N dass ¼ 0:166=ð4c þ hÞ mol g
1 Y ass ¼ ð4c þ hÞ=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

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
þ 4 Hþ ! 2 N2 þ 3 O2 þ 2 H2 O, ΔG0 ¼
116 kJðmol NO2
Þ
1 ; 2 N2 O ! 2 N2 þ O2 , ΔG ¼
104 kJðmol N2 OÞ
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 ΔG ¼
20 kJ molðCH4 Þ
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 study of other syntrophic associations (Jackson and McInerney 2002; Schink 1997, 2002). Another anaerobic reaction of a hydrocarbon of thermodynamic interest is the conversion of alkanes


813.3d

2 C3 H8 ðgÞ þ 2 H2 O

2 C3 H8 ðgÞ þ 5 SO4

þ 10 H

þ

! 5 CH4 ðgÞ þ CO2 ðgÞ

! 6 CO2 ðgÞ þ 5 H2 SðaqÞ þ 8 H2 O

! 3 CO2 ðgÞ þ 4 H2 O

2


! 7 CH4 ðgÞ þ CO2 ðgÞ

Propane C3 H8 ðgÞ þ 5 O2 ðgÞ

þ 14 H

4 C2 H6 ðgÞ þ 2 H2 O

4 C2 H6 ðgÞ þ 7 SO4

2


! 8 CO2 ðgÞ þ 7 H2 SðaqÞ þ 12 H2 O


63


117


2,108


36


73


1,467

! 4 CO2 ðgÞ þ 6 H2 O

Ethane 2 C2 H6 ðgÞ þ 7 O2 ðgÞ

CH4 ðgÞ þ SO4

þH

þ

2


CH4 ðgÞ þ SO4

þ


16.6d,e

! HCO3
þ HS
þ H2 O


344d

CH4 ðgÞ þ SO4 2


þ2H

2


þ


16.5d,e

! 8 FeCO3 þ 14 H2 O

CH4 ðgÞ þ 8 FeðOHÞ3 þ 7 CO2 ðgÞ


928.3

! HCO3
þ H2 S ðaqÞ þ H2 O

! 3 CO2 ðgÞ þ 4 N2 ðgÞ þ 10 H2 O

3 CH4 ðgÞ þ 8 NO2
þ 8 Hþ
21.2d

! 5 CO2 ðgÞ þ 4 N2 ðgÞ þ 14 H2 O

5 CH4 ðgÞ þ 8 NO3
þ 8 Hþ

! CO2 ðgÞ þ H2 SðaqÞ þ 2 H2 O

! HCO3 þ H þ H2 O
766.2


818.0d




þ

! CO2 ðgÞ þ 2 H2 O

Productsa

CH4 ðgÞ þ 2 O2 ðgÞ

Reactantsa Methane CH4 ðgÞ þ 2 O2 ðgÞ

Free energy change of reaction per mol hydrocarbonb, ΔG or ΔG0 (kJ mol
1 )

6


46


2,220


2


38


1,560


11.3


33.5


20.9


559


993


788


903.0


890.4

Enthalpy change of reaction per mol hydrocarbon, ΔH  (kJ mol
1 )

232

236


375

115

117


310

18


57

1


723


215


73


301


243

Entropy change of reaction per mol hydrocarbonb,c, ΔS or ΔS0 (J K
1 mol
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

42 F. Widdel and F. Musat

! 30 FeCO3 þ 48 H2 O

! 24 CO2 ðgÞ þ 15 H2 SðaqÞ þ 12 H2 O

! 15 CH4 ðgÞ þ 9 CO2 ðgÞ

! 7 CO2 ðgÞ þ 4 H2 O

! 35 CO2 ðgÞ þ 18 N2 ðgÞ þ 38 H2 O

! 36 FeCO3 þ 58 H2 O

! 14 CO2 ðgÞ þ 9 H2 S ðaqÞ þ 8 H2 O

! 9 CH4 ðgÞ þ 5 CO2 ðgÞ

4 C6 H6 ðlÞ þ 18 H2 O

Toluene C7 H8 ðlÞ þ 9 O2 ðgÞ

5 C7 H8 ðlÞ þ 36 NO3
þ 36 Hþ

C7 H8 ðlÞ þ 36 FeðOHÞ3 þ 29 CO2 ðgÞ

2 C7 H8 ðlÞ þ 9 SO4 2
þ 18 Hþ

2 C7 H8 ðlÞ þ 10 H2 O

! 30 CO2 ðgÞ þ 15 N2 ðgÞ þ 30 H2 O

5 C6 H6 ðlÞ þ 30 NO3
þ 30 Hþ

C6 H6 ðlÞ þ 30 FeðOHÞ3 þ 24 CO2 ðgÞ

! 12 CO2 ðgÞ þ 6 H2 O

Benzene 2 C6 H6 ðlÞ þ 15 O2 ðgÞ

4 C6 H6 ðlÞ þ 15 SO4 2
þ 30 Hþ

! 49 CH4 ðgÞ þ 15 CO2 ðgÞ

! 64 CO2 ðgÞ þ 49 H2 S ðaqÞ þ 68 H2 O

4 C16 H34 ðlÞ þ 30 H2 O

4 C16 H34 ðlÞ þ 49 SO4

2


þ 98 H

! 80 CO2 ðgÞ þ 49 N2 ðgÞ þ 134 H2 O

þ

! 32 CO2 ðgÞ þ 34 H2 O

5 C16 H34 ðlÞ þ 98 NO3
þ 98 Hþ


142


238


1,689


3,590


3,823


135


214


1,423


3,008


3,202


372


632


9,757


10,392


137

4 C6 H14 ðlÞ þ 19 SO4

! 19 CH4 ðgÞ þ 5 CO2 ðgÞ


4,023

4 C6 H14 ðlÞ þ 10 H2 O

! 12 CO2 ðgÞ þ 14 H2 O

n -Hexadecane 2 C16 H34 ðlÞ þ 49 O2 ðgÞ

þ 38 H

þ


238

2


! 24 CO2 ðgÞ þ 19 H2 SðaqÞ þ 28 H2 O

n -Hexane 2 C6 H14 ðlÞ þ 19 O2 ðgÞ

96

2


2,421


3,449


3,910

71


7


2,026


2,883


3,268

206


50


9,445


10,701

66


33


4,163

801

806


2,456

473


293

691

696


2,024

419


220

1,937

1,952

1,047


1,036

682

688


471

(continued)

2 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 43

! 50 CO2 ðgÞ þ 24 N2 ðgÞ þ 44 H2 O

! 48 FeCO3 þ 76 H2 O

! 10 CO2 ðgÞ þ 6 H2 S ðaqÞ þ 4 H2 O

! 6 CH4 ðgÞ þ 4 CO2 ðgÞ

C10 H8 ðcÞ þ 48 FeðOHÞ3 þ 38 CO2 ðgÞ

C10 H8 ðcÞ þ 6 SO4 2
þ 12 Hþ

C10 H8 ðcÞ þ 8 H2 O


186


313


2,247


4,782


5,093

Free energy change of reaction per mol hydrocarbonb, ΔG or ΔG0 (kJ mol
1 )

186

61


3,171


4,541


5,156

Enthalpy change of reaction per mol hydrocarbon, ΔH  (kJ mol
1 )

1,245

1,253


3,098

808


212

Entropy change of reaction per mol hydrocarbonb,c, ΔS or ΔS0 (J K
1 mol
1 )

b

Indicated standard states: g, gaseous; l, liquid; aq, aqueous, dissolved in water; c, crystalline If protons are involved, ΔG 0 (viz., for ½Hþ  ¼ 10
7 M, pH ¼ 7) is given    c Here calculated via ΔS ¼ ΔH
ΔG T d Standard free energy changes of reactions formulated with CO2 differ from those formulated with HCO3
because the reaction HCO3
þ Hþ ! CO2 ðgÞ þH2 O is exergonic under standard conditions at pH ¼ 7, with ΔG0 ¼
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

! 10 CO2 ðgÞ þ 4 H2 O

5 C10 H8 ðcÞ þ 48 NO3
þ 48 Hþ

Productsa

Reactantsa Naphthalene C10 H8 ðcÞ þ 12 O2 ðgÞ

Table 2 (continued)

44 F. Widdel and F. Musat

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

45

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  ΔG ¼
372 kJðmol C16 H34 Þ
1  ΔH ¼ þ206 kJðmol C16 H34 Þ
1

(9)

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 ðC16 H34 þ 16 H2 O ! 8CH3 COO
þ 8 Hþ þ 17 H2 Þ, (2) acetate-cleaving microorganisms which are either methanogens ðCH3 COO
þ Hþ ! CH4 þ CO2 Þ or additional syntrophs ðCH3 COO
þ Hþ þ 2 H2 O ! 2 CO2 þ 4 H2 Þ, and (3) H2-utilizing methanogens ðCO2 þ 4 H2 ! CH4 þ 2 H2 OÞ. 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).

3.2

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 ΔG of the reaction (and its share in the overall catabolic ΔG), 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 cðCyt cox =Cyt cred , E ¼ E0 ¼ þ0:245 VÞ would be thermodynamically allowed: CH4 þ 2 Cyt c½Fe3þ  þ H2 O ! CH3 OH þ 2 Cyt c½Fe2þ  þ 2 Hþ ΔG 0 ¼
16 kJðmol CH4 Þ
1 

(10)

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

46

F. Widdel and F. Musat ΔG of activation

Without catalyst S

Enzymatic, simple model

Enzymatic, elementary reactions

Net ΔG of activation reaction

1 2

ΔG

ΔG of catabolic reaction

3

4 5

7

8

9 P Proceeding reactions

Fig. 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

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 ¼ þ 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  ΔG 0 ¼
344 kJ mol-1

(11)

The sacrifice of energy to achieve activation via oxygenases is also reflected by the consumption of reducing equivalents detained from the energy-conserving 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.

10


35 to
39
31 to
35

R
CH2
CH3 þ
OOC
CH ¼ CH
COO
!
OOC
½ðRÞ
CH
ðCH3 ÞCH
HCH
COO


C6 H5 CH 3 þ
OOC
CH ¼ CH
COO
!
OOC
½C6 H5 CH 2 CH
HCH
COO


n-Alkane

Toluene


27 to
31

Addition to fumarate CH4 þ
OOC
CH ¼ CH
COO
!
OOC
½CH3 CH
HCH
COO
Methanee


335

C6 H6 þ O2 ! ½Intermediate; NADH
recycling ! o-C6 H4 ðOHÞ2

þ30


368

C15 H31
CH3 þ O2 þ NADH þ Hþ ! C15 H31
CH2 OH þ H2 O þ NADþ


344c

Free energy b  or change  ΔG
1 0 ΔG kJ mol

Methyl-coenzyme M reductase Methane CH4 þ CoM
S
S
CoB ! CoM
S
CH3 þ HS
CoB

nHexadecaned Benzene

Type of activation; compound Reactiona Mono- and dioxygenation Methane CH4 þ O2 þ NADH þ Hþ ! CH3 OH þ H2 O þ NADþ

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . (continued)

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

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 ¼ 7, if protons are involved)

2 47


31

Widdel et al. (2007) Widdel et al. (2007)

Widdel et al. (2007)

Reference

b

Fate of one reactant is visualized in bold ΔG 0 is indicated if protons are involved c Would be less exergonic with an electron donor of less negative redox potential than that of NADþ =NADH ð
0:320 VÞ d Sum of the following formal reactions: C15 H31
CH3 þ 0:5O2 ! C15 H31
CH2 OH , ΔG ¼
148:5 kJ mol
1 ; 0:5O2 þ NADH þ Hþ ! H2 O þ NADþ , ΔG0 ¼
219:6 kJ mol
1 (calculated from ΔE0 ¼ 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 have not been suggested or identified. The present free energy change is based on a calculation with oxaloacetate as a purely hypothetical carboxyl donor that may be energetically comparable to potent carboxyl donors such as carboxy-biotin

a

C6 H6 þ Carrier
COO
! C6 H5
COO
þ Carrier
H


4

Carboxylationf Benzene CH3
CH2
CH ¼ CH2 þ H2 O ! CH3
CH2
HCH
CH2 OH

Benzeneg


7

Free energy b  or change  ΔG
1 0 ΔG kJ mol

Type of activation; compound Reactiona Addition of water to isolated double bond CH3
CH2
CH ¼ CH2 þ H2 O ! CH3
CH2
HCH
CH2 OH Butenee

Table 3 (continued)

48 F. Widdel and F. Musat

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

49

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. Despite 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  ΔG ¼
818 kJðmol CH4 Þ
1

(12)

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 are 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 þ HS
CoB ! CoM
S
S
CoB þ CH4 , ΔG ¼
30 ½10 kJ mol
1 ; Shima and Thauer 2005; Thauer and Shima 2008). For methane activation, the standard free energy of the reverse Mcr (rMcr) reaction in methane oxidizing archaea would thus be þ30 ½10 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.

11

The utilization of chlorate by facultatively anaerobic bacteria for hydrocarbon metabolism (Chakraborty and Coates 2004; Tan et al. 2006) involves O2 that is generated from an intermediate ðClO2
! Cl
þ O2 Þ.

50

F. Widdel and F. Musat

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 (Krüger 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 non-methane 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 C6 H5
▪ 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 (Boll et al. 2018). 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.

4

Quantitative Aspects of Cell Synthesis

4.1

ATP and Growth Yields

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,

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 ΔG ¼ 0. In case of the reaction S ! P, the connection is

½P ½S eq

kS =K S



m ¼ kcat ¼ e
ΔG P =K P cat

m

=ðRT Þ

:

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

51

besides calculated free energy changes and ATP yields known from well-established 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 dissimilated (viz., the energy yielding) proportion of the substrate, Sdiss, would be a more meaningful definition: Y¼

 X  g mol-1 Stot

Y diss ¼

X  Sdiss

g mol-1



(13a; b)

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 SO4 2 ) 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 > (14) 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

(15)

Division by the obtained cell mass yields Stot Sdiss Sass ¼ þ X X X

(16)

1 1 Sass ¼ þ Y Y diss X

(17)

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 d ass ¼ 1=Y ass . This leads to

Toluene, O2

Pseudomonas putida 1.0*

92.1

100

1.28*

Toluene, O2

Pseudomonas putida

148 197

120*,g 94.7

0.50 1.20*

n-Heptadecane, O2 Benzene, O2

Pseudomonas nautica Pseudomonas putida

310 366 1240

204 226 401

Micrococcus cereficans Candida tropicalis Ideal aerobe

127f to 280f

0.60*,e to 1.32* 0.9* 1.0* 1.77

n-Octane, O2

159

185

164f to 563f

214

125

1.1*

Hydrocarbon and electron acceptor

Long-chain n-alkane mixture, O2 n-Hexadecane, O2 n-Hexadecane, O2 n-Hexadecane, O2

Nocardia sp.

Microorganism Aerobic Pseudomonas sp.

Growth yieldb (cell dry mass per amount of hydrocarbon) c molar, by mass   molar  , 
1
1 ðg g Þ Y g mol Y diss g mol
1

0.58 (58%)

0.54 (54%)

0.81 (81%) 0.48 (48%)

0.66 (66%) 0.62 (62%) 0.32 (32%)

0.77 (77%) to 0.50 (50%)

0.58 (58%)

Fraction of substrate catabolizedd, Sdiss/Stot

Einsele (1983) Einsele (1983) This article (see text) Bonin et al. (1992) Reardon et al. (2000) Reardon et al. (2000) Bordel et al. (2007)

Wodzinski and Johnson (1968) Wagner et al. (1969)

Reference

Table 4 A selection of growth yieldsa of aerobic and anaerobic microorganisms on hydrocarbons and calculated fraction of the dissimilated substrate

52 F. Widdel and F. Musat

0.83 0.037 0.059 0.31

Methane, SO4 2


n-Hexadecane, SO4 2


Toluene, SO4 2


0.70* 0.50*

Ethylbenzene, NO3


Toluene, O2 Naphthalene, O2

33*

13.8

13.5*,h 29*

0.6*

129*

91.6 82

0.59

88*

64.5 64.1

0.88 (88%)

0.97 (97%)

0.99 (99%)

0.68 (68%)

0.70 (70%) 0.78 (78%)

Rabus and Widdel (1995) Nauhaus et al. (2007) So and Young (1999) Rabus et al. (1993)

Dinkla et al. (2001) Wodzinski and Johnson (1968)

Only directly measured (“real”) growth yields are listed and not Ymax values obtained from extrapolation b Original value from the reference is indicated by asterisk; other values were calculated for this chapter (see text). Ydiss was 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; npentadecane, 558; n-hexadecane, 594; n-heptadecane, 630; benzene, 182; toluene, 218; ethylbenzene, 255 c Relative to dissimilated substrate d Calculated according to Sdiss =Stot ¼ Y=Y diss 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

a

Anaerobic Betaproteobacterium, strain EbN1 Archaea (ANME-2) Deltaproteobacteria Deltaproteobacterium, strain AK-01 Desulfobacula toluolica

Pseudomonas putida Pseudomonas sp.

2 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 53

54

F. Widdel and F. Musat

1 1 1 ¼ þ Y Y diss Y ass

(18)

Y ass  Y Y ass
Y

(19)

Rearrangement leads to Y diss 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 Y ass
Y ¼ Y ass Stot

(20)

For instance, for aerobic growth with hexadecane (M ¼ 226:45), a growth yield by mass of 1 g ðg C16 H34 Þ
1 has been reported, which equals a molar growth yield of Y ¼ 226 g ðmol C16 H34 Þ
1 . The assimilation equation is 17 C16 H34 þ 120 CO2 þ 54 H2 O ! 98 C4 H7 O3 dass ¼ 1:68  10
3 mol g
1 ; Y ass ¼ 594 g mol
1

(21)

Equation (19) yields Y diss ¼ 366 g mol
1. The fraction of the dissimilated substrate is Sdiss ¼ 0:62 ðor 62%Þ Stot

(22)

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

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

55

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 straightforward. From anaerobic pathways with biochemically known qATP, as for instance the homolactic fermentation of glucose ðqATP ¼ 2Þ, the amount of cell mass obtained per mol ATP, the so-called YATP, can be calculated from the determined growth yield via Y ATP ¼ Y diss =qATP. If for another bacterium of interest, the qATP is unknown but Ydiss has been determined, this should in principle allow to calculate the desired qATP parameter via qATP ¼ Y diss =Y ATP .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 nonproductive consumption of energy or ATP is interpreted as maintenance energy (Pirt 1965; Tempest and Neijssel 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, μ. 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/μ, the proportion of the substrate consumed for maintenance should become zero. At 1=μ ¼ 0 ðμ ¼ 1Þ the theoretically highest growth yield, Ymax (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 longchain 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 (β-oxidation, citric acid cycle, dehydrogenations with NADþ 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 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

56

F. Widdel and F. Musat O2 H2O

ATP

C15H31–CH2OH

C15H31–CH3

C15H31–CHO

NAD–H

NAD–H

7 (FADH2) + 7 NAD–H

15 OH2 + 32 NAD–H 30 e–

C15H31–CO–SCoA

NAD–H

Consumption not depicted 31 H2O

C15H31–COOH

Reducing equivalents

β–Oxidation

8 CH3–CO–SCoA

64 e–

47 H2O

8 (FADH2) + 24 NAD–H

Respiratory chain

TCA

16 CO2

23.5 O2 ≈ 26 ATP

≈ 91 ATP

8 ATP

≈ 124 ATP Catabolism: C16H34 + 24.5 O2 → 16 CO2 + 17 H2O ΔG° = –10 393 kJ (mol C16H34)

–1

Fig. 5 Reducing equivalents and ATP synthesis in the aerobic catabolism of hexadecane ðC16 H32 , M ¼ 226:45Þ. 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 Hþ =NADH and 6 Hþ =QH2 . A phosphorylation yield of 1 ATP per 3:5 Hþ was arbitrarily assumed here (based on the commonly assumed range of 1 ATP per 3 to 4 Hþ ). 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 (C6 H12 O6 , M ¼ 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

studies, a Ymax 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 ¼ 124 ðmol=molÞ. 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 ðmol ATPÞ
1. If a YATP of 10 g cell dry mass ðmol ATPÞ
1 is assumed that is likely for cell synthesis from the hexadecane-derived acetate units (Erickson 1981; Stouthamer 1988), this would lead to Y diss ¼ 1240 g cell mass ðmol C16 H34 Þ
1 . The Y value is obtained via a transformation of (19): Y¼

Y ass  Y diss Y ass þ Y diss

(23)

This yields (with the above Y ass ¼ 594 g mol
1 ) a value of Y ¼ 401 g mol
1 , which would be a yield by mass of 1.77 g cell mass ðg C16 H34 Þ
1 . This may be regarded as an “ideal” yield with hexadecane. The fraction of dissimilated hexadecane would be only

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

CH4 (g) a = 719 (ΔGf = –34.4)

–16.3 kJ +16.3 kJ

57

CH4 (g) a = 1.00 ΔGf ° = –50.75 kJ

Gas phase 0 kJ

0 kJ

0 kJ

0 kJ

Water phase

CH4 (g) a = 1.00 ΔGf ° = –34.4 kJ

–16.3 kJ +16.3 kJ

CH4 (aq) a = 0.00139 (ΔGf = –50.75)

All free energie values per mol

Fig. 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 ΔG oder ΔG 0 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 ΔGf 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

Sdiss ¼ 0:32 ðor 32%Þ Stot

(24)

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). In the environment and in cultures, microorganisms often obtain the limiting elements

0 kJ

–

H

+

0 kJ

H2O

+ 3.62 kJ

– 3.62 kJ

H

+

H2O – 3.62 kJ + + 3.62 kJ H

H2O

+ 3.62 kJ

–

H

+

0 kJ

H2O

–7

T

+ 4.72 kJ

– 4.72 kJ

H

+

H2O – 4.72 kJ + + 4.72 kJ H

H2O

–

H

+

0 kJ

H2O

HCO3 (aq) a = 1.249 (ΔGf = –591.57 kJ)

H

+

0 kJ

H2O

H2O (I), a = 1.00 , ΔGf° = –237.18 kJ T

0 kJ

CO2 (aq) a = 0.0346 (ΔGf = –394.36)

0 kJ

CO2 (g) a = 1.00 ΔG°f = –394.36 kJ

H (aq), a = 10 , ΔGf° = –39.97 kJ

+

HCO3 (aq) a = 1.00 ΔG °f = –586.85 kJ

H

+

0 kJ

H2O

+ 4.72 kJ

– 4.72 kJ

CO2 (aq) a = 0.232 (ΔGf = –389.64)

Gas phase

– 3.62 kJ

0 kJ

+ 4.72 kJ

– 4.72 kJ

Water phase pH = 7.0

0 kJ

CO2 (g) a = 6.7 (ΔG°f = –389.64 kJ)

Water phase pH = 7.0

Gas phase

+ 3.62 kJ

– 3.62 kJ

H2O +

+

H

CO2 (aq)

–

+

H

H2O

HCO3 (aq) a < 0.149

+

H

H2O

a < 0.0346

All free energie values per mol

H

H2O

Water phase pH = 7.0

Gas phase

Co2 (g) a < 1.00

Fig. 7 The three standard states (framed) of inorganic carbon (CO3 2
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 ¼ 7

HCO3 (aq) a = 4.31 (ΔGf = –583.23 kJ)

H

+

0 kJ

H2O

CO2 (aq) a = 1.00 ΔG°f = –386.02 kJ

0 kJ

CO2 (g) a = 28.9 (ΔGf = –386.02)

Natural systems

58 F. Widdel and F. Musat

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

59




as inorganic species (NH4 þ ,NO3
, H2 PO4
=HPO4 2 , Fe2þ , 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, ðCH2 OÞ106 ðNH3 Þ16 ðH3 PO4 Þ ¼ hC106 H263 O110 N16 Pi, 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 con  sumption of 1 g (1.3 ml) hexadecane would need 0.24 g 4:5  10
3 mol NH4 Cl and   0:04 g 0:3  10
3 mol KH2 PO4 : In a marine environment with for instance 1 μM combined nitrogen and 0.06 μM 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 (Harms et al. 2017). 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 (Ron and Rosenberg 2010).

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

60

F. Widdel and F. Musat

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.

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 ΔG of oxidation with O2 Per mass of Per volume of substance substance   ðkJ m
3 Þ kJ kg
1

Substance Gases (101 kPa) H2
117:6  103

a


9:7  103

ΔH of oxidation with O2 Per mass of substance  kJ kg
1
141:8  103


11:7  103

CH4


51:0  10


33:5  10


55:5  10


36:4  103

C3H8


47:8  103


86:2  103 a


50:3  103


90:8  103 a

NH3b


19:9  10


13:8  10

3


22:5  10


15:6  103 a

3

3

3

3a

3


19:3  103 H2Sc Solids or liquids Li
40:4  103


26:8  103 a


23:3  103


32:4  103 a


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


74:4  10

3


32:8  10


74:4  106

C6H14


46:7  103


30:8  106


48:3  103


31:9  106

C6H6


41:0  10


36:0  10

3


41:8  10


37:5  106

CH3OH


21:9  103


17:5  106


22:7  103


18:1  106

CH3CH2OH


28:8  10


22:8  10

3


29:7  10


23:5  106

C6H12O6(α-DGlucose) Mg


16:0  103


25:0  106


15:6  103


24:3  106


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


39:7  10

3


24:1  10


43:9  106

S


11:6  103


22:7  106


12:3  103


24:1  106

Fe


6:6  10


52:3  10


7:4  10


58:3  106

3

3

3

3

3

3

6

6

6

6

6

6

3

For convenience, ideal behavior assumed. In reality, the volumetric energy density will be somewhat higher b If N2(g) is produced c If H2SO4(l) (l) is produced

Formula mass   g mol
1 16.043

16.043 30.069

44.096

58.123

58.123 72.150 72.150 86.177

86.177 100.203 114.23

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Alkanes Methane (g)

Methane (aq) Ethane (g)

Propane (g)

n-Butane (g)

2-Methylpropane (g)

n-Pentane (l)

2-Methylbutane (l) n-Hexane (l)

2-Methylpentane (l) n-Heptane

n-Octane (l)


8.11e 1.0c 1.28e 6.41e


50.72c
50.8d
50.75f
34.4g
32.82c
32.6e
32.89f
23.49c
23.6d
23.4e
17.03c
17.2d
15.7e
20.9d
18.0e
9.3d
9.21e
14.6e
3.8d
4.28e

Free energy offormationfrom the  elements, ΔGf kJ mol
1

310.23c 310.1d


126.15c
125.6d
125e
134.2d
132e
173.1c
173.5d
179e
198.7c
198.8d
199e
204e
224.4c

(continued)

291e 328.6c 328e 361.1c

260e 296.1d 296e

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .


249.9c

269.91c 270.2d


103.85c
104.7d

294.6d 295e 262.7d

229.60c 229.1d

186.26c 186.2e

b, Entropy   S J K
1 mol
1


84.68c
83.8d


74.81c
74.9d

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Table 6 Thermodynamic properties of hydrocarbons and other compounds. Data are from other compilationsa

2 61

Formula mass   g mol
1 114.23 114.23 114.23 142.28 170.34 184.36 198.39 226.44 240.47 254.50 70.134 84.161

28.054 42.080

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

2-Methylheptane (l) 3-Methylheptane (l) 4-Methylheptane (l) n-Decane (l)

n-Dodecane (l)

n-Tridecane (l) n-Tetradecane (l) n-Hexadecane (l)

n-Heptadecane (l) n-Octadecane (c) Cyclopentane (l)

Cyclohexane (l)

Unsaturated hydrocarbons, nonaromatic Ethene (g)

Propene (g)

Table 6 (continued)

68.15c 68.4d 62.78c 62.8d 74.8e

53.9 36.4d 36.5e 26.8e 26.7d

e

52.26c 52.5d 20.42c 20.0d 20.4e


480e
569e
105.1d
106e
156c
156.4d

219.56c 219.3d 267.05c 266.6d 227e

204.4d

497e 204.3d

523e 555e

352e 358e 350e 425.5d 426e 490.6d


255.1c
252c
252c
300.9d

3.85e 4.68e 7.8e 17.5d 17.4e 28.1d 28.4e 33.8e 38.8e 49.8h 52.2i
350.9d
352e
378e
403e
454.4h

b, Entropy   S J K
1 mol
1

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

62 F. Widdel and F. Musat

255e 246.5d 246e 252.2d 247.4d


12.5e
24.4e

78.113

92.140

106.17 106.17 106.17 106.17 120.19 128.17 142.20 142.20 154.21

Toluene (l)

Ethylbenzene (l) o-Xylene (l)

m-Xylene (l) p-Xylene (l)

1,3,5-Trimethylbenzene (l)

Naphthalene (c)

1-Methylnaphthalene (l) 2-Methylnaphthalene (c) Biphenyl (c)

(continued)

254.8d 220d 205.9d

273.6d 273e 166.9d

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

189.4d 192.6d 254.2d

201d

103.9d

221d 219e

12.4d 8.08e

26.038

Ethyne (g) Aromatic hydrocarbons Benzene (l)


25.4e
24.4d
24.3e
63.4d
63.5e 78.53c 77.9d 56.3d 44.9d 99.4d

173.3c 173.4d

49.0c

124.3c 124.4d 124.5f 113.8d 110e 114.22f 120e 110.3d 111e 107.7d 110.1d

56.107

trans-2-Butene (g)

296.59c 296.5d 296e 200.94c

56.107

cis-2-Butene (g)

305.71c 305.6d 307e 300.94c 300.8d


0.13c 0.1d 1.17e
6.99c
7.1d
5.70e
11.17c
11.4d
10.1e 226.73c

71.39c 71.3d 72.0e 65.95c 65.9d 67.3e 63.06c 63d 64.3e 209.2c

56.107

1-Butene (g)

2 63

Formula mass   g mol
1 178.23 178.23

32.042 32.042 46.069 46.069 60.096 60.096 60.096 60.096 74.122 74.122 74.122 242.44 108.14

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Anthracene (c)

Phenanthrene (c)

Alcohols, phenolic compounds Methanol (l)

Methanol (aq) Ethanol (l)

Ethanol (aq) 1-Propanol (l) 1-Propanol (aq) 2-Propanol (l)

2-Propanol (aq) 1-Butanol (l)

1-Butanol (aq) 2-Butanol (l) 1-Hexadecanol (c)

Benzyl alcohol (l)

Table 6 (continued)


171.84f
177.0d
98.7d
98.8e
27.5d
27.3e


166.27c
166.8d
175.39f
174.78c
174.2d
181.75f
170.6d
175.81f
180.3d
182e
185.94f
162.5d


342.6d
686.7d
684e
160.7d

225.1d 451.9d 452e 216.7d

226.4d 252e

180.6d 180e


318.1d
319e
327.3d

194.6d

160.7c 161.0d

126.8c 127.2d

207.6d 207e 211.7d 212e

b, Entropy   S J K
1 mol
1


302.6d


277.69c
277.0d


238.66c
239.1d

129.2d 128e 116.2d 113e

286.0d 268.3d

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

64 F. Widdel and F. Musat

94.113

124.14 30.026

30.026 44.053 44.053 58.080 58.080 72.107 106.12 46.026

46.026 45.018

Phenol (s)

1,2-Dihydroxybenzene (s) Aldehydes, ketones Formaldehyde (g)

Formaldehyde (aq) Acetaldehyde (l)

Acetaldehyde (aq) Butyraldehyde (l) Acetone (l)

Acetone (aq) 2-Butanone (l)

Benzaldehyde (l) Carboxylic acids, carboxylates Formic acid (l)

Formic acid (aq) Formate
ðaqÞ


356.3j
351.04f
334.9j


361.35c
360e


102.53c
109.9d
111e
112.97f
130.54f
128.12c
128.3d
139.9f
127e
155.4c
155.8d
161.17f
151.4d
156e 9.4d


50.9c
50.4d
47.5e
47.6f
210.0d


424.72c
425.1d
423e
410.3j
410.3j


273.3d
279e
87.0d


247e
248.1c
242.1d


192.30c
191.8d

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . (continued)

163.7j 91.7j

128.95c

238.8d 241e

247e 200.4c 200.6d

160.2c 160.4d

218.77c

150.2d


361.1d
108.57c
116e

146c 144d 142e


165c
165.1d
163e

2 65

60.052

60.052 59.045 73.071 87.098

Acetic acid (l)

Acetic acid (aq) Acetate
ðaqÞ

Propionate
ðaqÞ

Butyrate
ðaqÞ

116.07 130.10 200.23 206.20

Succinate2
ðaqÞ

Methylsuccinate2
ðaqÞ

ð1
MethylpentylÞ succinate2
ðaqÞ

Benzylsuccinate2
ðaqÞ


521.1 to
525.4l


644.0 to
647.3l


681.6 to
685.5l


690.23f


604.21f


305.0f
245.3c
245.6f
229.3k


335.96f


23.0d
28e

242.6d 242e

452e 167.6c

178.7c 86.6c


485.76c
486.01c


882e
385.1c
385.2d

159.8c 159.9d


484.5c
484.4d


389.9c
390.2d
392e
396.46c
369.31c
369.41f
361.08f
352.63f

b, Entropy   S J K
1 mol
1

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

Hydrocarbon-derived nitrogen, oxygen, sulfur, and halocompounds Methylamine (g) 31.057 32.3d 27.5e þ 32.065
40.0f Methylammonium ðaqÞ

121.12 114.06

Fumarate2
ðaqÞ

256.43 122.12

Palmitic acid (c) Benzoic acid (c)

Benzoate
ðaqÞ

115.15

Hexanoate ðaqÞ




Formula mass   g mol
1

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Table 6 (continued)

66 F. Widdel and F. Musat

45.084 79.101 79.101 93.128 46.069 74.122 48.10 62.13 62.13 84.14 110.17 34.033 88.005 50.488 96.944 119.38 153.82 64.515

Ethylamine (g)

Pyridine (l) Pyridine (aq) Aniline (l)

Dimethyl ether (g)

Diethyl ether (l) Methanethiol (g)

Dimethyl sulfide (l) Ethanethiol (l) Thiophene (l)

Thiophenol (l)

Fluoromethane (g)

Tetrafluoromethane (g)

Chloromethane (g)

Dichloromethane (l) Trichloromethane (l) Tetrachlorromethane (l)

Chloroethane (g)


213.8d
222e
888.3d
862e
58.5d
58.1e
63.3e
71.2e
62.6d
68.4e
60.5d
53.0e

181.3d 177.1f 149.2d 148e
112.9d
114e
116.7d
9.9d 0.754e 5.72e
5.7e 121.2d 122e 134d

37.3d

31.3d 29.7e
184.1d
185e
279.3d
22.9d
12.4e
65.4e
73.7e 80.6d 81.7e 64.1d 62.8e
237.8d
247e
933.6d
908e
81.9d
82.0e
117e
132e
132.8d
139e
112.1d
105e


47.4d
48.5e 100.2d

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . (continued)

261.3d 262e 234.2d 233e 179e 203e 216.2d 214e 275.8d 275e

222.8d

222.8d

196e 207e 181.2d

253.1d 255.1d

191.4d 192e 267.1d

177.9d

289.9d

2 67

Formula mass   g mol
1 96.104 112.56

2.0158 1.0074 1.0074 12.011 12.011 28.011 44.010 44.010 61.017 60.009 28.0134 18.038 44.013

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Fluorobenzene (l) Chlorobenzene (l)

Inorganic compounds H2(g) Hþ ðaqÞ, pH ¼ 0

Hþ ðaqÞ, pH ¼ 7

C, graphite (c) C, diamond (c)

CO (g)

CO2(g)

CO2(aq)

HCO3
ðaqÞ

CO3 2
ðaqÞ

N2(g) NH4 þ ðaqÞ

N2O (g)

Table 6 (continued)


39.97l


137.17c
137.15f
394.36c
394.39d
385.98c
386.02f
586.77c
586.85f
527.81c
527.90f 0
79.31c
79.37e 104.20c 104.18f

0 2.900c

0

ΔGf

191.61c 113.4c 219.85c

82.05c


56.9c


677.14c 0
132.51c

91.2c

213.74c 213.80d 117.6c


393.51c
393.52d
413.80c
691.99c

197.67c

5.740c 2.377c

(ΔS 0 ) 134.06

130.684c 0

0 1.895c 1.897d
110.53c

0

0 0

206e 209.2d 194e


145e 11.0d 10.6e


69.0e 89.2d 93.7e 0 0


b, Entropy   S J K
1 mol
1

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

68 F. Widdel and F. Musat

62.005

31.999 18.015

32.06 34.08 34.08 33.072 96.06 18.999

NO3
ðaqÞ

O2(g)

H2O (l)

S, (α, rhombic; c)

H2S (g)

H2S (aq)

HS
ðaqÞ

SO4 2
ðaqÞ

F
ðaqÞ 35.453 54.937 114.95 86.937 55.846

Cl ðaqÞ

Mn2þ ðaqÞ

MnCO3 MnO2 Fe2þ ðaqÞ




46.006

NO2
ðaqÞ


131.23
131.3m
227.8j
228.0m
816.0m
465.1m
78.90c
78.87m c


33.56c
33.3d
27.83c
27.87d 12.08c 12.05m
744.53c
744.63f
278.79c


285.83c


237.13c
237.14d
237.178f
237.18m 0


223.3j
220.7m
889.3m
520.0m
89.10m


167.16

c

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . (continued)


84j
73.6m 100m 53m
137.7c

56.5c


13.8c

20.1c


909.27c
332.63c

62.08c

31.80c 32.056d 205.79c 205.7d 121c

205.138c 205.147d 69.91c 69.95d

125.2j 140m 146.4c 146.5j


20.63c
20.5d
39.7c
39.8d
17.6c

0


106.3j
104.6m
205.0c
206.7j
207.3m 0


37.2f,m
34.54j
108.74c
111.34f
110.7j 0

2 69

55.845 115.86 106.87 159.69 231.54

Fe3þ ðaqÞ

FeCO3 (siderite; c)

Fe(OH)3 (amorphous)

Fe2O3 (α, hematite; c)

Fe3O4 (magnetite; c)


315.9c 92.9j 105m 96j 87.40c


48.5c
748.2j
737.0m
824.8j
824.2c
824.6m
1118.4c
1115.7m


4.7c
4.60m
674.3j
666.7m
695j
699m
742.2c
742.7m
1015.4c
1012.6m 146.4c

b Entropy 
1,
1   S J K mol

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

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, ΔSf ; the latter can be used to prove     consistency of literature data via ΔGf ¼ ΔH f
298:15ΔSf . Example of n-hexane (C6H14): ΔSf ¼ 291:6
ð6  5:74Þ
ð7  130:68Þ ¼
657:6 J K
1    mol
1 . ΔGf ¼
198:8
ð298:15 
0:6576Þ ¼
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 ¼ 4:1868 J) k Widdel et al. (2007) l Free energy associated with dilution of 1 mol Hþ from a ¼ 1 to a ¼ 10
7 , which is R T ln 10
7 m Stumm and Morgan (1981)

Formula mass   g mol
1

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Table 6 (continued)

70 F. Widdel and F. Musat

2

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

71

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3

Physiology and Biochemistry of the Aerobic Methanotrophs Valentina N. Khmelenina, J. Colin Murrell, Thomas J. Smith, and Yuri A. Trotsenko

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Methane Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 sMMO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 pMMO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Methanobactins as Vital Chalkophores . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Methanol Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Formaldehyde and Formate Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Carbon Assimilation Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 The Serine Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 The RuMP Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 CBB Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Nitrogen Assimilation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Assimilation of Methylated Amines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 TCA Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Osmoadaptation Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Methanotrophs and Biotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Prospects for the Future . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

74 78 78 79 79 81 82 83 84 85 87 88 89 89 90 91 92 92 93

V. N. Khmelenina · Y. A. Trotsenko (*) G.K. Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino/Moscow, Russia e-mail: [email protected]; [email protected] J. Colin Murrell School of Environmental Sciences, University of East Anglia, Norwich Research Park, Norwich, UK e-mail: [email protected] T. J. Smith Biomolecular Sciences Research Centre, Sheffield Hallam University, Sheffield, UK e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_4

73

74

V. N. Khmelenina et al.

Abstract

Methanotrophs are a widely distributed group of aerobic bacteria that use methane as their source of carbon and energy. They play key roles in the global carbon cycle, including controlling anthropogenic and natural emissions of the greenhouse gas methane. Methanotrophs oxidize methane using the unique enzyme methane monooxygenase which exists in two structurally and biochemically distinct forms. One form, the membrane-associated or particulate methane monooxygenase (pMMO), is found in most known methanotrophs and is located in the cytoplasmic membrane. Another form, the soluble methane monooxygenase (sMMO), is found in some methanotrophs and is located in the cytoplasm. Both forms of MMO can co-oxidize a range of hydrocarbons and chlorinated pollutants and hence are interesting with respect to the biotechnological potential of methanotrophs. Methanol is further oxidized to formaldehyde, formate, and CO2, by specific methylotrophic enzymes, while biomass is built from formaldehyde, formate, CO2, or a combination thereof via three cyclic biochemical pathways: the ribulose monophosphate (RuMP) cycle, the serine pathway, and the Calvin-Benson-Bassham (CBB) cycle. The availability of genome sequences of methanotrophs enables postgenomic studies to investigate the regulation of methane oxidation in the laboratory and in the environment by natural methanotrophs and in laboratory or industrial conditions by platform organisms. Recent studies have included synthetic biology approaches and in future may incorporate the design of new pathways.

1

Introduction

Methane-oxidizing bacteria (methanotrophs) are a widely distributed group of aerobic microorganisms that use methane as their source of carbon and energy. Methanotrophic bacteria belong to the phyla Proteobacteria (Hanson and Hanson 1996) and Verrucomicrobia. They include extremely acidophilic methanotrophs (Op den Camp et al. 2009) and the candidate division NC10. NC10 includes a candidate oxygenic methanotroph that oxidizes CH4 by using O2 produced in situ from nitrite (Ettwig et al. 2010). Methanotrophs play a major role in the environment, oxidizing methane that is released into the environment due to the activity of methanogenic archaea and industrial human activity, thereby mitigating the effects of this potent greenhouse gas (Hanson and Hanson 1996). Methanotrophs of the phylum Proteobacteria are classified as type I or type II methanotrophs according to whether they belong to the γ- or α-class of the Proteobacteria, respectively. Methanotrophs can be isolated from many ecosystems including soils, peatlands, rice paddies, sediments, freshwater and marine systems, alkaline soda lakes, acidic hot springs, mud pots, cold ecosystems, and tissues of higher organisms (McDonald et al. 2008). Within the Gammaproteobacteria, there are currently 16 genera of methanotrophs within the family Methylococcaceae and 3 genera in the family

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Methylothermaceae (Table 1). The genera within the family Methylococcaceae are Methylobacter, Methylocaldum, Methylococcus, Methylogaea, Methyloglobulus, Methylomagnum, Methylomarinum, Methylomicrobium, Methylomonas, Methyloparacoccus, Methyloprofundus, Methylosoma, Methylosphaera, Methylosarcina, Methyloterricola, and Methylovulum. Methylohalobius, Methylomarinovum, and Methylothermus are the methanotrophs in the Methylothermaceae family. All methanotroph species of this phylum are highly specialized, to the point that they grow only on methane and its one-carbon derivatives such as methanol and cannot grow on complex, multicarbon substrates such as sugars or organic acids. They assimilate carbon predominantly via the RuMP pathway. Also, filamentous gammaproteobacterial methanotrophs of the genera Crenothrix and Clonothrix have been described, but they have been observed only in enrichments (Oswald et al. 2017). The Alphaproteobacteria currently include two methanotroph genera in the family Methylocystaceae, Methylosinus, and Methylocystis and three genera in the family Beijerinckiaceae, Methylocella, Methylocapsa, and Methyloferula. Alphaproteobacterial methanotrophs assimilate C1 compounds via the serine pathway; they possess either the ethylmalonyl-coenzyme A pathway for glyoxylate regeneration or a glyoxylate bypass (Chistoserdova et al. 2009). Beijerinckiaceae representatives of methanotrophs additionally contain the complete set of genes for the function of the Calvin-Benson-Bassham (CBB) cycle. Members of the genus Methylocella and some members of the genera Methylocystis and Methylocapsa are facultative methanotrophs that can grow on a narrow range of multicarbon compounds (acetate and several other organic acids, ethanol, and some short-chain alkanes) in addition to methane (Dunfield and Dedysh 2014). The alphaproteobacterial marine methanotroph within the genus, Methyloceanibacter, capable of methane oxidation by solely the soluble methane monooxygenase, has also been described (Vekeman et al. 2016). The non-proteobacterial methanotrophs of the phylum Verrucomicrobia, which constitute a relatively newly described clade of methane utilizers, include three genera: Methylacidiphilum, Methylacidimicrobium, and Methyloacida (Dunfield et al. 2007; Islam et al. 2008; Van Teeseling et al. 2014). These bacteria typically inhabit geothermal and acidic environments, where they thrive at pH 0.5–5 (optimum 2.0), 30–65  C (optimum temperatures between 35  C and 50  C). Unlike proteobacterial methanotrophs, which prefer to assimilate carbon from reduced forms of C1 carbon, the members of this group are autotrophs and use methane only as a source of energy, oxidizing it to CO2 and then fixing CO2 using the CBB cycle (Khadem et al. 2011). Finally, the methanotroph “Candidatus Methylomirabilis oxyfera” is a member of the deep-branching bacterial phylum NC10. It generates dioxygen from nitrite for in situ monooxygenation of methane despite being an obligate anaerobe (Ettwig et al. 2010). The complete list of methanotrophs validated today is presented on the Methanotroph Commons website http://www.methanotroph.org.

pMMO Not known pMMO +/ sMMO RuMP

pMMO +/ sMMO RuMP RuMP RuMP RuMP/CBB/ serine RuMP RuMP RuMP

pMMO pMMO

pMMO + sMMO pMMO pMMO pMMO pMMO + sMMO pMMO

γ-Proteobacteria γ-Proteobacteria

γ-Proteobacteria

γ-Proteobacteria pMMO γ-Proteobacteria pMMO γ-Proteobacteria pMMO + sMMO pMMO

γ-Proteobacteria γ-Proteobacteria

γ-Proteobacteria

γ-Proteobacteria γ-Proteobacteria

γ-Proteobacteria

γ-Proteobacteria γ-Proteobacteria

γ-Proteobacteria

Methylosoma Methylomicrobium

Methylomarinum Methyloparacoccus

Methylomonas

Methylosarcina Methylosphaera Methylococcus

Methylogaea

Methylovulum Methyloprofundus

Methylocaldum

Methyloglobulus Methylomagnum

Methyloterricola

RuMP/CBB

RuMP/CBB/ serine RuMP RuMP/CBB

RuMP RuMP

RuMP

pMMO

γ-Proteobacteria

C1 assimilation

MMO type

Phylogeny

Genus Family Methylococcacea Methylobacter

Table 1 Classification of genera of aerobic methanotrophs

Type I

Type I Type I

Type I

Type I Type I

Type I

Type I NDc Type I

Type I

Type I Type I

Type I Type I

Type I

ICM typea

nfH+

Yes No

No

nifH+ Yes

nifH+

No Yes Yes

Some

No No

Yes No

No

N2 fixation

61

47.7 63–64

57

49.3 40.5

63.1

54 43–46 59–66

51–59

50.9–51.7 65.6

49.9 49–60

49–54

G+C (mol %) Trophic niche

Thermophilic

Psychrotolerant Psychrotolerant

Not extreme

16:1 Psychrotolerant 16:1/16:0/ Not extreme 14:0 16:1/16:0 Not extreme

16:0/16:1/ 15:0 16:0/14:0 16:1/16:0/ 16:2 16:1

Some psychrophilic 16:1 Not extreme 16:1 Halotolerant/ alkalophilic 16:1/16:0 Halotolerant 16:1/16:0/ Not extreme 14:0 16:1 Some psychrophilic 16:1 Not extreme 16:1 Psychrophilic 16:1 Thermophilic

16:1

Major PLFAb

76 V. N. Khmelenina et al.

Verrucomicrobia

pMMO pMMO

Verrucomicrobia Verrucomicrobia CBB CBB

pMMO pMMO

b

CBB CBB

NA4 type I/ NA4 NA4

NA4 Type III NAd No

sMMO pMMO sMMO sMMO

α-Proteobacteria α-Proteobacteria α-Proteobacteria α-Proteobacteria Serine Serine CBB/serine Serine

Type II Type II

α-Proteobacteria pMMO +/ sMMO Serine α-Proteobacteria pMMO + sMMO Serine

Type I Type I Type I

Type I Type I

RuMP RuMP RuMP

pMMO +/ sMMO RuMP pMMO

γ-Proteobacteria γ-Proteobacteria

γ-Proteobacteria pMMO γ-Proteobacteria pMMO γ-Proteobacteria pMMO

ICM intracellular membrane PLFA phospholipid fatty acid c ND not determined d NA not applicable because ICMs are very limited in this genus

a

Methyloacida Phylum NC10 “Candidatus Methylomirabilis oxyfera”

Family Crenothrichaceae Crenothrix Clonotrix Family Methylocystaceae Methylocystis Methylosinus Family Beyerinkiacea Methylocella Methylocapsa Methyloferula Methyloceanibacter Family Methylacidiphilaceae Methylacidiphilum Methylacidimicrobium

Family Methylothermacea Methylothermus Methylohalobius Methylomarinovum

Yes

Yes Yes Yes Yes

Yes Yes

No No No

60.9–63.8

60–61 63.1 55.6–57.5 64

62–67 63–67

62.5 58.7 66

18:0

18:0 18:0

18:1 18:1 18:1 18:1

18:1 18:1

18:1/16:0 18:1 18:1/16:0

Acidophilic

Acidophilic Acidophilic

Acidophilic Acidophilic Acidophilic Not extreme

Some acidophilic Not extreme

Not extreme Not extreme

Thermophilic Halophilic Thermophilic/ halotolerant

3 Physiology and Biochemistry of the Aerobic Methanotrophs 77

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2

V. N. Khmelenina et al.

Methane Oxidation

Methanotrophs oxidize methane to methanol in the presence of oxygen, with the production of one water molecule and an input of two electrons (Fig. 1). The reaction is catalyzed by methane monooxygenase (MMO), which exists in two structurally and biochemically distinct forms, particulate methane monooxygenase (pMMO) and soluble methane monooxygenase (sMMO).

2.1

sMMO

sMMO comprises three components: the active site-containing hydroxylase (MMOH), a reductase (MMOR), and a regulatory protein (MMOB) which is required for efficient methane oxidation. MMOH is an (αβγ)2 homodimer, of which the α subunit contains a di-iron center coordinated by four glutamates, two histidines, and several water molecules. Electrons for methane oxidation are transferred from NADH to this active site of MMOH via FAD and [2Fe-2S] clusters in MMOR. The mechanism of sMMO has been reviewed in depth recently (Lawton and Rosenzweig 2016). Briefly, the di-iron(III) site of MMOH (MMOHox) is reduced by MMOR in two sequential electron transfer events to the di-iron(II) state (MMOHred). Oxygen reacts with MMOHred to form intermediate O, followed by the peroxo intermediates P*, a di-iron(II) species, and P, a peroxo-bridged di-iron (III) species that is converted to the di-iron(IV) intermediate Q, which is defined by its characteristic absorption feature at 420 nm. Intermediate Q is believed to react with methane resulting in the formation of the product complex, T (Brazeau and Lipscomb 2000; Lee et al. 1993). The structures of MMOHox and MMOHred have been determined by X-ray crystallography (Rosenzweig et al. 1993), but molecular details of the intermediates have been elusive due to their transient nature (Banerjee et al. 2015). MMOB increases the reaction rate of sMMO with dioxygen by 2–3 orders of magnitude. The binding of MMOB to MMOH alters the electronic

Fig. 1 Pathways of carbon metabolism in methanotrophs

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structure and reduces the reduction potential of the di-iron center (Lee et al. 2013; Wang and Lippard 2014; Banerjee et al. 2015). The facultative methanotroph Methylocella silvestris BL2 possesses two distinct di-iron center monooxygenase gene clusters, one encoding sMMO and the other encoding a propane monooxygenase (PrMO) (Crombie and Murrell 2014). During growth on a mixture of these gases, the strain efficiently consumes both gases at the same time. Such metabolic flexibility may be important in many environments where methane and short-chain alkanes co-occur.

2.2

pMMO

pMMO is a copper-containing, membrane-associated enzyme that consists of three polypeptides with molecular masses of 49, 27, and 22 kDa, encoded by the pmoB, pmoA, and pmoC genes usually organized in the operon pmoCAB. The enzyme has an (αβγ)3 stoichiometry. The N-terminus of PmoB is located in the periplasm, whereas the N-termini of PmoA and PmoC are cytoplasmic. The active site is a copper center coordinated by three histidine residues in the N-terminal periplasmic region of PmoB (Balasubramanian et al. 2010). A second metal-binding site is located in PmoC. The native reductant of pMMO may be ubiquinol generated by a type II NADH:quinone oxidoreductase. Another possibility is that methanol oxidation by methanol dehydrogenase (MDH) in the periplasm is coupled to methane oxidation, providing electrons via the electron acceptor of MDH, cytochrome cL. pMMO and MDH encoded by the mxa operon form a supercomplex anchored in the intracytoplasmic membranes, and that electron transfer from the PQQ-linked MDH to pMMO in vivo may drive the oxidation of methane (Culpepper and Rosenzweig 2014; Torre et al. 2015). Within individual genomes, pmoCAB operons are present often in multiple copies which can be substantially sequence divergent and might encode pMMO isozymes of alternative physiological functions. In Methylocystis strain SC2, the enzyme encoded by pmoCAB2 oxidizes methane at a lower apparent Km than the enzyme encoded by pmoCAB1 (Baani and Liesack 2008). Methanotrophs in the genera Methylomonas, Methylobacter, and Methylomicrobium also encode a sequencedivergent particulate monooxygenase (pXMO) whose genes are uniquely organized in the noncanonical form pxmABC and whose primary substrate could be a compound other than methane or ammonia (Tavormina et al. 2011). The growth on methanol of Methylomicrobium album BG8 possessing a pxm operon is enhanced by chloromethane (Han and Semrau 2000).

2.3

Methanobactins as Vital Chalkophores

In methanotrophs that produce both the soluble and particulate MMO, the expression of these enzymes is regulated by copper ions: pMMO is expressed during growth under high copper-to-biomass ratios, whereas the soluble form of the enzyme is

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expressed when the copper-to-biomass ratio is low (Murrell et al. 2000). Copper availability in the growth medium affects many other physiological features of methanotrophs: cell wall and membrane synthesis, poly-3-hydroxybutyrate accumulation, biomass yield, and carbon conversion efficiency (Kao et al. 2004; Pieja et al. 2011). It also controls the “surfaceome,” or proteins on the outer surface of the outer membrane, and the expression of several multi-c-type cytochromes and proteins involved in copper uptake (Bergmann et al. 1999; Karlsen et al. 2011; Shchukin et al. 2011). One way that methanotrophs meet their high copper requirement is via the biosynthesis and release of high-affinity copper-binding compounds called methanobactins (Mbns). Mbns, termed “chalkophores” because of their ability to bind copper, are low-molecular-mass (1021 М 1). Following binding, Mbns rapidly reduce Cu2+ to Cu1+. Mbns may have a more significant role in copper uptake in situ, where copper speciation and distribution will be much more complex and will include copper associated with a wide range of organic materials (e.g., humic and fulvic acids), as well as copper found either sorbed onto or as part of various mineral phases (Fru et al. 2011). Mbns are secreted into the medium in an apo-form and transported into bacterial cells as a Cu-Mbn complex. Mbns can extract copper from insoluble minerals and could be important for mineral weathering. In addition to binding copper, Mbns bind most transition metals and neartransition metals and protect the host methanotroph as well as other bacteria from toxic metals. In addition to Cu(II) or Cu(I), Mbns bind other metals including Hg (II), Ag(I), Au(III), Co(II), Cd(II), Fe(III), Hg(II), Mn(II), Ni(II), Pb(II), U(VI), or Zn (II) but not Ba(II), Ca(II), La(II), Mg(II), and Sr(II). However, the binding constants for other metals are lower than that for Cu(II). Therefore, the growth and activity of methanotrophs synthesizing Mbn can be accompanied by solubilization and in situ immobilization of many metals, thus decreasing their toxicity to other components of the microbial community. In particular, Mbn reduces Hg(II) into Hg(0); in this case mercury does not evaporate and remains associated with methanobactin and bound to biomass of methanotrophs. This defines the potential ecophysiological significance of Mbn-producing methanotrophs for bioremediation of ecosystems polluted with heavy metals (reviewed in DiSpirito et al. 2016). Several Mbn biosynthesis genes and sequences encoding peptide precursors have been identified. Along with MbnA and two putative genes participating in biosynthesis – MbnB and MbnC – the operon encodes an aminotransferase, a

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sulfotransferase, an FAD-dependent oxidoreductase, and transporters, including a multidrug resistance exporter and a TonB-dependent transporter (Semrau et al. 2013). Operons encoding the MbnA are present in non-methanotrophic bacteria as well, suggesting a broader role in and perhaps beyond copper acquisition. Mbn-like compounds are found in yeast mitochondria suggesting that these molecules are a more universal phenomenon. Mbn can be used for copper elimination from semiconductor industry wastes, treatment of Wilson disease, copper metabolism disorders in humans, and copper extraction from insoluble minerals (DiSpirito et al. 2016). A new family of copper storage proteins (Csps) which provide an internal copper source when copper becomes limiting has been discovered in Methylosinus trichosporium OB3b (Vita et al. 2015). It possesses three Csps: Csp1 and Csp2, which have predicted twin arginine translocase-targeting signal peptides and are therefore thought to be exported after folding, as well as the cytosolic Csp3. Csp1 forms a tetramer of four-helix bundles that can bind up to 52 Cu+ ions via Cys residues. Switchover to sMMO is accelerated in the Δcsp1 csp2 mutant compared to the wild type, suggesting that Mbn produced under such conditions can readily remove all Cu1+ from Csp1 and therefore may play a role in helping to utilize Csp1bound copper.

3

Methanol Oxidation

Methanol dehydrogenase (MDH) catalyzes the second step in microbial methane conversion. Methanotrophs and gram-negative methanol-utilizing bacteria possess an MDH with pyrroloquinoline quinone (PQQ) at its catalytic center. This MDH belongs to the broad class of eight-bladed β-propeller quinoproteins, which comprise a range of other alcohol and aldehyde dehydrogenases. A well-studied MDH is the heterotetrameric MxaFI-MDH comprised of two large catalytic subunits (MxaF) and two small subunits (MxaI). It binds calcium as a cofactor that assists PQQ in catalysis. The purified MxaFI is most active at alkaline pH (pH 9–11) in the presence of ammonium or methylamine as activator. MxaFI exhibits wide substrate specificity to primary C1–C5 alcohols with the highest affinity to methanol (Km = 20–70 μM). Oxidation of methanol is coupled with reduction of the prosthetic group PQQ to the corresponding quinol (PQQOH2) followed by a 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 (Anthony and Williams 2003). Another MDH, known as XoxF-MDH, is often present along with MxaFI in methanotrophs and various methylotrophs as well as in some gram-negative nonmethylotrophic bacteria (Chistoserdova 2011, 2016). XoxF sequences exhibit approximately 50% amino acid identity with MxaF sequences. XoxF-MDHs are homodimeric proteins lacking the small subunit and possess a rare-earth element (REE) instead of calcium. In the structure of XoxF from the thermoacidophilic methanotroph Methylacidiphilum fumariolicum, the ligands for binding Ln3+ ions can be recognized. Two of them are conserved with respect to those binding Ca2+ in

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MxaFI-type MDH, while an additional ligand is specific to the XoxF enzyme (Khadem et al. 2012). The accommodation of an REE requires the presence of a specific aspartate residue near the catalytic site. XoxF from M. fumariolicum has an extraordinary high affinity for methanol with an affinity constant as low as 0.8 μM; it does not need ammonium activation and is active at neutral pH (optimum pH of 7.0), conditions which correspond to natural condition within the cell. In all cases when tested, only the lighter lanthanides (such as La3+ and Ce3+) support activity, while the heavier lanthanides are inactive (Pol et al. 2014; Vu et al. 2016). The presence of a suitable REE confers a superior catalytic efficiency on XoxF-MDHs. Lanthanides increase xoxF expression and decrease mxa expression, completely blocking mxa expression at micromolar concentrations (Chu and Lidstrom 2016). The response regulator MxaB controls transcription of the two methanol dehydrogenase genes (and its own expression) in response to lanthanides. Besides its catalytic function, a regulatory function has been suggested for XoxF. In M. extorquens, while removal of either of the two highly similar XoxF proteins caused no phenotype in terms of expression of the Ca2+MDH, removal of both abolished transcription from mxaF. However, no DNA-binding motifs are present in its sequence; thus, a more complex regulatory mechanism must be involved. The rare-earth elements, such as lanthanum, cerium, praseodymium, and neodymium, are relatively abundant in the Earth’s crust. However, lanthanide concentrations found in the environment might not be sufficient to repress mxa transcription (Chu and Lidstrom 2016). XoxF-MDHs are abundant in genomes of methylotrophic bacteria and also in organisms that hitherto are not known for a methylotrophic lifestyle. xoxF-type genes have been detected at high abundance in natural communities of lake sediments, plant phyllosphere, nutrient-rich coastal ocean waters, and plume waters off a hydrothermal vent in the deep ocean sea sponge microbiome (Delmotte et al. 2009; Mattes et al. 2013; Moitinho-Silva et al. 2014; Sowell et al. 2011).

4

Formaldehyde and Formate Oxidation

A major portion of the reducing equivalents required for methane oxygenation is formed during formaldehyde oxidation to CO2. Three pathways for formaldehyde oxidation in methanotrophs are predicted: (1) oxidation through dye-linked heme-containing formaldehyde dehydrogenase (Patel et al. 1980), (2) tetrahydromethanopterin (H4MPT)- and tetrahydrofolate (H4F)-mediated C1 transfers (Vorholt 2002), and (3) the minor dissimilatory ribulose monophosphate (dRuMP) pathway (Trotsenko and Murrell 2008). The Xox-type and Mxa-type MDHs also play at least some role in formaldehyde oxidation. Moreover, the broad-specificity aldehyde dehydrogenases have been predicted by analyses of genomes, but their role in primary C1 oxidation remains to be validated. In proteobacterial methanotrophs, the H4MPT-mediated pathway generates 1 mol of NAD(P)H per 1 mol of formaldehyde converted to formate. The

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H4MPT-dependent formaldehyde conversion is initiated by the condensation of formaldehyde and the pterin cofactor to give the N5,N10-methylene derivative. The formaldehyde-activating enzyme (Fae) accelerates this spontaneous reaction in M. extorquens AM1, and its gene is present in all proteobacterial methanotrophs. N5,N10-methylene-H4MPT is oxidized to N5,N10-methenyl-H4MPT+ by NAD (P)+-dependent methylene-H4MPT dehydrogenase, MtdB, catalyzing an essentially irreversible exergonic reaction ( 13 kJ/mol). Methenyl-H4MPT cyclohydrolase (Mch) catalyzes conversion of methenyl-H4MPT into N5-formyl-H4MPT. Conversion of N5-formyl-H4MPT to formate is catalyzed by the formyltransferase complex, FhcA, FhcB, and FhcC, which exhibit sequence identity to the subunits FmdA, FmdB, and FmdC of formyl methanofuran (MFR) dehydrogenase from methanogenic and sulfate-reducing archaea. The H4MPT cofactor-based pathway for C1 transfer is not present in the genome of M. infernorum SolV. Formate produced through the H4MPT pathway may be oxidized into CO2 by NAD+-dependent formate dehydrogenase (FDH) or converted to methylene-H4F, the starting substrate for the serine cycle, via the reductive pathway involving formyl-H4F ligase (FtfL), methenyl-H4F cyclohydrolase (Fch), and methyleneH4F dehydrogenase (MtdA) (Crowther et al. 2008). This methylene-H4F synthesis pathway is dominating over assimilatory flux in M. extorquens AM1 in contrast to the direct condensation formaldehyde with H4F. Remarkably, Fch and MtdA enzymes present in all α- and γ-proteobacterial methanotrophs but verrucomicrobial methanotrophs do not possess methenyl-H4F cyclohydrolase and methylene-H4F dehydrogenase. Instead, like many other bacteria, verrucomicrobial methanotrophs apparently use the folD gene product to perform the same reactions. The formate dehydrogenase (FDH) serves as a terminal enzyme in the oxidative pathway. NAD+-dependent FDH purified from Ms. trichosporium OB3b consists of two types of polypeptides and functions in vitro as an electron donor for sMMO or nitrogenase (with the additional participation of ferredoxin-NAD+ reductase and ferredoxin). However, in genomes of methanotrophs, there are from one to four open reading frames encoding FDH-like proteins, and the roles of FDH isozymes warrant further study. The minor oxidative RuMP pathway is functioning in γ-proteobacterial methanotrophs. This route includes 3-hexulose 6-phosphate synthase (HPS), 6-phospho 3-hexulose isomerase (PHI), glucose-6-phosphate dehydrogenase (GPDH), and 6-phosphogluconate dehydrogenase (PGDH) and leads to the production of 2 mol of NAD(P)H and 1 mol of CO2 from 1 mol of formaldehyde.

5

Carbon Assimilation Pathways

Methanotrophs use three pathways for methane carbon assimilation: the serine cycle, the ribulose monophosphate (RuMP), and the ribulose bisphosphate (RuBP) cyclic pathways.

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The Serine Cycle

In α-proteobacterial methanotrophs, N5,N10-methylene-H4F is the entry point of reduced one-carbon compounds into the serine cycle (Fig. 2). N5,N10-methyleneH4F is formed either in the spontaneous reaction of formaldehyde with H4F or via an alternative route involving formyl H4F ligase. N5,N10-methylene-H4F reacts with glycine to produce serine by the action of the serine hydroxymethyltransferase (SHTM). The amino group of serine is then transferred by a specific serineglyoxylate aminotransferase (SGAT) to glyoxylate, thus forming glycine and hydroxypyruvate. Hydroxypyruvate reductase (HPR) converts hydroxypyruvate to glycerate, and ATP-glycerate kinase phosphorylates glycerate to 2-phosphoglycerate or 3-phosphoglycerate, followed by isomerization to phosphoenol pyruvate (PEP) which is subsequently carboxylated to oxaloacetate. The reduction of oxaloacetate 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 and 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. In the obligate methanotrophs lacking isocitrate lyase (ICL variant), glyoxylate can be regenerated via the formation of acetoacetyl-CoA and

Fig. 2 Pathways of carbon metabolism in gammaproteobacterial (Type I) methanotrophs. (Modified from Rozova et al. 2015a). TCA tricarboxylic acid, THF tetrahydrofolate, H4MPT tetrahydromethanopterin

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hydroxybutyryl-CoA, the known intermediates of the poly-β-hydroxybutyrate biosynthesis pathway, and also crotonyl-CoA and butyryl-CoA, the intermediates of fatty acid biosynthesis. This alternate glyoxylate regeneration pathway is now known as the ethylmalonyl-CoA pathway (EMCP) analogous to that described in Methylobacterium (Chistoserdova et al. 2009). EMCP shares reactions and enzymes with the serine cycle (malate thiokinase, malyl-CoA lyase), the tricarboxylic acid cycle (succinate dehydrogenase, fumarase), the polyhydroxybutyrate cycle (betaketothiolase, acetoacetyl-CoA reductase), and other metabolic pathways (ethylmalonyl-CoA mutase, propionyl-CoA carboxylase), in addition to the specific reactions such as ethylmalonyl-CoA mutase and crotonyl-CoA reductase/carboxylase. Some methanotrophs (Methylocella silvestris), however, use the glyoxylate shunt instead of the EMCP (Chen et al. 2010a). The serine cycle genes are typically located in clusters and are subject to coordinated regulation (Kalyuzhnaya and Lidstrom 2005). Interestingly, genomes of all alphaproteobacterial methylotrophs including methanotrophs, contain genes encoding phosphoribulokinase which phosphorylate ribulose-5-phosphate into ribulose-1,5-bisphosphate, regardless of the presence/absence of the functional CBB cycle. Ribulose-1,5-bisphosphate formed in the reaction is an essential metabolite for driving one-carbon assimilation via the serine pathway in M. extorquens AM1 (Ochsner et al. 2017).

5.2

The RuMP Cycle

The γ-proteobacterial methanotrophs assimilate methane carbon via the RuMP cycle (Fig. 3) that is initiated by a reaction catalyzed by 3-hexulosephosphate synthase (HPS) where formaldehyde is fixed with ribulose-5-phosphate to form (D-arabino)3-hexulose-6-phosphate. This very unstable product is rapidly isomerized to fructose-6-phosphate by 6-phospho-3-hexulose isomerase (PHI). In the sequenced genomes of γ-proteobacterial methanotrophs, there are from one to three operons consisting of hps and phi genes, and several species contain an additional hps-phi fused gene. HPS purified from M. capsulatus Bath has a high molecular mass (6  49 kDa subunits) that corresponded to the product of the hps-phi fused gene possessing synthase but is lacking in isomerase activity (Ferenci et al. 1974). The recombinant HPS obtained from Methylomicrobium alcaliphilum 20Z by expression of hps is a homodimeric enzyme (2  40 kDa) inhibited by AMP and ADP (Rozova et al. 2017). The recombinant bi-domain HPS-PHI protein of strain 20Z has low synthase activity but no isomerase activity. Disruption of the hps-phi fused gene did not affect the growth rate of the mutant M. alcaliphilum. Interestingly, the genomes of all RuMP pathway methylotrophs which are unable to grow on methane have no HPS-PHI fused proteins. The significance of the HPS-PHI bi-domain protein in methanotroph remains to be clarified. In contrast to HPS, the characteristics of PHI are poorly documented in methanotrophs. In the second part of the RuMP cycle, phosphohexoses are converted into C3 molecules via three mechanisms: the Entner-Doudoroff (ED) pathway,

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Fig. 3 Pathways of carbon metabolism in alphaproteobacterial (Type II) methanotrophs (Modified from Rozova et al. 2015a). EMC ethylmalonyl-CoA, TCA tricarboxylic acid, THF tetrahydrofolate, H4MPT tetrahydromethanopterin

Embden-Meyerhof-Parnas (EMP) glycolysis, and phosphoketolase pathway. In the ED pathway, fructose-6-phosphate is transformed via glucose-6-phosphate and 6-phosphogluconate into 2-keto-3-deoxy-6-phosphogluconate (KDPG), which is subsequently cleaved by KDPG-aldolase to pyruvate and glyceraldehyde3-phosphate (GAP). However, some γ-proteobacterial methanotrophs do not have ED-encoding genes. In the EMP pathway, fructose-6-phosphate is phosphorylated into fructose1,6-bisphosphate (FBP) by pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK). FBP aldolase cleaves FBP to GAP and dihydroxyacetone phosphate. In this pathway, the energy of PPi, a waste product of anabolic reactions, such as the synthesis of lipids, carbohydrates, proteins, and nucleic acids, is reutilized. Reutilization of PPi in the PPi-mediated glycolytic pathway significantly increases the predicted efficiency of one-carbon assimilation. Conversion of phosphoenolpyruvate into pyruvate is catalyzed by a pyruvate kinase that has unusual regulatory properties. The RuMP pathway intermediates such as glucose-6-phosphate, fructose6-phosphate, ribose-5-phosphate, ribulose-5-phosphate, or erythrose-4-phosphate

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stimulate activities of the enzyme from M. alcaliphilum 20-fold, whereas ATP, PPi, and Pi strongly inhibit the enzyme activity. Another PPi-dependent glycolytic enzyme, pyruvate-orthophosphate dikinase (PPDK), is encoded in genomes of methanotrophs facilitating full reversibility of the EMP pathway and its higher energetic efficiency in comparison to the classical glycolysis pathway. The functionality and predominance of the EMP pathway have been corroborated by transcriptomic studies, metabolomics, and 13C-label distribution analysis (Kalyuzhnaya et al. 2013). The third mechanism for phosphosugar cleavage in γ-methanotrophs involves phosphoketolase (Xfp) splitting fructose-6-phosphate (or xylulose-5-phosphate) into erythrose-4-phosphate (or glyceraldehyde-3-phosphate) and acetyl-phosphate. The reversible acetate kinase coded by the gene ack catalyzes ATP and acetate synthesis from acetyl-phosphate. The acetate kinase from M. alcaliphilum was 20-fold more active in the reaction of acetate and ATP synthesis compared to acetate phosphorylation and had a remarkably higher catalytic efficiency in this direction (Rozova et al. 2015b). In M. alcaliphilum 20Z, the xfp and ack genes are co-transcribed, and this phosphoketolase pathway is 1.5 times more efficient in anaerobic ATP generation compared to the classical glycolysis. Besides methanotrophs, most aerobic methylotrophs, chemoautotrophs, and cyanobacteria, i.e., microbes whose carbon assimilation proceeds by de novo formation of the C–C bond, possess the ack and xfp genes. The phosphoketolase pathway contributes to C2-compound production and bypasses the traditional glycolytic mechanism involving pyruvate dehydrogenase, thus preventing disruption of the C–C bond. Acetate accumulation during formaldehyde fermentation in cultures of M. alcaliphilum 20Z grown in microaerobic conditions suggests high metabolic flexibility that gives an advantage to gammaproteobacterial methanotrophs to survive in oxygen-limiting ecosystems. The biotechnological potential of the Xfp-based synthetic pathways has been highlighted in a recent publication (Bogodar et al. 2013). In the third part of the RuMP cycle, the primary acceptor of formaldehyde, ribulose-5-phosphate, is regenerated from glyceraldehyde-3-phosphate and fructose-6-phosphate in a series of transaldolase/transketolase reactions analogous to those in photo- and chemotrophic bacteria. Besides the RuMP cycle machinery, the key enzymes of the serine pathway, SHTM, SGAT, HPR, and malyl-CoA lyase, are encoded by genomes of all gammaproteobacterial methanotrophs sequenced so far, while the respective glyoxylate regeneration mechanism is not encoded (But et al. 2017).

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CBB Cycle

The methanotrophs of the phyla Verrucomicrobia and NC10 are autotrophic bacteria using the CBB cycle of CO2 fixation and using methane only as an energy source (Fig. 1). In contrast, in methanotrophs of the genera Methylococcus and Methylocaldum, three simultaneous pathways of C1 assimilation are probably

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functioning (Trotsenko and Murrell 2008). In these thermophilic/tolerant methanotrophs, the CBB cycle is a minor pathway whose contribution for C1 assimilation remains poorly understood. The ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) of Mc. capsulatus Bath has an α6β6 structure which differs from the typical α8β8 structure of the Form I RuBisCO found in Proteobacteria, Cyanobacteria, and higher plants. The genes encoding the large subunit (cbbL) and small subunit (cbbS) and putative regulatory gene (cbbQ) are colocated in one cluster. In Mc. capsulatus Bath and Methylocaldum szegediense O-12, RuBisCO activity is enhanced in response to temperature increase, thus providing a means for dissipation of excess heat energy (Eshinimaev et al. 2004).

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Nitrogen Assimilation

The ability to utilize atmospheric nitrogen (N2) as a sole nitrogen source has been experimentally proven in a number of proteobacterial methanotrophs as well as verrucomicrobial representatives (Murrell and Dalton 1983; Khadem et al. 2010). The genomes of many methanotrophs have a complete set of genes necessary for N2 fixation which encompass a gene cluster for iron-molybdenum-dependent nitrogenase (nifH, nifD, and nifK) and two additional clusters that contain genes for biogenesis of cofactors and electron transfer proteins, as well as a Mo/Fe nitrogenase-specific transcriptional regulator NifA. M. fumariolicum SolV is able to fix N2 under low oxygen concentration (0.5% O2 saturation) in chemostat cultures at a dilution rate of 0.017 h 1. The nitrogenase of M. fumariolicum SolV is extremely oxygen sensitive compared to those from the proteobacterial methanotrophs studied to date. Methanotrophs use ammonium, nitrates, and nitrites and often can grow in the presence of urea or some amino acids as nitrogen source. Gammaproteobacterial methanotrophs assimilate NH4+ mainly by reductive amination of pyruvate and/or α-ketoglutarate, whereas alphaproteobacterial methanotrophs use 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. Regulation of glnA in this methanotroph is analogous to that in enterobacteria and occurs via the Ntr system. In Mc. capsulatus Bath and other gammaproteobacterial methanotrophs grown on medium containing ammonia, the reductive amination of pyruvate (via alanine dehydrogenase) and/or 2-oxoglutarate (via glutamate dehydrogenase) occurs under high-ammonia growth conditions. In contrast, when grown under N2-fixing conditions, i.e., under ammonium limitation or on medium containing nitrate, the methanotrophs assimilate ammonia via the glutamate cycle (reviewed in Trotsenko and Murrell 2008). M. infernorum V4 can fix ammonia both through the glutamine synthesis reaction and through the carbamoyl-phosphate synthesis reaction. The latter substrate is used in the urea cycle, for which all genes are present except for the gene encoding arginase, which cleaves arginine into urea and ornithine.

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However, M. infernorum V4 encodes 4-aminobutyrate aminotransferase ArgD that can ultimately supply ornithine back to the cycle through a part of the TCA cycle and glutamate synthesis. Other methylotrophs possess neither arginase nor ArgD. Four predicted ammonium transporters have been identified in the genome of Mc. capsulatus Bath. Methanotrophs also possess genes encoding assimilatory nitrate reductase.

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Assimilation of Methylated Amines

Among methanotrophs, M. silvestris BL2 is able to grow on mono- or trimethylamine as a sole source of energy, carbon, and nitrogen. Also, trimethylamine N-oxide (TMAO) can serve as nitrogen source for this bacterium. Monomethylamine is metabolized via methylated amino acids in a metabolic pathway where γ-glutamylmethylamide (GMA) and N-methylglutamate (NMG) are intermediates (Chen et al. 2010b). However, methylamine dehydrogenase is not detectable. One of the key enzymes in this pathway, GMA synthetase, catalyzes the ATPdependent condensation of methylamine and glutamate, and a second enzyme, NMG dehydrogenase, delivers C1 unit from NMG into the serine cycle. Mutation of the GMA synthetase gene, gmas, abolishes growth of M. silvestris on methylamine. Eight genes of this pathway are co-transcribed as an operon. The flavin-containing trimethylamine monooxygenase (Tmm) oxidizes trimethylamine to trimethylamine N-oxide (TMAO) in M. silvestris BL2 (Chen et al. 2011). The Tmm contains the conserved sequence motif (FXGXXXHXXXF/Y) and typical domains for binding flavin adenine dinucleotide and nicotinamide adenine dinucleotide phosphate. TMAO demethylase (Tdm) catalyzes demethylation of TMAO to formaldehyde and dimethylamine (Zhu et al. 2014). It is a novel Zn2+ and Fe2+-dependent metalloprotein with the Zn2+/Fe2+/Tdm monomer ratio 1/1/1. It has been hypothesized that the oxygen atom is transferred from the substrate TMAO to produce formaldehyde via the high-valent iron-oxo intermediate (e.g., Fe(IV)oxo, Fe(V)-oxo) (Zhu et al. 2016). Hexameric Tdm of M. silvestris BL2 has a high affinity for TMAO (Km = 3.3 mM; Vmax = 21.7 nmol min 1 mg 1). It carries out an unusual O2-independent oxidative demethylation utilizing the substrate TMAO as a surrogate oxygen donor. Tdm of M. silvestris BL2 and eukaryotic Tdms have contrasting characteristics and no sequence homology.

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TCA Cycle

Since methanotrophs are able to obtain energy from the oxidation of reduced C1 compounds, the TCA cycle cannot be an obligatory means of energy generation. Due to low or zero activity of 2-oxoglutarate dehydrogenase in the α- and γ-proteobacterial methanotrophs, it has been assumed that the major function of the TCA cycle is to provide precursors for biomass synthesis in methanotrophs. Interestingly, the genome of the γ-proteobacterial methanotroph Methylomicrobium

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buryatense 5GB1 encodes three pathways for conversion of 2-oxoglutarate to succinyl-CoA or succinate: a classic 2-oxoglutarate dehydrogenase complex, 2-oxoglutarate ferredoxin oxidoreductase, and a pathway through succinate semialdehyde catalyzed by 2-oxoglutarate oxidase. All these sets of genes are expressed at levels similar to other TCA cycle genes. Moreover, operation of a complete, oxidative TCA cycle contributing about 45% of the total flux for de novo malate production has been revealed in M. buryatense 5GB1 by mutant analysis and monitoring of 13C-labeling patterns of metabolites in core metabolism (Fu et al. 2017). In M. buryatense 5GB1, the TCA cycle generates some energy for biosynthesis. In Ms. trichosporium OB3b, the TCA cycle enzyme malate dehydrogenase exhibits a remarkably higher catalytic efficiency in vitro for reduction of oxaloacetate than for oxidation of malate (Rozova et al. 2015a). The enzyme therefore allows the products of primary C1 assimilation to be converted to malate, a central intermediate of the serine pathway. In contrast, malate dehydrogenase from M. alcaliphilum 20Z displays about twofold higher catalytic efficiency toward malate oxidation over oxaloacetate reduction. These catalytic properties are in accordance with a high demand of the halotolerant bacterium for aspartate, which is a precursor of the osmoprotective compound ectoine. Overall, the biochemical properties of these two malate dehydrogenases suggest that the TCA cycle in obligate methanotrophs fulfills a predominantly anabolic function.

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Osmoadaptation Mechanisms

Methanotrophs inhabiting saline environments such as saline lakes and marine waters include species of the genera Methylomicrobium (M. alcaliphilum, M. buryatense, M. kenyense, and M. japanense), Methylobacter (M. marinus), and Methylohalobius (M. crimeensis). These halotolerant or halophilic methanotrophs synthesize and accumulate in their cytoplasm the cyclic amino acid ectoine (1,4,5,6-tetrahydro-2-methyl-4-pyrimidine carboxylic acid). Along with ectoine, M. alcaliphilum accumulates enhanced levels of glutamate and sucrose, the total intracellular concentration of which balances the external osmotic pressure. M. alcaliphilum 20Z is tolerant to 1.5 M NaCl and accumulates ectoine at up to 20% of cell dry weight and is a potential producer of this bioprotective compound for biotechnology. As in other halophilic bacteria, methanotrophs synthesize ectoine from aspartate and acetyl-CoA using three specific enzymes: diaminobutyric acid (DABA) aminotransferase (EctB), DABA acetyltransferase (EctA), and ectoine synthase (EctC). In M. alcaliphilum 20Z, the ectoine biosynthetic genes are organized in the ectR-ectABC-ask operon, which also encodes aspartokinase as well as a MarR-like negative transcriptional regulator EctR (Reshetnikov et al. 2011). The genome of the moderately halophilic methanotroph Methylohalobius crimeensis 10Ki encodes diverse genetic systems for osmotolerance, which

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include the ectABCD genes for ectoine and hydroxyectoine synthesis; a gene encoding a high-affinity importer of choline/glycine betaine driven by a sodiummotive force; three gene copies for choline dehydrogenase and a gene 40% identical to betaine aldehyde dehydrogenase from Bacillus subtilis, indicating possible glycine betaine synthesis from choline; and a pathway for sucrose synthesis and degradation/reutilization, including genes for sucrose-phosphate synthase, sucrose synthase, and fructokinase. Na+ export and use of a sodiummotive force is suggested by the presence of genes encoding a putative Na+/H+ antiporter localized within an ATP synthase-encoding gene cluster and a complete nqr gene cluster encoding Na+-pumping NADH:quinone oxidoreductase (Sharp et al. 2015). A mildly acidophilic, obligate methanotroph Methylocapsa palsarum NE2 harbors genes indicative of aerobic anoxygenic photosynthesis. This array of genes is highly similar to that in many plant-associated Methylobacterium species and includes genes encoding the light-harvesting complex (pufABCML), the reaction center (puhA), as well as genes involved in biosynthesis of bacteriochlorophyll and carotenoids (Miroshnikov et al. 2017). It remains to be seen if these genes are expressed under environmental conditions.

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Methanotrophs and Biotechnology

Methanotrophs are natural systems for the attenuation of methane emission and unique systems for methane-based bioconversions. The industrial potential of methanotrophs has been tested for the production of single-cell protein, polyhydroxyalkanoates, and lipids (potential biofuels) with methane as a feedstock for large-scale cultivation (Helm et al. 2006; Zuniga et al. 2013). Methanotrophs can generate components for nanotechnology applications (surface layers and nanoparticles), growth media, vitamin B12, and soluble metabolites (methanol, formaldehyde, organic acids, and ectoine) using methane as their carbon source. Other cell components, such as metal-chelating proteins (methanobactins), enzymes (methane monooxygenases), or heterologous proteins, hold promise as future products (DiSpirito et al. 2016; Strong et al. 2016). Advances in genetic methods for methane oxidizers have made possible the use of these bacteria as hosts for production of recombinant and heterologous proteins and low-molecular-mass products (de la Torre et al. 2015; Kalyuzhnaya et al. 2015; Kalyuzhnaya 2016). Genetically engineered methanotrophs can overproduce naturally occurring metabolites or non-native compounds, including such molecules as carotenoids, isoprene, 1,4-butanediol, farnesene, or lactic acid (Henard et al. 2016; Saville et al. 2014). Scenarios for generating multiple products from a single methanotroph are presented in the recent excellent publications (Gilman et al. 2015, 2017; Levett et al. 2016). A number of methane-driven transformations, such as epoxidation, calcium carbonate precipitation, bioleaching, and bioremediation, have also been explored (Jiang et al. 2010; Strong et al. 2015, 2016; Eswayah et al. 2017).

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Prospects for the Future

Our understanding of methanotroph diversity and methane oxidation mechanisms has changed dramatically during the last 20 years. New methanotroph strains with potentially novel metabolism have inspired new ideas for valorization of methane conversions and have stimulated new research into use of methanotrophs in biotechnology. Improved genetic systems together with highly active homologous expression systems for methanotrophs should allow metabolic engineering of methanotrophs, thus facilitating their considerable biotechnological potential. Structure-function studies on the sMMO will reveal more about the active site of this remarkable enzyme, and mutagenesis will enable its catalytic utility to be extended for the production of chiral alcohols and epoxides and degradation of larger aromatic compounds, particularly polyaromatic hydrocarbons. Another goal should be the expression of high-value heterologous proteins during the production of single-cell protein, thereby increasing the economic viability of large-scale bacterial fermentations during growth on methane. Isolation and characterization of new methanotrophs that can grow, for example, at high or low temperatures and more extreme pH may improve the potential use of methanotrophs in bioremediation and biocatalysis applications. Another much neglected area of research on methanotrophs is the study of membrane biogenesis. Despite their observation over 40 years ago, there is still much debate as to the exact function of the intracellular membranes of methanotrophs, and they provide an excellent subject for future study. The obligate versus facultative nature of methanotrophs can also be addressed more systematically through postgenomics, as can the pathways of formaldehyde assimilation and dissimilation. Proteomic analyses will provide a wealth of information for studying the regulation of methane oxidation and may provide insights into how methane oxidation is regulated under differing environmental conditions. Genome-wide transcriptomic studies, metabolomics, and 13C-label distribution analysis with methane-grown cultures will help us to understand regulation of metabolism in methanotrophs. Molecular biological and biochemical studies are needed to answer the long-standing question as to why obligate methanotrophs grow only on methane (and other one-carbon compounds) since they possess genes encoding putative membrane transport systems for organic acids and sugars. Future postgenomic studies, using methanotroph genome sequence information as a blueprint for hypothesis testing, will undoubtedly lead to further advances in our knowledge of the biology of these fascinating bacteria and allow further exploitation in biotechnology.

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Research Needs

More detailed physiological, biochemical, and molecular studies on the carbon assimilation and dissimilation pathways in obligate methanotrophs, especially newly isolated strains.

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Functional analysis of the genomes of newly isolated methanotrophs; transcriptomic and proteomic studies to investigate gene regulation in obligate methanotrophs. Synthetic biology of aerobic methanotrophs to engineer new pathways for production of commodity chemicals. Development of genetic techniques for key genera of aerobic methanotrophs in order to study environmental regulation of carbon and nitrogen metabolism. Environmental regulation of facultative methanotrophs and their growth on the various components of natural gas, methane, ethane, and propane.

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Biochemistry and Molecular Biology of Methane Monooxygenase Tim Nichol, J. Colin Murrell, and Thomas J. Smith

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Biochemistry of Particulate Methane Monooxygenase (pMMO) . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Biochemistry of Soluble Methane Monooxygenase (sMMO) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Molecular Biology and Regulation of Methane Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . 5 Methanotrophs in Biocatalysis and Bioremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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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 diiron monooxygenases. The expression of these enzymes in methanotrophs is often regulated by the availability of copper. The soluble methane monooxygenase has attracted significant attention T. Nichol (*) · T. J. Smith Biomolecular Sciences Research Centre, Sheffield Hallam University, Sheffield, UK e-mail: [email protected]; [email protected] J. C. Murrell School of Environmental Sciences, University of East Anglia, Norwich Research Park, Norwich, UK e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_5

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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 Gramnegative 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 (Fig. 1) (Anthony 1982; Dalton 2005; Hanson and Hanson 1996; Trotsenko and Murrell 2008; Lawton and Rosenzweig 2016). In methanotrophs there are two structurally and biochemically distinct forms of MMO, particulate methane monooxygenase (pMMO) and soluble methane monooxygenase (sMMO), which oxidize methane to methanol. pMMO is a

Fig. 1 Principal metabolic pathways of methanotrophic metabolism showing the roles of soluble and particulate methane monooxygenase (sMMO/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

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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 principal factor known to govern expression of the two types of MMO in these organisms is the concentration of available copper, and copper homeostasis is carefully regulated within methanotrophs through copper uptake systems and copper storage mechanisms (Vita et al. 2015, 2016; Gu and Semrau 2017). 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 previously the dogma was that all methanotrophs contained pMMO. More recently, however, the facultative type II methanotroph Methylocella silvestris and obligate methanotroph Methyloferula stellata have been shown to possess only the sMMO system and do not possess pMMO (Dedysh et al. 2005; Theisen et al. 2005; Vorobev et al. 2011; Crombie and Murrell 2014; Dedysh et al. 2015). While the majority of methanotrophs are aerobic, Methylomirabilis oxyfera appears to grow anaerobically through oxidation of methane via pMMO using O2 generated in situ from nitrite (Welte et al. 2016). The 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 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; Ross and Rosenzweig 2017), 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-β-D-maltoside as detergent (Smith and Dalton 1989), however, allows recovery of activity after solubilization, and subsequent development of purification protocols has 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 previously it was thought that the active site resided on this subunit.

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More recent structural studies suggest, however, that the active site may reside on the 49-kDa subunit or indeed may be shared between more than one subunit (reviewed in Hakemian and Rosenzweig 2007, Balasubramanian et al. 2010). The 49-, 27-, and 22-kDa components are encoded by the genes pmoB, pmoA, and pmoC, respectively, which are multicopy genes (see below) that are induced in response to growth of methanotrophs at high copper-to-biomass ratio. The crystal structure of pMMO, albeit of protein of rather low activity, showed that the enzyme has an (αβγ)3 stoichiometry and gave 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). While all active preparations of pMMO contain copper, the numbers and roles of copper ions in the active form of the enzyme continue to be debated, and it has also been suggested that iron plays a role in pMMO (Martinho et al. 2007 reviewed in Hakemian and Rosenzweig 2007; Semrau et al. 2010; Ross and Rosenzweig 2017). Recent quantum refinement of the crystal structure data suggests a mononuclear copper center in the crystallized form of the protein (Cao et al. 2018). An expression system for pMMO has been developed within Escherichia coli, which is capable of producing active PmoB, capable of methane oxidation (Balasubramanian et al. 2010). This has allowed further insights into the nature of the active site and enables future site-directed mutagenesis studies to elucidate the exact catalytic mechanism. Studies using protein refolding of truncated recombinant PmoB with metal ions suggest that only copper is required for catalysis and the addition of iron does not restore or increase activity (Balasubramanian et al. 2010). 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 catalyze 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).

3

Biochemistry of Soluble Methane Monooxygenase (sMMO)

sMMO is a three-component binuclear iron active center 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.

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2006; Nichol et al. 2015; Trehoux et al. 2016), 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 main components: (1) a 250-kDa hydroxylase with an (αβγ)2 structure (encoded by mmoX, mmoY, and mmoZ, respectively) – MmoX contains the binuclear iron active center 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 (Fig. 2) (Smith and Dalton 2004; Smith and Murrell 2008; Sazinsky and Lippard 2015; Sirajuddin and Rosenzweig 2015; Lee 2016). The 12-kDa component MmoD works in conjunction with the chalkophore methanobactin to regulate the expression of sMMO during low copper concentration (Semrau et al. 2013, 2018; DiSpirito et al. 2016). 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). Crystallography of the complex formed between the hydroxylase and protein B (Fig. 3) (Lee et al. 2013) indicates that binding of protein B induces changes in hydroxylase conformation that may allow substrate entry and product egress. They may also facilitate proton transfer required by the catalytic cycle. The catalytic cycle of sMMO has been extensively studied, and excellent progress has been made toward understanding the mechanism of oxygen and Fig. 2 Schematic of the sMMO enzyme complex

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Fig. 3 Structure of the hydroxylase component of sMMO of Mc. capsulatus (Bath). The α, β, and γ subunits are shown in blue, green, and yellow, respectively. The iron atoms of the diiron centers are shown as orange spheres. Protein B is shown as a pink ribbon diagram at its binding position to the α and β subunits

hydrocarbon activation at the binuclear iron center. 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; Sazinsky and Lippard 2015; Ross and Rosenzweig 2017). 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 center of compound Q is in a high-valent diferryl (FeIV-FeIV) state (Banerjee et al. 2015). It may have a six-membered ring-bridged structure rather than the four-membered ring “diamond core” structure proposed previously (Castillo et al. 2017). 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; Jasniewski and Que 2018). Radical, ionic, and concerted mechanisms have been suggested. Evidence from the use of radical clock substrates and theoretical studies suggests a reaction with multiple pathways and the possible involvement of a captive substrate-derived

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radical species (Sazinsky and Lippard 2015). Results using stopped-flow spectroscopy have established the involvement of quantum mechanical tunneling of hydrogen nuclei in breaking the C-H bond of methane (Zheng and Lipscomb 2006; Tinberg and Lippard 2010). 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 di-aromatics must be able to gain access to this cavity and the adjacent binuclear iron center (Zhang et al. 2017). The side chain of residue Leu 110 in the α-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 center 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, while in the “open” conformation, it permits a 2.6-Å diameter channel into the substratebinding 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). Sitedirected 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; Sigdel et al. 2015). Recently crystal structures of MmoB bound MmoH have indicated a change in the conformation of Phe 188 upon binding of MmoB. This suggests that Leu 110 and Phe 188 conformations, mediated by MmoB binding, have a role in controlling substrate access to the active site (Lee et al. 2013). At the time of writing, a study made available as a preprint (Cho et al. 2018) reports crystallographic data showing that binding of MmoD to the hydroxylase also opens this potential substrate access route, although the crystal structure data are not currently available. While 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 substratebinding pocket that can accommodate a wide range of oxidation substrates in addition to the natural substrate methane. Recent advances in understanding the interaction between the sMMO components may inform future mutagenesis studies to more effectively manipulate the selectivity and catalytic properties of the enzyme.

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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 the homologous genes amoCAB, encoding the ammonia monooxygenase (AMO) in nitrifying bacteria, has also been observed, and it has been suggested that both

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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. Type II methanotrophs with very different pmoA genes have also been 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). 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 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 σn-(σ54)-dependent promoter which requires a transcriptional activator for the formation of an active transcriptional complex. Located near the structural genes in the sMMO gene 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 σN (RpoN) to promoters which are regulated by this alternative σ 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 (Csaki 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 σN 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

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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 σ70 activated promoter located 50 of pmoC. In the absence of copper ions, pMMO genes are still expressed, albeit at lower levels, but the apoenzyme 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 multicarbon substrates such as acetate (Crombie and Murrell 2014). 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). Recent evidence suggests a model for the copper switch mechanism and regulation of the sMMO operon and pMMO operon which involve methanobactin and MmoD. At low copper ion concentration, MmoD acts to repress the pMMO operon and also upregulates expression of the mbn operon to produce methanobactin. Methanobactin in turn increases the expression of the mmo (sMMO-encoding) operon which further represses pMMO expression. In the presence of excess copper ions, methanobactin is bound to copper and is unable to upregulate sMMO expression. The MmoD protein also binds copper and is unable to repress pMMO (Semrau et al. 2013, 2018; DiSpirito et al. 2016). The identification of a constitutively sMMO-expressing mutant of Ms. trichosporium with a deletion of part of the copD gene led to the suggestion that the copCD system is involved in copper regulation in methanotrophs (Kenney et al. 2016). The copCD genes encode for a copper-binding protein and inner membrane protein, respectively, and are utilized by other bacteria for copper uptake. However specific knockout mutants of copCD in M. trichosporium OB3b suggest this is not the case (Gu et al. 2017).

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

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the cells. In addition, the presence of oxygen-stable hydrogenase activity in methanotrophs enables hydrogen to be used as the reductant. 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 nonenzymatic 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 in the 1990s by Dalton and co-workers toward developing sMMO-expressing 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 were filed worldwide. The process was also evaluated for production of 1,2-epoxybutane from but-1-ene and acetaldehyde from ethane. sMMO and pMMO are attractive biocatalysts for conversion of methane to the liquid fuel methanol, which is fuel with a higher energy density than methane that is also easier to store and transport (Bjorck et al. 2018). Development of a suitable cellfree MMO system or cells engineered to minimize onward metabolism of methanol may enable such technology. Recently a site-directed mutagenesis study of MmoX has identified a mutant R98L that abolishes a salt bridge on the periphery of the hydroxylase (Fig. 4). This mutant has increased activity toward aromatic substrates and altered regioselectivity for more precise hydroxylation of the substrate biphenyl (Lock et al. 2017). This unique enzyme co-oxidizes a wide range of organic substrates, and it will be

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Fig. 4 Structure of the hydroxylase component of sMMO from Ms. trichosporium OB3b showing the position of the mutated residue Arg 98 and Asp 365, with which it forms an ionic interaction

interesting to ascertain if its 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. 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-catalyzed 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 methane-oxidizing 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-tobiomass ratio (to promote sMMO expression) in order to purify effluent contaminated with TCE and cis-1,2-dichloroethylene (cDCE). Here, up to 99% removal of

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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 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). By utilizing gene probe hybridization, it was suggested that the majority of TCE biodegradation at a field test site in Carolina, USA, was carried out by sMMO-expressing bacteria (Hazen et al. 2009). It has been shown that a facultative methanotroph Methylocystis strain SB2 constitutively expresses pMMO when grown on multicarbon substrates and is able to degrade a variety of chlorinated hydrocarbons including TCE, trans-dichloroethylene, vinyl chloride, and 1,1,1trichloroethane (Im and Semrau 2011; Yoon et al. 2011). Increasing understanding of the way methanotrophs expressing pMMO and sMMO interact with other microorganisms in complex communities is expected to lead to further exploitation of cells expressing these enzymes in bioremediation and other biotechnologies. Other possibilities for bioremediation using methanotrophs include use of sMMO-expressing 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. Recently an expression system for pMMO has been developed within Escherichia coli, which is capable of producing active PmoB, capable of methane oxidation (Balasubramanian et al. 2010). This has allowed further insights into the nature of the active site and enables future site-directed mutagenesis studies to elucidate the exact catalytic mechanism. The exact nature and function of the copper centers in pMMO still need to be further 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 MMO regulation, copper sequestration, and delivery of copper ions to the active site of pMMO which is still unclear (DiSpirito et al. 2016; Kenney and Rosenzweig 2018). 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 multicarbon

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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. The ability of methanotrophs to produce valuable bioproducts using methane as a feedstock is another focus for biotechnological research. It has been shown that methanotrophic bacteria can produce a variety of valuable products such as liquid biofuel, polyhydroxyalkanoate bioplastics, single-cell protein for animal feed, and other bioproducts such as ectoine and vitamin B12 (Strong et al. 2015; Pieja et al. 2017; Cantera et al. 2018), and this is showing great potential for future development of MMO and methanotrophs as a commercially viable biotechnology.

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Trotsenko YA, Murrell JC (2008) Metabolic aspects of aerobic obligate methylotrophy. Adv App Microbiol 63:183–229 Vita N, Platsaki S, Baslé A, Allen SJ, Paterson NG, Crombie AT, Murrell JC, Waldron KJ, Dennison C (2015) A four-helix bundle stores copper for methane oxidation. Nature 525:140–143 Vita N, Landolfi G, Baslé A, Platsaki S, Lee J, Waldron KJ, Dennison C (2016) Bacterial cytosolic proteins with a high capacity for Cu (I) that protect against copper toxicity. Sci Rep 6:39065 Vorobev AV, Baani M, Doronina NV, Brady AL, Liesack W, Dunfield PF, Dedysh SN (2011) Methyloferula stellata gen. nov., sp. nov., an acidophilic, obligately methanotrophic bacterium that possesses only a soluble methane monooxygenase. Int J Syst Evol Microbiol 61:2456–2463 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 Welte KU, Rasigraf O, Vaksmaa A, Versantvoort W, Arshad A, Op den Camp HJM, Jetten MSM, Lüke C, Reimann J (2016) Environ Microbiol Rep 8:941–955 Yoon S, Im J, Bandow N, DiSpirito AA, Semrau JD (2011) Constitutive expression of pMMO by Methylocystis strain SB2 when grown on multi-carbon substrates: implications for biodegradation of chlorinated ethenes. Environ Microbiol Rep 3:182–188 Zahn JA, DiSpirito AA (1996) Membrane-associated methane monooxygenase from Methylococcus capsulatus (Bath). J Bacteriol 178:1018–1029 Zhang S, Karthikeyan R, Fernando SD (2017) Low-temperature biological activation of methane: structure, function and molecular interactions of soluble and particulate methane monooxygenases. Rev Environ Sci Biotechnol 6:611–623 Zheng H, Lipscomb JD (2006) Regulation of methane monooxygenase catalysis based on size exclusion and quantum tunneling. Biochemistry 45:1685–1692

5

Enzymes for Aerobic Degradation of Alkanes in Bacteria Renata Moreno and Fernando Rojo

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Uptake of n-Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 The Low Water Solubility of Hydrocarbons Presents a Problem for their Uptake by Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Transport of Hydrocarbons Through the Cell Envelope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Chemotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 n-Alkane Degradation Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Hydroxylation of n-Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Alkane Hydroxylases Related to Methane Monooxygenase . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 The AlkB Family of Alkane Hydroxylases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Cytochrome P450 Alkane Hydroxylases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Alkane Hydroxylases for Long-Chain n-Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Several Alkane Hydroxylases Frequently Coexist in a Single Bacterial Strain . . . . . 5 Metabolism of the Alcohols and Aldehydes Derived from the Oxidation of n-Alkanes . . 6 Degradation of Branched-Chain Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Applications of Alkane Oxidation Enzymes in Biotransformations of Industrial Interest 8 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

118 119 119 120 122 123 124 125 126 128 129 130 131 133 133 134 134

Abstract

Alkanes are major constituents of crude oil but they are also present at low concentrations in diverse noncontaminated habitats since many living organisms produce them as chemoattractants or as agents that help to protect against water loss. Although the metabolism of these compounds poses problems (mainly to do with their hydrophobicity), many microorganisms can use them as a carbon and energy source. This chapter examines how bacteria metabolize n-alkanes R. Moreno · F. Rojo (*) Centro Nacional de Biotecnología, CSIC, Madrid, Spain e-mail: [email protected]; [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_6

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aerobically, paying particular attention to the enzymes involved in the initial oxidation of the alkane molecule – the most critical step given that n-alkanes are chemically rather inert.

1

Introduction

Alkanes are saturated hydrocarbons. They can be linear (n-alkanes), cyclic (cycloalkanes), or branched (iso-alkanes). These compounds are major constituents of crude oil but are also produced by many living organisms, including bacteria, green algae, plants and animals, in which they serve as chemoattractants or in protection against water loss, insects, or pathogens (Post-Beitenmiller 1996; Schirmer et al. 2010; Lea-Smith et al. 2015; Pedrini et al. 2013). Alkanes are therefore found in small amounts in most soils and water, in which ongoing biosynthesis and biodegradation keep their concentrations relatively constant. Alkanes are highly reduced molecules with a high energy and carbon content; they are therefore good carbon and energy sources for any microorganisms able to metabolize them. However, alkane metabolism is in no way straightforward. These compounds are very hydrophobic and their solubility in water therefore very reduced, hindering their uptake. The compounds generated during alkane metabolism are often hydrophobic too, leading to their accumulation in the plasma membrane and alterations in the latter’s fluidity. Further, alkanes are chemically rather inert, and need to be activated (an energy-costly process) before they can be metabolized. Even so, many bacteria, filamentous fungi, and yeasts have acquired the ability to degrade alkanes and use them as a carbon source (van Beilen et al. 2003; Wentzel et al. 2007). A typical soil, sand, or ocean sediment contains 104–106 hydrocarbon degrading microorganisms per gram (Rosenberg 1993), and considerably more in oil-polluted sites (Harayama et al. 2004). Many alkane-degrading bacteria have a very versatile metabolism, and alkanes are but some of many substrates that can serve as carbon sources (Harayama et al. 2004; Margesin et al. 2003). In fact, alkanes are not usually the most preferred substrates; cells tend to use other compounds before turning to them. How this is achieved is analyzed in detail in the following chapter in this book by Moreno and Rojo. However, hydrocarbonoclastic bacteria are highly specialized in the degradation of hydrocarbons, and play a key role in their removal from polluted environments (Harayama et al. 2004; Head et al. 2006; Yakimov et al. 2007). A particularly well-studied example is that of Alcanivorax borkumensis, a marine bacterium that can assimilate linear and branched alkanes, but which is unable to metabolize the aromatic hydrocarbons, sugars, amino acids, or fatty acids, etc., commonly used as carbon sources (Schneiker et al. 2006; Yakimov et al. 1998). Alcanivorax spp. are present in low numbers in nonpolluted sea water, probably living on the alkanes continuously produced by algae and other marine organisms. After a spill of crude oil, however, Alcanivorax strains become predominant and are believed to play an important role in the natural bioremediation

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process (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 this respect (Coulon et al. 2007; McKew et al. 2007a, b). Alkanes can be metabolized aerobically or anaerobically. The present chapter deals only with aerobic degradation; anaerobic degradative pathways are covered in a separate chapter in this book series. Several reviews are available that focus on different features of the pathways and enzymes involved in alkane degradation (Coon 2005; van Beilen and Funhoff 2007; van Hamme et al. 2003; Wentzel et al. 2007; Rojo 2009; Wang and Shao 2013). The present chapter emphasizes recent developments in n-alkane metabolism in Eubacteria; although some Archaea can use alkanes and other hydrocarbons as carbon sources under aerobic conditions (Tapilatu et al. 2010; Fathepure 2014), the enzymes involved have not been studied in detail and are not covered here. The regulation of the expression of the genes involved in alkane degradation is treated in a separate chapter in this volume. The degradation of methane, which is oxidized via a specialized enzyme, is also covered in a separate chapter.

2

Uptake of n-Alkanes

2.1

The Low Water Solubility of Hydrocarbons Presents a Problem for their Uptake by Microbes

The water solubility of n-alkanes decreases exponentially as their molecular weight increases (Eastcott et al. 1988; see Table 1); the solubility of n-alkanes with more than nine carbon atoms is so low that their uptake by microorganisms is clearly compromised. This poses a problem for their biodegradation. The precise manner in which n-alkanes enter the cell is poorly understood, although the mechanisms involved probably differ depending on species, the molecular weight of the alkane, and the physicochemical characteristics of the environment (Wentzel et al. 2007). Table 1 Water solubility of representative n-alkanes (at 25  C) n-Alkane Propane Hexane Nonane Dodecane Hexadecane Eicosane Hexacosane

Carbon atoms 3 6 9 12 16 20 26

Data obtained from Eastcott et al. (1988)

Molecular weight 44.1 86.2 128.3 170.3 226.4 282.6 366.7

Solubility (mol L 1) 5  10 3 1.4  10 4 10 6 2  10 8 2  10 10 10 12 4  10 16

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Only low molecular weight n-alkanes are sufficiently soluble to be taken up by diffusion from the water phase. Microorganisms gain access to mediumand long-chain n-alkanes in the form of small, pseudosolubilized hydrocarbon droplets, or by adhering to droplets much larger than themselves (a process used by cells able to develop a hydrophobic surface) (Beal and Betts 2000; Hua and Wang 2013, 2014). Most bacteria able to degrade n-alkanes secrete surfactants of different chemical nature that facilitate the emulsification of hydrocarbons (Hommel 1990; Ron and Rosenberg 2002). Biosurfactants are believed to increase the surface area that hydrophobic compounds can expose to the water phase, thereby improving 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 n-alkanes such as hexadecane (Beal and Betts 2000; Noordman and Janssen 2002). However, in soils and elsewhere, the usefulness of surfactants in the uptake of n-alkanes is less evident (Holden et al. 2002). Pseudomonas aeruginosa produces rhamnolipid surfactants that stimulate the uptake of hexadecane via a process that requires energy (Beal and Betts 2000; Noordman and Janssen 2002). Rhamnolipids can disperse hydrocarbons in liquid cultures to generate hexadecane droplets in the sub-micromolar range, which clearly increases the availability of the hydrocarbon to the bacterium (Cameotra and Singh 2009). Efficient emulsification requires the production of relatively large amounts of the surfactant, which in turn requires high population densities of surfactant-producing microorganisms. This suggests that the role of surfactants at low cell densities might be different to emulsification (Ron and Rosenberg 2001). Indeed, surfactants facilitate adhesion to and detachment from surfaces or 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 n-alkane-degrading bacteria that also behave as opportunistic pathogens, such as P. aeruginosa, these properties of biosurfactants might facilitate infection; they might therefore also be considered virulence factors (Zulianello et al. 2006). The uptake of hydrocarbons is thus just one of the processes in which the properties of surfactants may be useful.

2.2

Transport of Hydrocarbons Through the Cell Envelope

After contact is established between cells and hydrocarbon molecules, the latter need to gain access to the plasma membrane where they will be processed by enzymes that initiate their oxidation (see Sects. 3 and 4). Three mechanisms might be involved: (a) passive diffusion of the hydrocarbons through the cell envelope, (b) passive uptake facilitated by protein transporters, or (c) energy-dependent active transport (reviewed by Hua and Wang 2014). In Gram-negative bacteria, the membrane is a strong barrier to both hydrophilic and hydrophobic molecules, and contains many proteins that facilitate the import and export of different molecules across it. These can be general porins, substrate-specific transporters, or active energy-dependent transporters (reviewed in Nikaido 2003 and van den Berg 2005). General porins do

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not bind their substrate, but rather allow the passive passage of small solutes by diffusion along their concentration gradients. Substrate-specific transporters have low affinity binding sites for particular compounds, and allow the entry of molecules when their concentration gradient across the membrane is shallow. Finally, active energy-dependent transporters bind substrates with high affinity and selectivity, and transport them against a concentration gradient using energy provided by the inner membrane protein TonB. Transporters for hydrophilic molecules have been extensively studied, but much less information is available on those dealing with hydrophobic molecules such as hydrocarbons. The best known example is that of the FadL protein from Escherichia coli, an outer membrane protein that allows the import of long-chain fatty acids via diffusion facilitated by spontaneous conformational changes that require no exogenous energy input (van den Berg 2005). FadL also mediates the uptake of n-alkanes with eight or ten carbon atoms (C8 and C10, respectively, where the subindex indicates the number of carbon atoms of the alkane molecule), which is severely reduced if the fadL gene is inactivated (Call et al. 2016). FadL homologs are widespread in Gram-negative bacteria. One such homolog is the AlkL protein involved in the alkane degradation pathway and encoded within the OCT plasmid of Pseudomonas putida GPo1. AlkL has been shown to greatly improve the import of fatty acid methyl esters and C7–C16 n-alkanes in both this bacterium and E. coli (Julsing et al. 2012; Grant et al. 2014). In the latter species, AlkL was found necessary for the uptake of C12–C16 n-alkanes, but not for n-octane, probably because it can be transported by FadL (Grant et al. 2014). In the hydrocarbonoclastic bacterium Alcanivorax dieselolei B5, three FadL-like outer membrane proteins named OmpT-1, OmpT-2, and OmpT-3 are required for the uptake of n-alkanes (Wang and Shao 2014). Compared to cultivation with acetate, the transcription of ompT-1 was strongly induced in cells growing on C24–C34 nalkanes or pristane, while the use of C8–C16 n-alkanes as the carbon source led to significantly reduced induction. Nevertheless, the expression of ompT-2 and ompT-3 was efficiently induced by all these hydrocarbons. Inactivation of the ompT-1 gene impaired growth on C28, C32, C36 n-alkanes or pristane, although C8–C24 n-alkanes could still be efficiently used. In contrast, the inactivation of ompT-3 impaired growth on C8–C12 n-alkanes, but did not affect the assimilation of C16, C24 n-alkanes and pristane. A mutant strain lacking OmpT-2 did not grow on C16–C24 n-alkanes, and showed slow growth on n-alkanes with under 16 and more than 24 carbon atoms. Transport studies confirmed these proteins to be involved in the selective uptake of these hydrocarbons. The above authors concluded that OmpT-1 is used for the uptake of n-alkanes of over 28 carbon atoms and pristane, that OmpT-2 is preferentially used for C16–C24 n-alkanes, and that OmpT-3 takes care of the uptake of C8–C12 n-alkanes. Finally, transport studies on the n-octadecane-degrading strain Pseudomonas sp. DG17, using 14C n-octadecane as a substrate, showed that, when the alkane concentration was higher than 4.5 μmol/L, its uptake was driven by a facilitated passive mechanism that required no supply of external energy. It was also insensitive to chemicals such as carbonyl cyanide m-chlorophenyl hydrazone (CCCP), which

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uncouples the proton gradient of the membrane and ultimately inhibits ATP synthesis. However, when the concentration of octadecane was about ten times lower, transport was strongly inhibited by CCCP (Hua et al. 2013). This suggests that the facilitated diffusion of n-alkanes by FadL-like proteins is probably enough to allow growth at the expense of these compounds when they are present at micromolar concentrations. At nanomolar concentrations, however, energy-driven active transport systems become necessary, but these have not yet been identified.

2.3

Chemotaxis

Many motile bacteria have systems for detecting and responding to the presence of specific chemicals in their environment, swimming towards or away from them in a process called chemotaxis (reviewed in Wadhams and Armitage 2004; Hazelbauer et al. 2008). Depending on whether the flagellum rotates in an anticlockwise or clockwise direction, cells swim in a straight line or stop and tumble. Cells sense chemicals via dedicated transmembrane receptors known as methylaccepting chemotaxis proteins (MCPs). Studies in E. coli have shown that, in response to reduced binding of the attractant to the MCPs, CheA histidine kinase autophosphorylates and transfers the phosphoryl group to the CheY and CheB response regulators. Phosphorylated CheY interacts with components of the flagellar motor, promoting a change in the rotational direction of the flagellum from anticlockwise to clockwise, causing cell tumbling. Phosphorylated CheB is a methylesterase that demethylates the MCP receptors, reducing their ability to induce CheA autophosphorylation, thereby resetting the system. Binding of the specific ligands to the MCPs inhibits the autophosphorylation of CheY and, therefore, reduces the frequency of motor switching, allowing swimming towards the attractant. This basic plan seems to be conserved across bacteria, although some chemotaxis pathways are more complex. Many chemicals eliciting an attractive response can be used as a carbon and energy source. Chemotaxis can therefore help bacteria find food supplies and, for low water-solubility chemicals, improve the rate of substrate acquisition. Several hydrocarbons elicit chemotaxic responses, although most of the literature refers to aromatic hydrocarbons such as naphthalene or toluene (Parales and Harwood 2002; Shingler 2003). Some reports indicate that n-alkanes also induce chemotaxis (Lanfranconi et al. 2003; Wang and Shao 2014). A recent study in Alcanivorax dieselolei B5 showed that the sensing of n-alkanes is connected to their uptake and metabolism by a signal transduction network (Wang and Shao 2014). In brief, mutagenesis studies identified an MCP protein and two additional components of the chemotaxis machinery, the inactivation of which abolished chemotaxis towards n-alkanes and significantly reduced growth on C8–C32 n-alkanes and pristane. When purified, the identified MCP protein can bind all these hydrocarbons, but not acetate, which is also a carbon source for this bacterium. Several genes belonging to the chemotaxis machinery, including the mentioned MCP receptor, were expressed more strongly when cells were grown on n-alkanes than when grown on

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acetate. An outer membrane protein named OmpS was also identified and shown to be essential for the assimilation of C8–C32 n-alkanes and pristane. The above authors proposed that a receptor for these hydrocarbons triggers the transport/ assimilation process via a mechanism still not understood. Inactivation of the gene coding for OmpT-1 porin, which transports C28, C32, C36 n-alkanes and pristane, abolished chemotaxis towards these hydrocarbons. Similarly, the absence of OmpT-2 or OmpT-3 abolished chemotaxis towards the n-alkanes they transport. It therefore seems that the sensing, chemotaxis, uptake, and assimilation of n-alkanes are interdependent processes in A. dieselolei, and probably in other Alcanivorax strains as well. In other bacterial species, chemotaxis can be dependent or independent of the assimilation of the attractant, depending on the compound and the strain in question (Pandey and Jain 2002; Sarand et al. 2008; Luu et al. 2015).

3

n-Alkane Degradation Pathways

In Eubacteria, the aerobic degradation of n-alkanes usually 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 β-oxidation to generate acetyl-CoA (van Hamme et al. 2003; Watkinson and Morgan 1990; Wentzel et al.

Fig. 1 The most common pathways for the degradation of n-alkanes by terminal and subterminal oxidation

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2007; Rojo 2009). Subterminal oxidation has been reported as well in some microorganisms (Fig. 1; Britton 1984; Kotani et al. 2003, 2006; Whyte et al. 1998). While the oxidation of fatty alcohols and fatty acids is widespread among microorganisms, the activation of the alkane molecule requires enzyme systems that are much less common.

4

Hydroxylation of n-Alkanes

In bacteria, the initial terminal hydroxylation of n-alkanes can be performed by enzymes belonging to different families (Table 2; van Beilen and Funhoff 2007; van Beilen et al. 2003). Microorganisms that degrade short-chain n-alkanes (C2–C4) have enzymes related to methane monooxygenases, while those that degrade medium- (C5–C11) or long-chain (>C12) n-alkanes commonly contain membrane, non-heme iron monooxygenases related to the well-characterized Pseudomonas putida GPo1 AlkB alkane hydroxylase. However, some strains contain alkane hydroxylating enzymes that belong to a family of soluble P-450 cytochromes and that are active against C5–C11 n-alkanes. Finally, several strains that assimilate n-alkanes of more than 18 carbon atoms contain alkane hydroxylases that seem to be unrelated to any of those mentioned above. Several yeasts assimilate n-alkanes as

Table 2 Enzyme classes oxidizing n-alkanes Enzyme class PRM, propane monooxygenase sBMO, butane monooxygenase pBMO, butane monooxygenase CYP153 CYP116B5 CYP52 AlkB-related AlmA LadA Dioxygenase PT7_2466 monooxygenase

Characteristics Non-heme iron monooxygenase similar to sMMO Non-heme iron monooxygenase similar to sMMO Copper-containing monooxygenase similar to pMMO Soluble cytochrome P450 (class I) Self-sufficient cytochrome P450 (class VII) Membrane-bound cytochrome P450 Non-heme iron monooxygenase Flavin-binding monooxygenase Thermophilic flavin-dependent monooxygenase Copper flavin-dependent dioxygenase Rieske-type monooxygenase

Substrate length C3

Host Bacteria

C2–C9

Bacteria

C2–C9

Bacteria

C5–C12 C14–C16

Bacteria Bacteria

C10–C16 C3–C13 or C10–C20 C20–C36 C10–C30

Yeasts Bacteria

C10–C30 C5–C24

Bacteria Bacteria

Bacteria Bacteria

The substrate range is approximate; upper and lower limits may vary in different strains. See text for details

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well. In those studied, the enzymes involved in the initial oxidation of the alkane molecule belong to the microsomal cytochrome P450 family (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 thought, 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 n-alkanes, but not on methane (Ashraf et al. 1994). The enzymes that initially oxidize these n-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 n-alkanes in the C1–C4 range, while some methanotrophs additionally produce a soluble form (sMMO) that is active against a wider range of substrates, oxidizing C1–C7 n-alkanes to the corresponding alcohols (Green and Dalton 1989). Thauera butanivorans, previously known as Pseudomonas butanovora (Anzai et al. 2000; Dubbels et al. 2009), can grow on C2–C9 n-alkanes using a pathway that sequentially oxidizes the terminal methyl group of the hydrocarbon (Arp 1999). The first enzyme in the pathway, butane monooxygenase (BMO), is a non-heme iron monooxygenase (similar to sMMO) that hydroxylates C2–C9 n-alkanes (Sluis et al. 2002). This enzyme has three components: a dinuclear iron-containing monooxygenase (BMOH) that in turn contains three different polypeptides, an NADHoxidoreductase (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 may require the assistance of a chaperonin-like protein, BmoG (Kurth et al. 2008). Gordonia sp. TY-5, which can grow on propane, contains an enzyme with sequence similarity to sMMO, but which has a very narrow substrate range: it can only oxidize propane and does so at the subterminal 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 coding for this propane monooxygenase have also been found in two propane-utilizing species, Mycobacterium sp. TY-6 and Pseudonocardia sp. TY-7 (Kotani et al. 2006). In the former, propane is oxidized at the terminal position, while in the latter both terminal and subterminal oxidation occurs. The BMOs of two other strains, Mycobacterium vaccae JOB5 and Nocardioides sp. CF8, have also been analyzed. That of M. vaccae JOB5 shows properties similar to sMMO (Hamamura et al. 1999), while that of Nocardioides sp. CF8 is a copper-containing enzyme formed by three subunits that share distant but significant similarity to other members of the pMMO family (Hamamura and Arp 2000; Hamamura et al. 1999; SayavedraSoto et al. 2011).

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The AlkB Family of Alkane Hydroxylases

The most extensively characterized alkane degradation pathway is that encoded within the OCT plasmid of P. putida GPo1 (formerly Pseudomonas oleovorans GPo1). It was originally characterized by Coon and colleagues (Baptist et al. 1963) and has become a model system (van Beilen et al. 1994, 2001). The first enzyme of this pathway is an integral-membrane non-heme di-iron monooxygenase, named AlkB, that hydroxylates n-alkanes at the terminal position. This requires the assistance of two soluble electron transfer proteins named rubredoxin (AlkG) and rubredoxin reductase (AlkT). Rubredoxin reductase, via its cofactor FAD, transfers electrons from NADH to rubredoxin, which in turn transfers the electrons to AlkB (see Fig. 2). The biochemical properties of AlkB have been analyzed in detail. Genetic studies have shown it to have six transmembrane segments and a catalytic site that faces the cytoplasm. The active site includes four histidine-containing sequence motifs that are conserved in other hydrocarbon monooxygenases and that chelate two iron atoms (Fig. 3; Shanklin et al. 1994; van Beilen et al. 1992b). The di-iron 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 is reduced to H2O by electrons provided by rubredoxin. Oxidation is regio- and stereospecific (van Beilen et al. 1996). The P. putida GPo1 AlkB alkane hydroxylase oxidizes propane, n-butane (Johnson and Hyman 2006), and C5–C13 n-alkanes (van Beilen et al. 2005b). All these nalkanes support growth. Methane, ethane, and n-alkanes longer than C13 are not oxidized. Mutagenesis studies allowed the identification of a residue, Trp55, which appears to limit the size of the alkane molecules that AlkB can oxidize, since, when replaced by Ser or Cys, the substrate range increases to include C14 and C16

Fig. 2 Oxidation of n-alkanes by alkane hydroxylases of the AlkB family (left) and the bacterial cytochrome P450 family (right). AH, membrane-bound alkane hydroxylase; Rub rubredoxin, RubR rubredoxin reductase, Cyt P450 soluble cytochrome P450, Fdx ferredoxin, FdxR ferredoxin reductase. The gray bar represents the plasma membrane; the phospholipid layer facing the cytoplasm is marked as “In”

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Fig. 3 Proposed structure of the P. putida GPo1 membrane-bound AlkB alkane hydroxylase. The gray bar represents the plasma membrane. The four histidine clusters (H) believed to bind the two iron atoms at the catalytic site are indicated, as is the proposed position of residue Trp55 (W55), which extends its bulky side group towards the hydrophobic pocket in which the alkane molecule is believed to fit (Adapted from van Beilen et al. (2005b) and Rojo (2005))

n-alkanes (van Beilen et al. 2005b). It has been proposed that the AlkB active site might be a deep hydrophobic pocket formed by the alignment of the six transmembrane helices, and that the alkane molecule slides 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 protrudes into the hydrophobic pocket, impeding n-alkanes longer than C13 from entering any deeper into the pocket, and thus 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 n-alkanes to fit into the hydrophobic pocket. Recent genetic and modeling studies support this idea (Alonso et al. 2014). More than 400 AlkB homologs are known (Marín et al. 2001, 2003; Smits et al. 1999, 2002, 2003; van Beilen et al. 2002b, 2004; Nie et al. 2014a). They have been found in both Gram-positive and Gram-negative microorganisms and show wide sequence diversity (van Beilen et al. 2003; Lo Piccolo et al. 2001; Nie et al. 2014a). Interestingly, only a few of these AlkB enzymes oxidize C5–C13 n-alkanes, as does P. putida GPo1 AlkB; most members of this family prefer n-alkanes longer than C10. The rubredoxin that transfers electrons to the AlkB active site is a small redoxactive iron-sulfur protein. The AlkG rubredoxin of P. putida GPo1 is unusual in that it contains two rubredoxin domains, AlkG1 and AlkG2, connected by a linker;

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rubredoxins from other microorganisms have only one of these domains. Several rubredoxins present in Gram-positive and Gram-negative alkane-degrading bacteria were cloned and analyzed in complementation assays for their ability to substitute P. putida GPo1 AlkG. Interestingly, they clustered into two groups. AlkG1-type rubredoxins could not transfer electrons to the alkane hydroxylase, while AlkG2type enzymes were able to do so and could therefore act as substitutes (van Beilen et al. 2002a). In some cases, the AlkB-type hydroxylase and the rubredoxin fuse into a single polypeptide, the rubredoxin domain being needed for the activity of the hydroxylase domain (Hamamura et al. 2001; Bihari et al. 2011; Nie et al. 2011; Nie et al. 2014a). In addition, fusion proteins containing an N-terminal ferredoxin domain, a central ferredoxin reductase domain, and a C-terminal alkane-hydroxylase domain have been identified during DNA sequence searches of genomic and metagenomic databanks (Nie et al. 2014a). AlkG1-type rubredoxins have other roles as well; in fact, rubredoxin-rubredoxin reductase systems are present in many other organisms that are unable to degrade n-alkanes. 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). The structure of the rubredoxin-rubredoxin reductase complex, which has been resolved in P. aeruginosa, seems to be optimized for the rapid transport of reducing equivalents to the final receptor (Hagelueken et al. 2007).

4.3

Cytochrome P450 Alkane Hydroxylases

The cytochromes P450 are heme-thiolate proteins that catalyze the oxygenation of many compounds. Found across all the kingdoms of life, they can be grouped into 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 n-alkanes contain alkane hydroxylases that belong to a distinct family of bacterial soluble cytochrome P450 monooxygenases. The first to be characterized from a biochemical and genetic perspective was CYP153A1 from Acinetobacter sp. EB104 (Maier et al. 2001), but similar enzymes have been found in strains of Mycobacterium, Rhodococcus, and Dietzia, and in several Gram-negative bacteria including hydrocarbonoclastic bacteria such as Alcanivorax spp. (Sekine et al. 2006; van Beilen et al. 2005a, 2006; Schneiker et al. 2006; Wang et al. 2010; Scheps et al. 2011; Nie et al. 2014b). These P450 cytochromes require the presence of ferredoxin and of ferredoxin reductase to transfer electrons from NAD(P)H to the cytochrome (Fig. 2). Complementation assays have shown many of these P450 proteins can functionally substitute for P. putida GPo1 AlkB, revealing them to be true alkane hydroxylases (van Beilen et al. 2006). Cytochrome P450 from Mycobacterium sp. HXN-1500 has been purified and shown to hydroxylate C6–C11 n-alkanes to 1-alkanols with high affinity and regioselectivity (Funhoff et al. 2006). Fusion proteins containing an N-terminal cytochrome P450 domain, a ferredoxin reductase domain, and C-terminal ferredoxin

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domain have been detected in genomic and metagenomic databanks (Nie et al. 2014a). In addition, a class VII cytochrome P450 was identified in Acinetobacter redioresistens S13 with an N-terminal heme domain and a C-terminal reductase domain comprising NADPH-, FMN-, and [2Fe-2S]-binding sites (Minerdi et al. 2015). This cytochrome is therefore catalytically self-sufficient and does not require ferredoxin reductase. Its heterologous expression in E. coli showed it to hydroxylate C14 and C16 n-alkanes. As mentioned above, several yeasts can assimilate n-alkanes. 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). These receive electrons from NADPH via FAD- and FMN-containing reductases. A detailed description of the yeast enzymes involved in the oxidation of n-alkanes is provided by R. Fukuda in a separate chapter in this book.

4.4

Alkane Hydroxylases for Long-Chain n-Alkanes

Several bacterial strains have been reported to assimilate n-alkanes larger than C20 (for a compilation see Wentzel et al. 2007). In some cases, the enzymes responsible for the oxidation of such n-alkanes, which are solid at room temperature, have been characterized. In Acinetobacter sp. M1, which can grow on C13–C44 n-alkanes, several alkaneoxidizing enzymes have been detected. Two of these, 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+, and does not receive electrons from NADH. It is therefore clearly unrelated to the AlkB family of hydroxylases (Maeng et al. 1996) and has been proposed to be a dioxygenase that oxidizes C10–C30 n-alkanes generating n-alkyl hydroperoxides that render the corresponding aldehyde. Acinetobacter sp. DSM 17874 also contains at least three n-alkane-oxidizing enzymes. Two are AlkB paralogs similar to the AlkMa and AlkMb enzymes described above, and oxidize C10–C20 n-alkanes (Throne-Holst et al. 2006). The third, a flavin-binding monooxygenase named AlmA, oxidizes C20 to >C32 n-alkanes (Throne-Holst et al. 2007). Genes homologous to almA have been identified in several other long-chain n-alkane degrading strains, including Acinetobacter sp. M1 and several Alcanivorax species (Liu et al. 2011; Wang and Shao 2012a, b). A different long-chain alkane hydroxylase has been characterized in the hemophilic bacterium Geobacillus thermodenitrificans NG80–2 (Feng et al. 2007). Termed LadA, it oxidizes C15–C36 n-alkanes, generating the corresponding primary alcohols. Its crystal structure has been resolved, showing it to belong to the bacterial luciferase family of proteins, which are two-component flavin-dependent oxygenases (Li et al. 2008). LadA is believed to oxidize n-alkanes via a mechanism similar to that of other flavoprotein monooxygenases; its ability to recognize and hydroxylate long-chain n-alkanes probably lies in the way it captures these molecules.

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Finally, an alkane monooxygenase different to any mentioned above was recently described in Pusillimonas sp. T7–7, a Gram-negative, cold-tolerant bacterium that can assimilate C5–C30 n-alkanes. The protein was purified and shown to belong to the Rieske non-heme iron monooxygenase family (Li et al. 2013). It contains two different subunits, and works in association with a ferredoxin and an NADHdependent reductase. The purified enzyme was shown to hydroxylate C5–C24 n-alkanes. Several bacterial strains can degrade >C20 n-alkanes using enzyme systems that are yet to be characterized. It is likely that new enzyme classes responsible for the oxidation of these high molecular weight n-alkanes will be found in the near future.

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 well-characterized alkane degrader P. putida GPo1. However, it is rather common to find strains that contain more than one alkane oxidation system. In many cases these alkane oxidation enzymes have different substrate ranges or different induction patterns. The Alcanivorax species characterized to date typically contain two AlkB-related alkane hydroxylases, one to three cytochrome P450s involved in alkane degradation, and an alkane hydroxylase similar to AlmA (Hara et al. 2004; Sabirova et al. 2006; Schneiker et al. 2006; van Beilen et al. 2004; Liu et al. 2011; Wang and Shao 2012a, b). The presence of multiple alkane oxidation determinants in a single strain occurs both in hydrocarbonoclastic bacteria and in bacterial species that have a versatile metabolism. For example, P. aeruginosa PAO1 and RR1 contain two AlkB-related alkane hydroxylases that are differentially regulated (Marín et al. 2001; Stover et al. 2000), while genomic and proteomic analyses of P. aeruginosa SJTD-1 have identified two AlkB-like monooxygenases, two cytochrome P450s of the CYP153 family, and one AlmA-like monooxygenase (Liu et al. 2014, 2015). 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 n-alkanes, and a third that oxidizes C32–C36 n-alkanes (Throne-Holst et al. 2007). Besides carrying two AlkB-related hydroxylases, Acinetobacter sp. M1 also contains a dioxygenase that oxidizes long-chain n-alkanes (Maeng et al. 1996; Tani et al. 2001), and has 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 n-alkanes of different size ranges (Hamamura et al. 2001; Kotani et al. 2006). Dietzia sp. DQ12–45-1b has an AlkB-like alkane hydroxylase and a cytochrome P450 of the CYP153 family; the former is responsible for the hydroxylation of n-alkanes longer than C14, while the latter deals with those shorter than C10 (Nie et al. 2014b). Rhodococcus sp. Q15 and NRRL B-16531 contain at least four AlkB-related alkane hydroxylases (Whyte et al. 2002); in the latter strain, two additional cytochrome P450s of the CYP153 family have also been detected

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(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 very divergent alkane degradation genes in a single bacterial strain suggests that horizontal transfer has greatly facilitated their spread. 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). The four AlkB-related proteins present in Rhodococcus sp. Q15 and NRRL B-16531 are as divergent as all hydroxylases analyzed from Gram-positive strains (Whyte et al. 2002). A similar conclusion was reached in a more recent analysis comparing a set of 458 AlkB-type alkane hydroxylases from 369 genomes belonging to 51 bacterial genera: the topology of a phylogenetic tree based on the AlkB sequences did not match that of the 16S rRNA genes (Nie et al. 2014a). Some alkane degradation genes have been found on transposons (van Beilen et al. 2001) and 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 n-Alkanes

The primary fatty alcohols generated by the terminal oxidation of n-alkanes are further oxidized to aldehydes by alcohol dehydrogenase (ADH). There are several kinds of ADH. Some use NAD(P)+ as an electron acceptor, while others use cytochromes or ubiquinone. Most NAD(P)+-independent ADHs contain pyrroloquinoline quinone (PQQ) as a prosthetic group, and are commonly named quinoprotein ADHs. Many bacteria contain several ADHs that can be used for the assimilation of distinct alcohols. For example, T. butanivorans 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. BDH contains PQQ and heme c as prosthetic 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 against C2–C8 primary alcohols and C3–C8 secondary alcohols (Vangnai et al. 2002). Growing cells in butane leads to the 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 the assimilation of butane, although the simultaneous inactivation of both genes renders cells unable to grow on this substrate (Vangnai et al. 2002). When T. butanivorans was grown on 2-butanol and lactate, two additional NAD+-dependent secondary ADHs were

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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, but their precise roles have not been reported (Vangnai et al. 2002). It is worth noting that BOH and BDH are active against aldehydes. Acinetobacter calcoaceticus HO1-N contains at least two ADHs. One of them requires NAD+ and shows a preference for decanol. The other requires NADP+ and shows greater activity against tetradecanol. An aldehyde dehydrogenase active against 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 P. putida GPo1 OCT plasmid. The alcohol dehydrogenase AlkJ is necessary for growth on n-alkanes only if the chromosomal AlcA alcohol dehydrogenase is inactivated by mutation (van Beilen et al. 1992a). This again indicates a redundancy in these enzymes. Similarly, the plasmid-encoded AlkH aldehyde dehydrogenase is not essential for growth on n-alkanes, which agrees with the presence of several aldehyde dehydrogenases in the P. putida GPo1 chromosome (van Beilen et al. 1994). The secondary alcohols generated by subterminal oxidation of n-alkanes are turned into ketones by alcohol dehydrogenases (Fig. 1). Gordonia sp. TY-5, a bacterium that can grow on propane and C13–C22 n-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 these three secondary ADHs, which are all expressed in propane-grown cells, ADH1 seemed to play the major role under the conditions in the latter report. 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 aldehyde oxidation are further metabolized by β-oxidation, generating Acyl-CoA, which enters the tricarboxylic acid cycle. However, when the carbon source is in excess relative to nitrogen, many bacteria use 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 granules (Alvarez and Steinbuchel 2002; Grage et al. 2009; Waltermann et al. 2005). These compounds can then serve as endogenous carbon and energy sources during starvation periods. The formation of storage lipids is common among hydrocarbon-utilizing marine bacteria. Alcanivorax strains, for example, accumulate triacylglycerols and wax esters when growing on pyruvate or n-alkanes (Kalscheuer et al. 2007). In addition, P. putida GPo1, a soil bacterium, forms intracellular inclusions of poly-β-hydroxyoctanoate when grown on n-octane (de Smet et al. 1983), and Acinetobacter sp. M-1 forms wax esters when growing on hexadecane (Ishige et al. 2000, 2002).

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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), and even much more complex compounds such as pristane (reviewed in Britton 1984; Watkinson and Morgan 1990). Alcanivorax spp. 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 terminal and a di-terminal oxidation of the hydrocarbon molecule, rendering mono- and dicarboxylic acids (Watkinson and Morgan 1990; Nhi-Cong et al. 2010). However, a subterminal oxidation pathway for pristane has also been proposed (Nhi-Cong et al. 2009; Yang et al. 2014).

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. Alcohols derived from n-alkanes are valuable products in the pharmaceutical, cosmetics, and food industries. Alkane hydroxylases frequently oxidize not only their natural substrates but other compounds as well, albeit with reduced efficiency, further increasing their potential usefulness in industry (van Beilen and Funhoff 2005). P. putida GPo1 AlkB can, for example, generate epoxides from alkenes and other chemicals with a terminal double bond, oxidize alcohols to aldehydes, and catalyze 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 stereospecific 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 in high enantiomeric excess. Optically active epoxides can be used to generate a number of chemicals that are useful precursors from which to derive several products of added value. The set-up of a cost-effective high-scale process based on alkane hydroxylases is complicated, however, due to practical issues such as substrate uptake, the toxicity of the substrate and/or the product generated, uncoupling, oxygen mass transfer, low turnover with some compounds, regeneration of redox cofactors, and problems related to product recovery (reviewed in Soussan et al. 2016). Considerable efforts have been made in the optimization of these processes. The use of alkane hydroxylases from thermophilic microorganisms may help solve some of these problems since high temperatures can increase the solubility of n-alkanes and reduce their viscosity.

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Research Needs

Despite extensive research on alkane degradation by bacteria, many features remain poorly understood, including how n-alkanes are incorporated or transported into the cell (which may differ between n-alkanes and microorganisms). The enzymes for the degradation of short- and medium-chain length n-alkanes are rather well characterized, although there is a paucity of structural data. However, some findings indicate that, in several microorganisms, C20–C50 n-alkanes are probably oxidized by enzymes yet to be identified. The question of why bacterial strains frequently contain several different or related alkane hydroxylases that have very similar substrate specificities is also intriguing. It may be that these hydroxylases differ in aspects that are still unknown but that are important in cell biology. Finally, the use of alkane hydroxylases for biotransformations of industrial interest – an area of great potential – still has to resolve several technical issues that limit efficiency. Acknowledgments Work in the author’s laboratory is funded by the Spanish Ministry of Economy and Competitiveness (grant BIO2015-66203-P) and the European Commission VII Framework Program (grant number 312139).

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Enzymes for Aerobic Degradation of Alkanes in Yeasts Ryouichi Fukuda and Akinori Ohta

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Uptake of n-Alkanes by Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Terminal Hydroxylation of n-Alkanes by Cytochrome P450 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Oxidation of Fatty Alcohols to Fatty Aldehydes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Oxidation of Fatty Aldehydes to Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Activation and Utilization of Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

A wide variety of yeasts can utilize n-alkanes as sole carbon and energy sources. The degradation pathways of n-alkanes in yeasts and the enzymes associated with these pathways have been studied intensively in the ascomycetous yeasts, Candida tropicalis, Candida maltosa, and Yarrowia lipolytica, for biotechnological applications, such as conversion of n-alkanes to proteins or useful compounds, as well as for elucidating the metabolism of hydrophobic substrates by fungi. Here, we describe the aerobic degradation pathway of n-alkanes in yeasts and the enzymes that catalyze the reactions involved in the degradation. In n-alkaneassimilating yeasts, incorporated n-alkanes are hydroxylated to fatty alcohols by cytochromes P450 of the CYP52 family in the endoplasmic reticulum (ER). Fatty

R. Fukuda (*) Department of Biotechnology, The University of Tokyo, Tokyo, Japan e-mail: [email protected] A. Ohta Department of Biological Chemistry, College of Bioscience and Biotechnology, Chubu University, Kasugai, Aichi, Japan e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_7

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alcohols are oxidized in the ER or the peroxisome to fatty aldehydes and finally to fatty acids, which are then activated to acyl-CoAs and metabolized by β-oxidation or used for lipid synthesis.

1

Introduction

n-Alkanes are common compounds in nature, and a variety of microorganisms, including bacteria, yeasts, and filamentous fungi, can utilize these compounds as their sole carbon and energy sources. The assimilation of n-hexadecane has been assessed in ~700 yeast species among the 1,270 yeasts listed in The Yeasts: A Taxonomic Study, and ~180 of these 700 species have been shown to have the ability to assimilate n-hexadecane (Kurtzman et al. 2011). These ~180 yeasts belong to 28 genera of ascomycetous and basidiomycetous yeasts including Candida, Debaryomyces, Metschnikowia, Yarrowia, and Cryptococcus (Fig. 1). Neither the Candida tropicalis Candida maltosa Candida parapsilosis 0.02 963 Lodderomyces elongisporus Candida dubliniensis 1000 Candida albicans Scheffersomyces stipitis Meyerozyma guilliermondii Debaryomyces hansenii Millerozyma farinosa Saccharomyces cerevisiae Yarrowia lipolytica Candida apicola Starmerella bombicola Metschnikowia pulcherrima Schizosaccharomyces pombe Cryptococcus_musci 987

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Fig. 1 Phylogenetic tree of n-alkane-assimilating yeasts. Phylogenetic tree of D1/D2 regions of 26S ribosomal DNA of the yeasts that can assimilate n-alkanes and the model yeasts, Saccharomyces cerevisiae and Schizosaccharomyces pombe, was constructed using ClustalW (DDBJ, v2.1) and drawn using NJplot. The scale bar indicates 0.02 substitutions per site. The bootstrap values by 1000 repetitions are indicated. The accession numbers of sequences of D1/D2 regions from GenBank are as follows: Candida albicans (U45776), Candida apicola (U45703), Candida dubliniensis (U57685), C. maltosa (U45745), Candida parapsilosis (U45754), Candida tropicalis (U45749), Cryptococcus musci (KC585415), Debaryomyces hansenii (U45808), Lodderomyces elongisporus (U45763), Metschnikowia pulcherrima (U45736), Meyerozyma guilliermondii (U45709), Millerozyma farinosa (U45739), Saccharomyces cerevisiae (U44806), Scheffersomyces stipitis (U45741), Schizosaccharomyces pombe (U40085), Starmerella bombicola (U45705), and Yarrowia lipolytica (U40080)

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n-Alkane

Plasma membrane

ER

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Peroxisome CH2OH

CH2OH

FADH

FAOD CHO

CHO

FALDH

FALDH

COOH

ACS Acyl-CoA

Lipids

COOH

ACS Acyl-CoA β-Oxidation

Acetyl-CoA Fig. 2 Proposed pathway of n-alkane metabolism in yeasts. Incorporated n-alkanes are hydroxylated to fatty alcohols by cytochromes P450 of the CYP52 family. Fatty alcohols are oxidized to fatty aldehydes by fatty alcohol dehydrogenase (FADH) in the ER or fatty alcohol oxidase (FAOD) in the peroxisome. Fatty aldehydes are oxidized to fatty acids by fatty aldehyde dehydrogenase (FALDH) in the ER or the peroxisome. Fatty acids are activated to acyl-CoAs by acyl-CoA synthetase (ACS) and are metabolized through β-oxidation pathway in the peroxisome or utilized for membrane or storage lipid synthesis

model yeast Saccharomyces cerevisiae nor Schizosaccharomyces pombe, however, can assimilate n-alkanes. The metabolic pathway of n-alkanes has been studied intensively in Candida tropicalis (Tanaka and Fukui 1989), Candida maltosa (Mauersberger et al. 1996), and Yarrowia lipolytica (Barth and Gaillardin 1996; Barth and Gaillardin 1997; Fickers et al. 2005; Fukuda 2013; Fukuda and Ohta 2013; Nicaud 2012) and has attracted considerable attentions owing to its involvement in the production of singlecell protein (SCP) and other useful materials, including long-chain dicarboxylic acids and tricarboxylic acid (TCA) cycle intermediates, from n-alkanes (Fickers et al. 2005; Tanaka and Fukui 1989). In order to establish and improve systems for the production of useful materials from n-alkanes using yeasts, the n-alkane metabolic pathway and associated enzymes must be understood in detail. In C. tropicalis, C. maltosa, and Y. lipolytica, incorporated n-alkanes are sequentially oxidized to fatty acids, which are then metabolized via the β-oxidation pathway in the peroxisome or utilized for the synthesis of membrane or storage lipids (Fig. 2). This chapter discusses the pathway of n-alkane oxidation to fatty acids and the subsequent activation of fatty acids to acyl-CoAs, as well as the enzymes catalyzing these reactions in yeasts. The metabolism and utilization of fatty acids in yeasts will be described in another chapter.

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Uptake of n-Alkanes by Yeasts

n-Alkanes exhibit poor solubility in water, and it has been proposed that yeasts facilitate the uptake of n-alkanes by two mechanisms, which are not mutually exclusive. In the first mechanism, reported for some species of n-alkane-assimilating yeasts, bioemulsifiers or biosurfactants are secreted by yeasts to solubilize n-alkanes (Barth and Gaillardin 1996; Tanaka and Fukui 1989): Y. lipolytica secretes a 28-kDa emulsifier, named liposan, composed of 83% carbohydrate and 17% protein (Cirigliano and Carman 1984, 1985), while Starmerella bombicola and Candida apicola produce the biosurfactant, sophorolipid (Van Bogaert et al. 2007). The physiological roles of these bioemulsifiers and biosurfactants in the uptake of n-alkanes, however, remain to be examined. In the second mechanism, n-alkaneassimilating yeasts adhere to n-alkane droplets to enhance the uptake of n-alkanes. Protrusions or slime-like outgrowths have been observed on the cell surface of C. tropicalis, C. maltosa, and Y. lipolytica cultured in the presence of n-alkanes (Kim et al. 2000; Mauersberger et al. 1996; Osumi et al. 1975; Tanaka and Fukui 1989). The slime-like outgrowths have been shown to reach the cell membrane through electron-dense channels in C. tropicalis, while the endoplasmic reticulum (ER) was found close to the cell membrane beneath such channels (Osumi et al. 1975). From these observations, it was suggested that n-alkanes are attached to the protrusions or slime-like outgrowths and are thereby transported through the channels to the ER, where they are hydroxylated to fatty alcohols (Sect. 3). The molecular structures of these protrusions or slime-like outgrowths and their roles in n-alkane assimilation, however, are not well understood. Two models have been proposed for the uptake of n-alkanes by yeasts: one is a passive, diffusion-like mechanism facilitated by the hydrophobic properties of nalkanes, while the other is an active, energy-dependent mechanism mediated by transporter proteins. Although the molecular mechanism of n-alkane uptake by yeasts is still unclear, the uptake of 14C-labeled n-hexadecane by Y. lipolytica was shown to be upregulated by the incubation with n-decane and inhibited by KCN and 2,4-dinitrophenol (Bassel and Mortimer 1985). Mutations in the 16 loci were also shown to significantly reduce n-hexadecane uptake in Y. lipolytica (Bassel and Mortimer 1985). These results support the model of n-alkane incorporation across the plasma membrane by one or more energy-dependent transporters. A Y. lipolytica strain with an insertion mutation in ABC1 encoding an ATP-binding cassette (ABC) transporter was shown to exhibit defective growth on n-hexadecane, but not on n-decane; however, the involvement of Abc1p in the uptake of n-hexadecane remains to be investigated (Thevenieau et al. 2007).

3

Terminal Hydroxylation of n-Alkanes by Cytochrome P450

Incorporated n-alkanes are transported from the plasma membrane to the ER, where they are hydroxylated to fatty alcohols by cytochromes P450 (P450s) belonging to the CYP52 family using molecular oxygen (Fig. 2) (Nelson 2009). Genes encoding

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CYP52-family P450s have been identified in various n-alkane-assimilating yeasts, including C. tropicalis (Sanglard et al. 1987; Seghezzi et al. 1992; Seghezzi et al. 1991), C. maltosa (Ohkuma et al. 1991, 1995), Candida albicans (Kim et al. 2007; Panwar et al. 2001), Candida dubliniensis, Candida parapsilosis, Debaryomyces hansenii (Yadav and Loper 1999), Lodderomyces elongisporus, Meyerozyma guilliermondii, Scheffersomyces stipitis, Starmerella bombicola (Van Bogaert et al. 2009), and Y. lipolytica (Fig. 3) (Fickers et al. 2005; Hirakawa et al. 2009; Iida et al. 1998, 2000). A striking feature of the CYP52-family P450s in these yeasts is that they exist as multiple paralogs of the CYP52-family P450 gene. Eight genes encoding CYP52-family P450s have been identified in C. tropicalis and C. maltosa, and Y. lipolytica has twelve CYP52-family P450 genes. Furthermore, C. albicans, C. dubliniensis, C. parapsilosis, D. hansenii, L. elongisporus, M. guilliermondii, and S. stipitis have more than four genes encoding CYP52family P450s. During the evolution of n-alkane-assimilating yeasts, the CYP52family P450 genes may have multiplicated and functionally diversified to efficiently metabolize n-alkanes and their metabolites or to detoxify them (see following paragraph). The CYP52-family P450s of C. maltosa are encoded by ALK1–ALK8, and quadruple deletion of ALK1, ALK2, ALK3, and ALK5 was shown to cause defects in the utilization of n-alkanes for growth (Ohkuma et al. 1998). In Y. lipolytica, the CYP52-family P450s are encoded by ALK1–ALK12, and a mutant in which all twelve ALK genes are deleted completely lost the ability to grow on n-alkanes (Takai et al. 2012). These results clearly indicate the essential roles of the CYP52family P450s in the assimilation of n-alkanes (Fig. 4). The Y. lipolytica deletion mutant of twelve ALK genes was furthermore shown not to be able to grow on ndecane even in the presence of glucose, suggesting that n-decane is toxic to the yeast cells and that the CYP52-family P450s are involved in the detoxification of n-decane (Takai et al. 2012). Substrate specificities have been studied in a subset of the CYP52-family P450s of C. tropicalis, C. maltosa, C. albicans, and Y. lipolytica, and it has been shown that they have distinct substrate preferences (Fig. 3) (Eschenfeldt et al. 2003; Iwama et al. 2016; Kim et al. 2007; Ohkuma et al. 1998; Zimmer et al. 1996). In accordance with their involvement in n-alkane metabolism, some CYP52-family P450s were shown to preferentially hydroxylate n-alkanes, while, in contrast, a subset of CYP52-family P450s preferred hydroxylation of the ω-terminal end (ω-hydroxylation) of fatty acids. Some CYP52-family P450s hydroxylated both n-alkanes and the ω-terminal end of fatty acids. The amino acid sequences of Alk4 (CYP52A7) and Alk5 (CYP52A8) of C. tropicalis (Seghezzi et al. 1992); Alk5 (CYP52A9), Alk7 (CYP52A10), and Alk8 (CYP52A11) of C. maltosa (Zimmer et al. 1996, 1998); and CYP52A21 of C. albicans (Kim et al. 2007) – all of which were shown to preferentially hydroxylate the ω-terminal ends of fatty acids – showed significant sequence similarities (Fig. 3). Interestingly, it has been reported that CYP52A3 of C. maltosa catalyzes the oxidation of fatty alcohol and fatty aldehyde in addition to hydroxylating n-alkane and the ω-terminus of fatty acids (Scheller et al. 1998).

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CYP52A1 (Ct Alk1) CYP52A4 (Cm Alk3) CYP52A22 (Ca) CYP52A2 (Ct Alk2) CYP52A23 (Ca) CYP52A5 (Cm Alk2) CYP52A6 (Ct Alk3) CYP52A24 (Ca) CYP52A3 (Cm Alk1) CYP52A44 (Dh) CYP52A53 (Ss) CYP52A54 (Ss) CYP52A56 (Ss) CYP52A43 (Dh) CYP52A57 (Ss) CYP52A55 (Ss) CYP52A45 (Dh) CYP52A47 (Dh) CYP52A46 (Dh) CYP52A7 (Ct Alk4) CYP52A8 (Ct Alk5) CYP52A9 (Cm Alk5) CYP52A21 (Ca) CYP52A11 (Cm Alk8) CYP52A10 (Cm Alk7) CYP52D2 (Ct) CYP52D1 (Cm Alk4) CYP52F1 (Yl Alk1) CYP52F10 (Yl Alk9) CYP52F2 (Yl Alk2) CYP52F11 (Yl Alk10) CYP52F3 (Yl Alk3) CYP52F9 (Yl Alk12) CYP52F4 (Yl Alk4) CYP52F5 (Yl Alk5) CYP52F7 (Yl Alk7) CYP52F6 (Yl Alk6) CYP52F8 (Yl Alk8) CYP52C1 (Ct Alk7) CYP52C3 (Ca) CYP52C2 (Cm Alk6) CYP52B1 (Ct Alk6) CYP52S1 (Yl Alk11) CYP52E3 (Sb) CYP52M1 (Sb) CYP52N1 (Sb) CYP51 (Yl)

AF A AF A A

F F F F F F AF AF AF A AF F F F AF F

F F

Fig. 3 Phylogenetic tree of the CYP52-family P450s in yeasts. Phylogenetic tree of the CYP52family P450s of n-alkane-assimilating yeasts was constructed using ClustalW (DDBJ, v2.1) and drawn using NJplot. CYP51 of Y. lipolytica was used as an out-group. The scale bar indicates 0.05 substitutions per site. The bootstrap values by 1000 repetitions are indicated. The accession numbers of sequences from UniProtKB are as follows: CYP52A21 (Q59K96), CYP52A22 (Q5AAH7), CYP52A23 (Q5AAH6), CYP52A24 (Q5A8M1), CYP52C3 (Q5AGW4), CYP52A3 (P16496), CYP52A5 (Q12581), CYP52A4 (P16141), CYP52A9 (Q12586), CYP52A10 (Q12588), CYP52A11 (Q12589), CYP52C2 (Q12587), CYP52D1 (Q12585), CYP52A1 (P10615), CYP52A2 (P30607), CYP52A6 (P30608), CYP52A7 (P30609), CYP52A8 (P30610), CYP52B1 (P30611), CYP52C1 (P30612), CYP52D2 (Q874J0), CYP52A43 (Q6BVP2), CYP52A44 (Q6BVH7), CYP52A45 (Q6BNW0), CYP52A46 (Q6BNV9), CYP52A47 (Q6BNV8), CYP52A53 (A3LRT5), CYP52A54 (A3LR60), CYP52A55 (A3LS01), CYP52A56 (A3LZV9), CYP52A57 (A3LSP0), CYP52E3 (B8QHP3), CYP52M1 (B8QHP1), CYP52N1 (B8QHP5), CYP52F1 (O74127), CYP52F2 (O74128), CYP52F3 (O74129), CYP52F4 (O74130), CYP52F5 (O74131),

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In Y. lipolytica, eleven CYP52-family P450s belong to the CYP52F subfamily and one belongs to the CYP52S subfamily. The CYP52F-subfamily P450s, Alk1p–Alk10p and Alk12p, appear to constitute a monophyletic clade in the phylogenetic tree of the CYP52-family P450s (Fig. 3). The CYP52F-subfamily P450s of Y. lipolytica can also be classified into four groups: P450s with significant n-alkanehydroxylating activity, P450s with significant hydroxylating activity for the ω-terminus of dodecanoic acid, P450s with significant hydroxylating activity for both n-alkanes and dodecanoic acid, and P450s with faint or no oxidizing activity for these substrates. Alk1p, Alk9p, Alk2p, and Alk10p, which have been shown to exhibit substrate preferences for n-alkanes, share significant sequence similarities, while Alk5p and Alk7p, which have ω-hydroxylation activities to dodecanoic acid, are structurally similar (Iwama et al. 2016). Alk proteins that were shown to catalyze the oxidation of n-alkanes showed distinct preferences for different n-alkane chain lengths. Alk1p and Alk3p were shown to hydroxylate n-alkanes of various carbon numbers. Alk10p, too, oxidized n-alkanes of a wide range of lengths, but was shown to preferentially oxidize shorter-chain n-alkanes. Alk2p, Alk6p, and Alk9p, on the Fig. 4 n-Alkane metabolism in Y. lipolytica. See text for details

n-Alkane

ER Alk proteins

Plasma membrane Peroxisome CH2OH

CH2OH

Adh? Fadh? Alk proteins?

Fao1

CHO

CHO

Hfd1, Hfd2A, Hfd4?

Hfd1, Hfd2A, Hfd2B, Hfd3

Faa1

Fat1

COOH

Acyl-CoA

Lipids

COOH

Acyl-CoA β-Oxidation

Acetyl-CoA

ä Fig. 3 (continued) CYP52F6 (O74132), CYP52F7 (O74133), CYP52F8 (O74134), CYP52F10 (A0A0K2S2A7), CYP52F11 (Q6CDW4), CYP52S1 (Q6CCE5), CYP52F9 (Q6CGD9), and CYP51 of Y. lipolytica (Q6CFP4). Species are indicated in parentheses as follows: C. albicans (Ca), C. maltosa (Cm), C. tropicalis (Ct), D. hansenii (Dh), S. stipitis (Ss), S. bombicola (Sb), and Y. lipolytica (Yl). P450s that were shown to catalyze the oxidation of n-alkanes or ω-termini of fatty acids are indicated as A or F, respectively

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other hand, preferred longer-chain n-alkanes. In Y. lipolytica, the genes encoding CYP52-family P450s are likely to have multiplicated after diverging from ancestral n-alkane-assimilating yeasts carrying one or a small number of genes encoding CYP52-family P450s.

4

Oxidation of Fatty Alcohols to Fatty Aldehydes

In n-alkane-assimilating yeasts, fatty alcohols are thought to be oxidized to fatty aldehydes by NAD+- or NADP+-dependent fatty alcohol dehydrogenase (FADH) in the ER or by H2O2-producing fatty alcohol oxidase (FAOD) in the peroxisome (Fig. 2) (Barth and Gaillardin 1996; Fickers et al. 2005; Fukuda 2013; Fukuda and Ohta 2013; Mauersberger et al. 1996; Tanaka and Fukui 1989). Such FAODs have been detected and characterized in several n-alkane-assimilating yeasts including C. tropicalis, C. maltosa, C. parapsilosis, and Y. lipolytica (Dickinson and Wadforth 1992; Kemp et al. 1994; Mauersberger et al. 1992). The FAOD-coding genes FAOT, FAO1, and FAO2 have been identified in C. tropicalis (Cheng et al. 2005; Eirich et al. 2004; Vanhanen et al. 2000). A FAOT deletion mutant exhibited defective growth on n-octadecane but not on shorter-chain n-alkanes or fatty acids, suggesting that FAOD encoded by FAOT is involved in the oxidation of fatty alcohol that is produced in the metabolism of n-octadecane and that one or more other enzymes are involved in the oxidation of shorter-chain n-alkanes (Cheng et al. 2005). In the genome sequence of Y. lipolytica, eight alcohol dehydrogenase genes, ADH1–ADH7 and FADH, and a fatty alcohol oxidase gene, FAO1, were identified (Gatter et al. 2014). Among these genes, a triple deletion mutant of ADH1, ADH3, and FAO1 showed severely defective growth on 1-dodecanol and 1-tetradecanol, but not on dodecanoic acid or tetradecanoic acid, suggesting that Adh1p, Adh3p, and Fao1p are involved in the assimilation of exogenous fatty alcohols (Iwama et al. 2015). Microscopic observation suggested that Fao1p localizes in the peroxisome. Adh1p and Adh3p are cytosolic proteins, but substantial amounts of these proteins were recovered in the membrane fraction of cell extracts, raising the possibility that Adh1p and Adh3p transiently localize to the membranes, possibly to the ER. A deletion mutant of ADH1–ADH7, FADH, and FAO1 exhibited slightly defective growth on n-decane and n-dodecane, but not on longer-chain n-alkanes. These results imply that any one or more of the enzymes encoded by these genes catalyze the oxidation of fatty aldehydes produced in the metabolism of shorter-chain n-alkanes (Fig. 4), but that one or more other enzymes are also involved in the oxidation of fatty alcohols derived from n-alkanes.

5

Oxidation of Fatty Aldehydes to Fatty Acids

In n-alkane-assimilating yeasts, fatty aldehydes generated during n-alkane metabolism are oxidized to fatty acids by fatty aldehyde dehydrogenase (FALDH) in the ER or in the peroxisome (Fig. 2). The model yeast S. cerevisiae has a single

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FALDH-coding gene, HFD1. S. cerevisiae Hfd1 and its mammalian ortholog ALDH3A2 are involved in the conversion of hexadecenal to hexadecenoic acid during the degradation of sphingosine-1-phosphate, a metabolite of sphingolipids and a second messenger involved in various cellular processes (Nakahara et al. 2012). ALDH3A2 also plays a role in the degradation of phytanic acid in the peroxisome (Ashibe et al. 2007; Verhoeven et al. 1998) as well as in protection from oxidative stress associated with lipid peroxidation (Demozay et al. 2004). Mutations in ALDH3A2 have been shown to cause Sjögren-Larsson syndrome (De Laurenzi et al. 1996). Hfd1 and ALDH3A2 belong to a superfamily of NAD (P)+-dependent aldehyde dehydrogenases (Sophos et al. 2001). In n-alkane-assimilating yeasts, FALDH activities have been reported in Candida intermedia, C. tropicalis, and Y. lipolytica (Liu and Johnson 1971; Ueda and Tanaka 1990; Yamada et al. 1980). Genes encoding FALDHs that are involved in the assimilation of n-alkanes were identified in Y. lipolytica (Iwama et al. 2014). The genome of Y. lipolytica contains four orthologs, HFD1–HFD4, of S. cerevisiae HFD1 and mammalian ALDH3A2. A Y. lipolytica mutant lacking all four HFD genes did not grow on n-alkanes of 12–18 carbons and showed severe growth defects on n-alkanes of 10 and 11 carbons. The expression of any one of these genes, however, restored the growth of the deletion mutant on n-alkanes. Furthermore, bacterially produced Hfd proteins exhibited dehydrogenase activities to dodecanal and tetradecanal in vitro. Fluorescence microscopic analysis suggested that Hfd1p localizes to the ER and the peroxisome and that Hfd3p localizes to the peroxisome. Two HFD2 transcript variants, which encode Hfd2Bp containing a peroxisomal targeting signal 1 (PTS1)-like sequence at its C-terminus and Hfd2Ap without PTS1, were generated from HFD2. Hfd2Ap has been suggested to localize in the ER and the peroxisome, while Hfd2Bp localizes to the peroxisome. These results imply that Hfd proteins are involved in the oxidation of fatty aldehydes produced during metabolism of n-alkanes in the ER and the peroxisome (Fig. 4) (Iwama et al. 2014); however, growth of the quadruple deletion mutant of HFD genes on dodecanal or tetradecanal indicated that one or more other enzymes are involved in the assimilation of exogenous fatty aldehydes. The n-alkane-assimilating yeasts, C. tropicalis, C. albicans, C. dubliniensis, C. parapsilosis, L. elongisporus, and M. guilliermondii, have multiple FALDH genes. Although involvement of these FALDH genes in the assimilation of n-alkanes remains to be examined, it is possible that they were multiplicated to efficiently degrade fatty aldehydes produced during the metabolism of n-alkanes.

6

Activation and Utilization of Fatty Acids

Fatty acids play a critical role as hydrophobic moieties in lipid molecules that constitute biological membranes or are used as energy and carbon sources via β-oxidation. Fatty acids are also involved in a variety of cellular processes as precursors of signaling molecules and hormones as well as by acylation of proteins. Fatty acids are used for these processes in the forms of acyl-CoAs, which are

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synthesized from fatty acids and coenzyme A by acyl-CoA synthetase (ACS). Multiple ACS isozymes are encoded by the genomes of eukaryotes including yeasts, and some ACS isozymes exhibit distinct substrate specificities and subcellular distributions (Black and DiRusso 2007; Soupene and Kuypers 2008; Watkins and Ellis 2012). Y. lipolytica has five ACSs, Faa1p and Fat1p–Fat4p, and ten ACS-like enzymes, Aa11p–Aal10p (Dulermo et al. 2014, 2016; Tenagy et al. 2015; Wang et al. 2011). A deletion mutant of FAT1 exhibited severely defective growth on n-decane of 10 and 12 carbons and partially defective growth on 14 and 16 carbons. The FAA1 deletion mutant exhibited retarded growth on n-alkane of 16 carbons, while deletion mutants of other ACS genes did not show any defects on n-alkanes. In addition, a double deletion mutant of FAT1 and FAA1 showed severe growth defects on nalkanes of 10–18 carbons (Tenagy et al. 2015). These results suggest that Faa1p and Fat1p play critical roles in the activation of fatty acids produced during n-alkane assimilation (Fig. 4). The wild-type strain of Y. lipolytica was shown to grow in the presence of cerulenin, an inhibitor of fatty acid synthesis, when n-octadecane was supplemented, suggesting that the stearic acid produced by the oxidation of n-octadecane is activated to stearoyl-CoA and that stearoyl-CoA or its derivatives support the growth of Y. lipolytica. The FAA1 deletion mutant, however, did not grow in the presence of cerulenin and n-octadecane, suggesting that FAA1 is involved in the activation of fatty acids derived from n-alkanes for essential cellular processes including membrane lipid synthesis. Fluorescent microscopic observation and fractionation analysis of cell extracts suggested that Fat1p localizes in the peroxisome, in agreement with the presence of a PTS1-like sequence at its C-terminus, while Faa1p localizes in the cytosol and to membranes. Roles of ACS isozymes of other n-alkane-assimilating yeasts in the metabolism of n-alkanes remain to be elucidated.

7

Research Needs

The oxidation pathway of n-alkanes to fatty acids in yeasts has been studied primarily in C. tropicalis, C. maltosa, and Y. lipolytica. Apart from the enzymes that are involved in the oxidation of fatty alcohols produced through the hydroxylation of n-alkanes, the enzymes that catalyze the steps in the oxidation of n-alkanes to fatty acids have been identified. As mentioned already, CYP52A3 of C. maltosa reportedly catalyzes the cascade of the sequential oxidation of n-hexadecane to hexadecanoic acid (Scheller et al. 1998). Accordingly, the CYP52-family P450s may catalyze the sequential oxidation of fatty alcohols to fatty acids in other n-alkane-assimilating yeasts. The aerobic degradation of n-alkanes and the enzymes involved in this degradation have been studied in only a limited subset of ascomycetous yeasts (C. tropicalis, C. tropicalis, and Y. lipolytica), while this process in other yeasts, particularly basidiomycetous yeasts, remains to be characterized. The metabolism of n-alkanes is regulated at the transcriptional level in C. tropicalis, C. maltosa, and Y. lipolytica. Transcription of genes encoding a subset of enzymes involved in the n-alkane metabolism is upregulated in the presence of

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n-alkanes in these yeasts (Endoh-Yamagami et al. 2007; Hirakawa et al. 2009; Iida et al. 1998, 2000; Kobayashi et al. 2013, 2015; Mori et al. 2013; Ohkuma et al. 1991, 1995; Sanglard et al. 1987; Yamagami et al. 2004). Details of the transcriptional regulation of n-alkane metabolic genes in yeasts are described and discussed in other chapters. Two organelles, the ER and the peroxisome, are involved in the metabolism of n-alkanes in yeasts. Fundamental and critical questions that remain to be answered about these organelles in the context of n-alkane metabolism in yeasts are (1) how n-alkanes are imported into cells and transported to the ER and (2) how hydrophobic metabolites, fatty alcohols, fatty aldehydes, and fatty acids are transported from the ER to the peroxisome during n-alkane metabolism. C. maltosa and C. tropicalis produce dicarboxylic acids from n-alkanes and excrete them into culture medium (Arie et al. 2000; Tanaka and Fukui 1989). In a dicarboxylic acid-hyperproducing mutant of C. maltosa, the transcription of a CmCDR1 gene encoding an ABC transporter is highly activated in the later phase of culture on n-dodecane (Sagehashi et al. 2013). It would be of interest to test whether overexpression of CmCDR1 improves the efficiency of dicarboxylic acid production from n-alkane. The uptake, intracellular transport, and excretion of lipophilic compounds in eukaryotic cells are important and fundamental issues of basic cell biology. The elucidation and subsequent optimization of the mechanisms of those processes will contribute to improvements in efficiency of the production of useful compounds from n-alkanes.

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Aerobic Degradation of Aromatic Hydrocarbons D. Pérez-Pantoja, B. González, and Dietmar H. Pieper

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Catabolism of Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Peripheral Reactions Preparing Aromatic Hydrocarbons for Ring-Cleavage . . . . . . . . 2.2 Side Chain Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Central Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 CoA Dependent Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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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 aromatic hydrocarbons are transformed by peripheral reactions to a restricted range of central intermediates, which are subject to ring-cleavage and funneling into the Krebs cycle. Key enzymes in aerobic aromatic degradation are D. Pérez-Pantoja Departamento de Bioquímica y Biología Molecular, Facultad de Ciencias Biológicas, Universidad de Concepción, Concepción, Chile e-mail: [email protected] B. González Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile e-mail: [email protected] D. H. Pieper (*) Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_10

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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. CoA dependent aerobic routes to allow ringcleavage 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 (Pérez-Pantoja et al. 2008). 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 (Duarte et al. 2014).

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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 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; Duarte et al. 2014). 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- cis-dihydroxycyclohexa-3,5-diene (benzoate- cis-dihydrodiol, see Fig. 1) (Reiner and Hegeman 1971). A benzoate dihydrodiol dehydrogenase catalyzes the dehydrogenation to a β-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 (αβ)3 quaternary structure (Wolfe et al. 2002), with each α-subunit containing a mononuclear non-heme iron active site and a Riesketype (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 parasubstituted 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

Fig. 1 Dendrogram showing the relatedness among oxygenase α-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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(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-carboxy-1,2-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 1,2-dioxygenases. 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 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 2,3-dihydroxy-2,3dihydro-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 α-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. Anthranilate dioxygenase of Burkholderia cepacia DBO1 is a three-component 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 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).

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Phylogenetic analyses (see Fig. 1) show that the α-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 (Duarte et al. 2014). 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 α-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-dihydroxy-3,5-cyclohexadiene-1,4-dicarboxylate as product, which is dehydrogenated, as described above for benzoate dihydrodiol, to a β-ketoacid which spontaneously decarboxylates to protocatechuate (Schläfli 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 (Schläfli et al. 1994). In contrast to salicylate 1- and 5-hydroxylase or anthranilate dioxygenase, terephthalate dioxygenase seems to be active as a two-component dioxygenase consisting of α- and β-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 α-subunits, a feature shared with carbazole dioxygenases, 3-chlorobenzoate 4,5-dioxygenase and isophthalate dioxygenase from C. testosteroni YZW-D (Wang et al. 1995). 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 α-subunit being closely related to α-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-3,4dihydro-3,4-dihydroxyphthalate 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

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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)-3,5cyclohexadiene-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.

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 (αβγ)2 quaternary structure and a diiron center contained in each α-subunit. Theses monooxygenases are classified according to their α-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

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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 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. 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).

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Fig. 2 Dendrogram showing the relatedness of single component flavoprotein monooxygenases. Reactions catalyzed by enzymes indicated are given to the exterior of the figure

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-3hydroxybenzoate (Entsch and van Berkel 1995). The purification from several bacterial sources and the presence of pobA homologous genes in Actinomycetes, α, β, and γ- Proteobacteria is indicative of a broad distribution of this enzyme. In a dendrogram of single-component 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 were 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).

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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. 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. 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; Ferrández 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 MhaB, described as an essential coupling protein, for activity. Thus it constitutes a novel type of two-component hydroxylase (AriasBarrau et al. 2005), distinct from the more classical two-component flavoprotein monooxygenases described below. 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).

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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. 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 2.2.1

Side Chain Processing

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 membraneassociated 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 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

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Fig. 3 Peripheral reactions involved in oxidation of methyl groups in methyl-substituted aromatic hydrocarbons. Reaction intermediates are shown in brackets

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 α subunit containing the active site and a FAD covalently linked to tyrosine, and a c-type cytochrome containing β 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 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).

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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 non-heme 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 α-subunits and shares similarity with phthalate 4,5-dioxygenases (see Sect. 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 2,6-dihydroxybenzoate, among other aromatic acids. Decarboxylases involved in the elimination of the carboxyl group from the aromatic nucleus in these pathways have been reported as non-oxidative (reductive) decarboxylases that do not require the external addition of any cofactor for its activity. Phthalate is metabolized by two different dioxygenase-initiated pathways (see Sect. 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

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Fig. 4 Metabolism of methoxylated aromatic hydrocarbons by Sphingomonas paucimobilis SYK-6. Enzymes catalyzing a given reaction only at slow rate are shown in italics

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 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.

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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 plantassociated 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 β-oxidative route (Overhage et al. 1999). p-Coumarate, caffeate and ferulate are transformed to 4-hydroxybenzoate, 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

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Fig. 5 Peripheral reactions in the bacterial degradation of ferulate, caffeate, p-coumarate, phenylalanine and tryptophan

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

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hydroxycinnamoyl-CoA thioesters that can be channeled to protocatechuate (Smith et al. 2003).

2.2.5

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 α-keto acid dependent oxygenases that typically require an α-keto acid (almost exclusively α-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 α-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 hemecontaining 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 3,4-dioxygenase (see Sect. 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.

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2.3

Central Reactions

2.3.1

Intradiol Ring-Cleavage Pathways

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 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,cismuconate, 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-1,2addition-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 enollactone (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

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Fig. 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

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 (Pérez-Pantoja et al. 2008). Enol-lactone is

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hydrolyzed by enol-lactone hydrolases which are assumed to use a Ser-His-Asp catalytic triade (Schlömann 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).

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 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, with reactions analogous to those of the classical 3-oxoadipate pathway (Prucha et al. 1997) being encoded by the mml gene cluster (Marin et al. 2010). In R. rhodochrous N75 hydrolysis of the lactone ring obviously occurs from 3-methylmuconolactone-CoA (Cha et al. 1998).

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. 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 nitroaromatic degradation.

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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) or naphthalene. 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; Duarte et al. 2014). 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). 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-2hydroxymuconic semialdehyde (from 4-methylcatechol) are preferentially degraded via the oxalocrotonate branch (Harayama et al. 1987). Since 2-hydroxy-6-oxo-2,4heptadienoate, 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, 4-dienoate 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.

Fig. 7 Extradiol ring-cleavage pathways involved in the degradation of catechol, 3-methylcatechol, protocatechuate, homoprotocatechuate, 2,3-dihydroxyphenylpropionate, 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

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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 β-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; Duarte et al. 2014) (Fig. 8). The protocatechuate 4,5-dioxygenolytic ring-cleavage product 4-carboxy-2hydroxymuconate-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-

Fig. 8 Dendrogram showing the relatedness of type II (LigB) extradiol dioxygenases. The physiological function of BphC6 2,3-dihydroxybiphenyl 1,2-dioxygenase from R. rhodochrous K37 remains to be established, but is possibly involved in fluorene degradation. (Taguchi et al. 2004)

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dihydroxyphthalate dehydrogenase (see Sect. 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-2hydroxymuconate) 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 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-hydroxy-2-oxoadipate by 4-oxalomesaconate hydratase (Hara et al. 2000). Finally, 4-carboxy-4-hydroxy2-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

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; Duarte et al. 2014). 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 Sect. 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).

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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; Duarte et al. 2014) (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-2,4dienoate (Fig. 7). The respective hydrolases show some substrate selectivity for the carboxylate of the side chain 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-hydroxy-6-ketonon2,4-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 Sect. 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-1,9-dioate hydrolases of Actinobacteria. Further metabolism of 2-hydroxypent-2,4-dienoate occurs as described above (see Sect. 2.3.2). Degradation of Gallate Only poor information is available so far on the metabolism of gallate (3,4,5trihydroxybenzoate). 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 Sect. 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

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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 β-subunit of protocatechuate 4,5-dioxygenases, whereas the C-terminal region has similarity to the corresponding small α-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. 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 β-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 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 Sect. 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. 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 chloroor nitroaromatics. Chlorobenzoquinol (and also benzoquinol) produced during the catabolism of γ-HCH (hexachlorocyclohexane) in S. japonicum UT26 is subject to direct ring cleavage by LinE extradiol dioxygenase, a type I extradiol dioxygenase. In contrast, benzoquinol 1,2-dioxygenase from the 4-hydroxyacetophenonedegrading P. fluorescens ACB has been shown to be an α2β2 heterotetramer where the α- and β-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

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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.

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 (Duarte et al. 2014). 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-1-carboxylate structural motif (Vaillancourt et al. 2006). 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, 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 two-domain 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

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Fig. 9 Metabolism of gentisate, homogentisate and salicylate via 1,2-dioxygenolytic cleavage.

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 (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 gram-positive 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). The Homogentisate Pathway Homogentisate is the central metabolite formed during degradation of aromatic amino acids phenylalanine and tyrosine in several microorganisms (see Sect. 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

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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). Direct Cleavage of Salicylate by Salicylate 1,2-Dioxygenase A 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-1-carboxylate 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. The 3-Hydroxyanthranilate Pathway 3-Hydroxyanthranilate is a central intermediate of tryptophan degradation via the kynurenine pathway (see Sect. 2.2.5) (Fig. 5) and of the biosynthetic pathway from tryptophan to quinolinate, 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-3carboxymuconic 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 Sect. 2.3.4.4) (Fig. 7). 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

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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 Sect. 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, an 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 phenylacetyl-CoA by a phenylacetate-CoA ligase (Mohamed 2000). Like other CoA ligases (see Sect. 2.2.4), this enzyme belongs to the AMP-forming acyl-CoA ligases, which catalyze thioesterification via a two-step process in which an acyladenosine monophosphate (AMP) intermediate is formed in the first step, followed by formation of the acyl-CoA ester and release of AMP. Phenylacetyl-CoA 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 α, β, and y subunits of the heteromultimeric diiron oxygenase component of the oxygenase. PaaB 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 1,2-dihydroxy-1,2dihydrophenylacetyl-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

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Fig. 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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β-oxidation-like pathway via β-ketoadipyl-CoA (Ismail et al. 2003) (Fig. 10) and a β-ketoadipyl-CoA thiolase that catalyses the last step of the phenylacetate catabolic pathway, i.e., the thiolytic cleavage of beta-ketoadipyl-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-2,3dihydroxybenzoyl-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 α 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-5oxocyclohex-1-enecarboxyl-CoA via 2-amino-5-oxocyclohex-1,3-dienecarboxylCoA (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 β-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).

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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 dead-end 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

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reconstruction studies linking in vivo with in silico catabolic properties and transcriptional studies would help to address this point. Moreover, 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 new isolates, it seems obvious that our current knowledge cover only a small proportion of the broad microbial degradative potential.

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8

Biosynthesis and Insertion of Heme Katrin Müller, Toni Mingers, V. Haskamp, Dieter Jahn, and Martina Jahn

Contents 1 2 3 4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Multiple Pathways for Heme Biosynthesis: An Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Two Routes for 5-Aminolevulinic Acid Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Common Part of Heme Biosynthesis from 5-Aminolevulinic Acid to Uroporphyrinogen III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 The Classical Pathway via Protoporphyrin IX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 The Alternative Pathway via Coproporphyrin III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 The Second Alternative Pathway via Siroheme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Heme b Insertion by the Heme Chaperone HemW . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

The red, iron containing tetrapyrrole heme is an essential cofactor of enzymes involved in the electron transport chain of energy generation and used for catalyzing chemically challenging reactions of the metabolism. It is also used for diatomic gas transport (O2, CO, CO2, NO, N2O), catalysis, and detection. Multiple transcriptional regulators and transporters bind heme. This chapter

K. Müller · T. Mingers · V. Haskamp · M. Jahn (*) Institute of Microbiology, Braunschweig University of Technology, Braunschweig, Germany e-mail: [email protected]; [email protected]; [email protected]; [email protected] D. Jahn Institute of Microbiology, Braunschweig University of Technology, Braunschweig Integrated Center of Systems Biology BRICS, Braunschweig, Germany e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_17

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focuses on the highly unusual pathways for heme biosynthesis and the integration of protoheme into target proteins. Today, three different biosynthetic routes for heme formation are known. The general precursor molecule of all tetrapyrroles 5-aminolevulinic acid is formed by two different pathways starting either with glutamyl-tRNA or succinyl-CoA and glycine. The conversion of 5-aminolevulinic acid to uroporphyrinogen III is common to all biosynthetic paths. Then the pathway branches to a classical route via protoporphyrin and two currently known alternative routes via coproporpyhrin III and siroheme. Various steps are catalyzed by up to three structurally unrelated enzymes. Finally, formed protoheme (heme b) gets actively inserted into proteins by the “Radical SAM” protein HemW. A detailed description of involved intermediates, enzymes, and their mechanisms are depicted below.

1

Introduction

Hemes are red colored, iron containing porphyrins which belong to the class of tetrapyrroles. The word heme is derived from the Greek αἷμα haima which stands for “blood.” These molecules are all composed of four pyrrole rings connected by methine bridges. The centrally coordinated iron can be used as electron source or sink and usually switches between the Fe(II), Fe (III), and Fe (IV) state. In humans, defects of the enzymes involved in heme biosynthesis lead to severe diseases called porphyrias (Kaufholz et al. 2013b). Overall, multiple different closed circular and open chain tetrapyrroles are known. Cyclic tetrapyrroles reveal characteristic reduction states of the ring system and typical metal irons chelated in the center (Dailey et al. 2017; Heinemann et al. 2008; Jahn and Jahn 2012; Layer et al. 2010). The pyrrole moieties are substituted by propionate, vinyl, acetate, and methyl groups. Prominent members of the tetrapyrroles are the magnesium coordinating, green bacteriochlorophylls and chlorophylls (Bröcker et al. 2012; Wang and Grimm 2015), the light absorbing pigments of oxygenic and anoxygenic photosynthesis. The yellow nickel containing coenzyme F430 plays an essential role in methanogenesis as part of the enzyme methyl coenzyme M (CoM) reductase (Moore et al. 2017; Zheng et al. 2016). Siroheme and heme d1 are technically no hemes (Boss et al. 2017). Siroheme is an iron containing isobacteriochlorin and serves as prosthetic group of assimilatory sulfite and nitrite reductases. Heme d1 is an iron containing dioxoisobacteriochlorin and the prosthetic group of the dissimilatory cd1 nitrite reductase. The cobalt containing corrins of the cobalamin or vitamin B12 class are part of methyltransferases, reductive dehalogenases, isomerases, and radical S-adensoyl-L-methionine (SAM) enzymes (Moore et al. 2017; Smith et al. 2018). However, the most ubiquitously found and versatile tetrapyrroles are hemes. They participate in various cellular processes including electron transfer, gas sensing and transport, signaling, catalysis, and transcriptional regulation.

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Multiple Pathways for Heme Biosynthesis: An Overview

For many decades, a unique biosynthetic pathway for the biosynthesis of hemes in all organisms was proposed (Bogorad 1958; Bogorad and Granick 1953; Hoare and Heath 1958). First variations were observed in the late 1980s for the formation of the general precursor 5-aminolevulinc acid (ALA, (Jahn 1992). Originally, the presence of aminolevulinic acid synthase was assumed to be ubiquitous throughout all kingdoms of life (Figs. 1 and 2). But for plants and later on for most bacteria and archaea, a different so-called C5-pathway, originating from the C5 skeleton of glutamate and proceeding via a glutamyl-tRNA intermediate, was discovered (Czarnecki and Grimm 2013; Jahn et al. 1992). Later in the biosynthetic pathway, variation occurred at the enzymatic steps known to require molecular oxygen, the coproporphyrinogen III (HemF) and protoporphyrinogen IX oxidase (HemY) reactions (Fig. 2). For their anaerobic metabolism, bacteria had developed the “Radical SAM” enzyme coproporphyrinogen III dehydrogenase (HemN) and an electron chain coupled flavin protoporphyrinogen IX oxidase (HemG) (Boynton et al. 2009; Layer et al. 2004, 2005; Möbius et al. 2010). In the meantime, a third protoporphyrinogen IX oxidase (HemJ) was discovered in cyanobacteria (Kato

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Fig. 1 Current knowledge about the biosynthesis of biologically active tetrapyrroles

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Glutamate

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Fig. 2 The threealternative pathways of heme biosynthesis. Typical intermediates and the end products are boxed

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et al. 2010; Skotnicova et al. 2018). With increasing access to multiple genomes, heme-synthesizing bacteria and archaea were discovered that lack genes for the enzymes of the late classical biosynthetic route to heme. Investigations from the late 1990s already proposed an alternative route via precorrin 2, an intermediate of the branch of tetrapyrrole biosynthesis toward cobalamin (B12), F430, siroheme, and heme d1 (Ishida et al. 1998). Novel enzyme activities were measured in Desulfovibrio vulgaris; however, due to the isolation of the reaction products as oxidized ester, their exact nature was not determined with absolute certainty. It was demonstrated in 2006 for the archaeon Methanosarcina barkeri and in 2009 for D. vulgaris that heme can be synthesized from precorrin 2 (Buchenau et al. 2006; Lobo et al. 2009). A novel pathway via siroheme was proposed and demonstrated (Bali et al. 2011; Kuhner et al. 2016; Storbeck et al. 2009). Additionally, in many Gram-positive bacteria, genes for coproporphyrinogen III oxidases/dehydrogenases were missing. Recently, for these organisms a novel pathway with coproporphyrin III as intermediate was described (Dailey et al. 2015; Lobo et al. 2015). Here, a coproporphyrinogen III oxidase produces this tetrapyrrole which in turn is subjected to iron insertion and final decarboxylation to yield heme (Dailey et al. 2015; Hansson et al. 1997a; Hobbs et al. 2016, 2017; Lobo et al. 2014). Interestingly, the siroheme and coproporphyrin pathways to heme share the final step, the decarboxylation of Fe-coproporphyrin III (coproheme) to form heme (Lobo et al. 2014). In summary, today we know three different pathways for the formation of heme with typical intermediates protoporphyrin, siroheme and coproporphyrin (Fig. 2).

3

Two Routes for 5-Aminolevulinic Acid Formation

The C5 compound ALA represents the general precursor of all known tetrapyrroles. Two different pathways for its formation are known. The most likely older pathway uses the C5 skeleton of glutamate as precursor (Beale and Castelfranco 1973). Glutamate is loaded onto tRNAGlu by glutamyl-tRNA synthetase, normally involved in protein biosynthesis (Schulze et al. 2006). An active site cysteine residue of glutamyl tRNA reductase (HemA, GtrR, GluTR) attacks the ester bond between the α-carbonyl of glutamate and tRNAGlu with the formation of an enzyme-bound thioester intermediate (Fig. 3) and the release of free tRNAGlu (Moser et al. 1999; Randau et al. 2004; Schauer et al. 2002). Hydride transfer from NADPH yields glutamate-1-semialdehyde (Lüer et al. 2007). Next, the pyridoxal-50 -phosphate/pyridoxamine-50 -phosphate-dependent glutamate-1-semialdehyde-2,1-aminomutase (HemL, GsaM) catalyzes the intramolecular transfer of an amino group using a modified aminotransferase mechanism (Grimm et al. 1992; Ilag and Jahn 1992). The product of both reactions is ALA. Currently, two crystal structures of GtrR and multiple structures of GsaM exist (Hennig et al. 1997; Li et al. 2018; Moser et al. 2001; Schulze et al. 2006; Zhao et al. 2014). Both enzymes form

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Fig. 3 Enzyme mechanisms of glutamyl-tRNA reductase (top) and 5-aminolevulinic acid synthase (bottom)

a stable channeling complex to protect the water-labile glutamate-1-semialdehyde intermediate (Lüer et al. 2005). In a second, often termed Shemin pathway, 5-aminolevulinic acid synthase (HemA, AlaS) catalyzes the condensation of the C4 compound succinyl-CoA and the C2 amino acid glycine with elimination of CO2 to form ALA (Fig. 2 (Gibson et al. 1958; Kikuchi et al. 1958; Shemin and Rittenberg 1945)). The pyridoxal50 -phosphate-dependent enzyme proceeds after lysine binding through an internal aldimine, after pro-R-hydrogen abstraction a quinonoid I intermediate is formed prior to succinyl-CoA binding of the 2-amino-3-ketoadipate and after the release of

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coenzyme A the quinonoid II intermediates (Fig. 3). Decarboxylation and protonation yields ALA (Kaufholz et al. 2013a; Stojanovski et al. 2014). Two crystal structures of the enzyme are known (Astner et al. 2005; Brown et al. 2018).

4

The Common Part of Heme Biosynthesis from 5-Aminolevulinic Acid to Uroporphyrinogen III

The monopyrrole porphobilinogen is formed via the asymmetric condensation of two ALA molecules (Dresel and Falk 1953; Granick 1954) by porphobilinogen synthase (Fig. 2, HemB, PbgS). For this purpose, the enzyme contains two ALA binding sites, termed A and P sites, referring to the contribution to the acetate or propionate moiety of porphobilinogen (Fig. 4). ALA 1 is bound via a Schiff base to a P site lysine, while ALA2 is also bound to a lysine residue of the sometimes metal containing A site (Spencer and Jordan 1995). This second Schiff base in the A site is converted to an enamine. In an aldol addition reaction, the C3 of ALA2 attacks the C4 of ALA1 with the formation of a C-C bond. Now the amino group of ALA 1 targets the Schiff base at the C4 atom of ALA2 yielding a C-N bond with the release from the lysine (Fig. 4). Subsequently, lysis of the remaining bond to the other lysine and aromatization of the formed ring system leads to porphobilinogen (Frere et al. 2002; Jaffe 2004). Overall three different binding sites for metal ions have been detected. These sites are filled in multiple combinations by zinc and magnesium (Jaffe 2016). Multiple crystal structures for the usually octameric enzymes have been elucidated (Erskine et al. 1997; Frankenberg et al. 1999; Frere et al. 2005; Jaffe et al. 2000). For the human enzyme, an equilibrium of functionally distinct octameric, hexameric, and two different dimers was described (Breinig et al. 2003; Jaffe and Lawrence 2012). Interestingly, the PbgS from Rhodobacter capsulatus is a hexamer without any metals (Bollivar et al. 2004). During the next two enzymatic steps, the first circular closed tetrapyrrole uroporphyrinogen III is formed by hydroxymethylbilane synthase (previously porphobilinogen deaminase, HemC, HmbS) and uroporphyrinogen III synthase (HemD, UroS). Originally, these reactions were believed to be catalyzed by one enzyme. However, at end of the 1960s, both activities were separated (Stevens and Frydman 1968). During the 1970s, the new intermediate hydroxymethylbilane (pre-uroporphyrinogen) was identified via 13C NMR and shown to be the substrate for uroporphyrinogen III synthase (Burton et al. 1979; Jordan and Seehra 1979). Hydroxymethylbilane synthase catalyzes the polymerization of four porphobilinogen pyrroles starting with ring A followed by rings B,C, and D of the final tetrapyrrole (Battersby et al. 1979; Jordan and Seehra 1979). In 1987, the existence of covalently attached dipyrromethane cofactor, formed by the enzyme from two molecules of porphobilinogen, serving as primer for the polymerization reaction, was shown (Azim et al. 2014; Hart et al. 1987; Jordan et al. 1988; Warren and Jordan 1988). During catalysis first the dipyrromethane cofactor is assembled once after ribosomal enzyme formation. Incoming porphobilinogens are deaminated and polymerized at the cofactor. The roles of conserved aspartic acid and arginine

Fig. 4 Enzymatic reaction for the formation of porphobilinogen from two molecules of 5-aminolevulinic acid by porphobilinogen synthase

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amino acid residues have been described (Bung et al. 2018; Pluta et al. 2018; Woodcock and Jordan 1994). Finally, the bond between the cofactor and the tetrapyrrole is hydrolyzed and hydroxymethylbilane is released (Fig. 5). In water, it would cyclize to fully symmetric uroporphyrinogen I, an inhibitor of the next enzyme, uroporphyrinogen III synthase. Multiple crystal structures are available (Azim et al. 2014; Gill et al. 2009; Roberts et al. 2013). Recently, a regulation of hydroxymethylbilane synthase by heme was proposed (Uchida et al. 2018). Uroporphyrinogen III synthase catalyzes the cyclization of linear hydroxymethylbilane with the inversion of ring D to form the cyclic but asymmetric uroporphyrinogen III. Uroporphyrinogen III as an intermediate of heme biosynthesis was proposed in the 1950s (Bogorad 1958). The enzyme was characterized and a mechanism proposed in the 1960s (Levin 1968; Mathewson and Corwin 1961). Catalysis starts with the loss of the hydroxyl group at ring A with the formation of the first azafulvene intermediate (Fig. 5). The reaction of the azafulvene with the substituted α-position of the D ring results in the formation of a spirocyclic pyrrolenine intermediate. A second azafulvene intermediate is formed on ring C by the breakage of the bond between rings C and D. This azafulvene reacts in the final steps with the free a-position before deprotonation, and rearrangement leads to the formation of uroporphyrinogen III (Hawker et al. 1998; Stark et al. 1985, 1986, 1993). Crystal structures for the enzyme have been published (Mathews et al. 2001; Peng et al. 2011; Schubert et al. 2008). Due to the low degree of amino acid conservation, the gene for the protein of a plant enzyme was discovered decades later (Tan et al. 2008).

5

The Classical Pathway via Protoporphyrin IX

Here, uroporphyrinogen III is first subject to a stepwise decarboxylation of each of the four pyrrole ring acyl side chains at the C2, C7, C12, and C18 to four methyl groups (Mauzerall and Granick 1958). The reaction is catalyzed by uroporphyrinogen III decarboxylase (HemE, UroD), which starts at ring D and proceeds clockwise via ring A and B to ring C (Jackson et al. 1976). The enzyme does not require any associated cofactor. The enzyme is a homodimer with juxtaposed and facing active sites. Two major different models were proposed for enzyme activity. In the first model, the homodimer shuttles a single substrate molecule forth and back between both active sites without solvent contact (Phillips et al. 2003; Whitby et al. 1998). In the second model, solely one active site of one subunit is used, the substrate is rotated by 90 following each decarboxylation. Interestingly, a dimer of an active and an inactive uroporphyrinogen III decarboxylase subunit is still able to perform the complete reaction pointing toward model 2 (Phillips et al. 2009). Several crystal structures have been reported (Fan et al. 2007; Martins et al. 2001; Phillips et al. 2003; Whitby et al. 1998). A uroporphyrinogen III decarboxylase structure with the product coproporphyrinogen III revealed that the substrate/product binds in a dome-shaped structure with the four NH groups facing a 2x hydrogen bond distance to a conserved aspartate residue. Furthermore, three conserved

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Porphobilinogen a

b

c

1-hydroxymethylbilane

Hydroxymethylbilane

Uroporphyrinogen III

Azafulvene intermediate

spirolytic pyrrolenine intermediate

Azafulvene intermediate

Fig. 5 Assembly of hydroxymethylbilane at a dipyrromethane cofactor by hydroxymethylbilane synthase (top) and the enzymatic conversion of hydroxymethylbilane to uroporphyrinogen III with the inversion of ring D catalyzed by uroporphyrinogen III synthase (bottom)

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arginines, one conserved histidine, and one tyrosine residues were proposed to be involved in catalysis. One of the arginine residues was identified as the general acid catalyst. Upon its protonation the decarboxylation reaction becomes rate-limiting (Silva et al. 2010). Regardless of the exact mechanism, the enzyme was suggested as a “benchmark” for catalytic proficiency among enzymes without cofactors due to a calculated enzyme enhancement value of the various decarboxylation reaction of around 1017 (Lewis and Wolfenden 2008). During the next biosynthetic step toward the formation of heme, the propionate side chain of coproporphyrinogen III ring A and B undergoes an oxidative decarboxylation to the corresponding vinyl groups by two different, structurally not related enzymes. The oxygen-dependent cofactor-free coproporphyrinogen III oxidase (HemF, CpgC) and the radical SAM enzyme coproporphyrinogen III dehydrogenase (HemN, CgdH) are catalyzing the conversion of coproporphyrinogen III into protoporphyrinogen IX. The general reaction was discovered in the 1950s (Granick and Mauzerall 1958) and the corresponding oxidase enzyme was described shortly thereafter (Sano and Granick 1961). First, it was shown that coproporphyrinogen III oxidase catalyzes the decarboxylation of ring A prior to that of ring B with the intermediate haderoporphyrinogen (Cavaleiro et al. 1974; Elder and Evans 1978). A mechanism for the oxygen-dependent reaction by the cofactor-free enzyme was proposed (Lash 2005; Silva and Ramos 2008). During the first step a base catalyzed deprotonation of the pyrrole NH-group yields an azacyclopentadienyl anion, which reacts with molecular oxygen at the α-position to form a pyrrole peroxide anion. Next, a proton at the β-positon of the substrate gets abstracted by the peroxide via a six-membered ring transition state with the formation of an exocyclic double bond (Fig. 6). Elimination of CO2 and H2O2 with the following bond rearrangements results in the formation of the vinyl group of the product protoporphyrinogen IX (Breckau et al. 2003). Solved crystal structures were without substrate (Lee et al. 2005; Phillips et al. 2004). Nevertheless, an aspartate and two conserved arginine residues were proposed to be in catalysis and substrate binding (Stephenson et al. 2007). Many bacteria possess anaerobic heme containing respiratory chains for energy generation. Consequently, an oxygen-independent coproporphyrinogen III conversion was needed. At the end of the 1960s Tait described such enzyme activity for Rhodobacter sphaeroides (Tait 1969, 1972). Around 10 years later the identical stereochemistry of an initial pro-S-hydrogen abstraction at the β-carbon as observed for the oxygen-dependent catalysis was described for the coproporphyrinogen III dehydrogenase (Seehra et al. 1983). Similarly, haderoporphyrinogen was identified as reaction intermediate (Rand et al. 2010). Genetic approaches led to the isolation of the corresponding genes in the 1990s (Lieb et al. 1998; Troup et al. 1995; Xu and Elliott 1994). Intensive biochemical and structural analysis of recombinant coproporphyrinogen III dehydrogenase from Escherichia coli identified the protein as “Radical SAM” enzyme. It carries a [4Fe-4S] cluster coordinated by three cysteine residues and one S-adenosyl-L-methionine (SAM) molecule (Layer et al. 2002, 2005; Lieb et al. 1998; Troup et al. 1995; Xu and Elliott 1994). The reaction starts with the reduction of the [4Fe-4S] cluster by an unknown electron donor. The electron is subsequently transferred to SAM, which in turn undergoes a homolytic

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Fig. 6 Enzymatic conversion ofcoproporphyrinogen III in coproporphyrinogen III by the coproporphyrinogen III oxidase (top) and the coproporphyrinogen III dehydrogenase (bottom)

cleavage with the generation of methionine and a 5' deoxyadenosyl radical. The highly reactive radical abstracts stereo-specifically the pro-S-hydrogen at the β-carbon (Fig. 6). Finally, elimination of CO2, transfer of the remaining electron to a yet unknown electron acceptor, and structural rearrangements with formation of the vinyl group finalize the reaction which has to occur twice (Layer et al. 2004, 2006). The crystal structure revealed the presence of two SAM molecules, which would allow a complete reaction cycle without release of a reaction intermediate (Layer et al. 2003). The six electron oxidation of protoporphyrinogen IX to the red colored protoporphyrin IX is catalyzed by three distinct enzymes all named protoporphyrinogen IX oxidase (HemY or PgoX, HemG or PgdH1, HemJ or PgdH2). Under aerobic conditions the reaction can occur auto-catalytically. Already in the early 1960s the corresponding enzyme was discovered (Porra and Falk 1961, 1964; Sano and Granick 1961). The FAD-containing, oxygen-dependent PgoX is found in all heme-synthesizing eukaryotes and a few Gram-negative bacteria. Currently, the crystal structure of three enzymes are known (Corradi et al. 2006; Koch et al. 2004; Qi et al. 2002), however, without bound substrate or product. A model for substrate binding was verified via kinetic studies (Heinemann et al. 2007). Due to the FAD cofactor one can assume that the reaction proceeds via three two-electron steps from porphyrinogen via tetrahydro and dihydro intermediates to the fully oxidized porphyrin. Kinetic studies revealed that three meso-carbon hydride ions are removed in a sequential fashion with the concomitant removal of the NH proton (Akhtar 2003). Alternatively, it was proposed that all hydride abstractions occur at the C-20

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meso-carbon including total ring hydrogen rearrangement via enamine-imine tautomerizations (Koch et al. 2004). Based on the PgoX structure, a complex with the following enzyme ferrochelatase was proposed and finally demonstrated (Koch et al. 2004; Masoumi et al. 2008). Again, PgoX is oxygen-dependent, and bacteria require an alternative system for anaerobic heme biosynthesis. First description of an oxygen-independent system channeling the six abstracted electrons into respiratory chains was reported in the 1970s from Jacobs and Jacobs (Ishihara et al. 1995; Jacobs and Jacobs 1978; Jacobs et al. 1970, 1971). In E. coli, the corresponding gene hemG was mapped and the corresponding mutant used for cloning of the gene (Nishimura et al. 1995; Sasarman et al. 1993). A detailed biochemical characterization of PgdH1 followed at the beginning of this decade (Boynton et al. 2009; Möbius et al. 2010). The FMN-containing protein belongs to the class of long chain flavodoxins. It was shown that the abstracted six electrons are transferred via ubiquinone to terminal oxidases (Cyo, Cyd) under aerobic conditions and via menaquinone to nitrate and fumarate reductase under anaerobic conditions (Möbius et al. 2010). A similar mechanism as proposed for PgoX is also possible for PgdH1 (Fig. 7). Interestingly, the only eukaryotic organism utilizing PgdH1 is the Leishmania major (Zwerschke et al. 2014). For the third enzyme PgdH2 (HemJ), little is known. The corresponding hemJ gene was discovered in the cyanobacterium Synechocystis 6803 (Boynton et al. 2011; membrane

cytoplasm

Cyo

H+

H2O

or

Aerobic conditions

O2

Cyd

H+ Q

QH2

HemG

FMN

protoporphyrinogen IX protoporphyrin IX

PgdH1 MQH2

MQ

Anaerobic conditions Frd or H+

Nar

fumarate succinate nitrate nitrite

H+

ADP + Pi ATPase ATP

Fig. 7 Respiratory chain coupled, six electron oxidation of protoporphyrinogen IX to protoporphyrin IX by the FMN containing protoporphyrinogen IX oxidase (HemG, PgdH1)

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Kato et al. 2010). The corresponding membrane protein contained heme and was proposed to interact with coproporphyrinogen III oxidase (Skotnicova et al. 2018). Protoporphyrin ferrochelatase (HemH, PpfC) catalyzes the final step of heme biosynthesis, the insertion of ferrous iron into protoporphyrin IX with the formation of protoheme (heme b). The first description of ferrochelatase from avian erythrocytes dates back to the 1950s (Ashenbrucker et al. 1956). Eukaryotic enzymes are usually membrane associated and contain with the exception of the plant enzymes one [2Fe-2S] cluster of unknown function per subunit (Shepherd et al. 2006). Bacterial enzymes are found with and without the cluster. The tetrapyrrole binds to the enzyme in an open conformation. Binding of the substrate triggers a rearrangement of a hydrogen bond network among conserved active site amino acid residues. Possibly, due to an abstraction of one pyrrole hydrogen a closed conformation is induced. Now the macrocycle is engulfed which causes an approximately 12 degree distortion of the bound tetrapyrrole. This distortion obviously facilitates metal chelation by the porphyrin with the simultaneous displacement of the second pyrrole hydrogen to a conserved histidine residue. The imidazole ring of the histidine moves, which causes structural rearrangements of the enzyme to adapt to the release conformation. Finally, heme b is released (Medlock et al. 2007, 2009; Sigfridsson and Ryde 2003; Wang et al. 2009, 2013).

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The Alternative Pathway via Coproporphyrin III

Gram-positive bacteria utilize an alternative, only recently discovered pathway for heme biosynthesis. Coproporpyhrinogen III synthesized via the classical pathway gets oxidized to coproporphyrin III followed by metal insertion with the formation of Fe-coproheme III. Finally, the propionate side chains at ring A and B of Fe-coproheme III get decarboxylated to form heme b (Dailey et al. 2015; Hansson et al. 1997b; Hobbs et al. 2016, 2017; Lobo et al. 2015). Obviously, oxidation of the ring system of the porphyrinogen to form a porphyrin occurs at the level of coproporphyrinogen III instead of protoporphyrinogen IX as seen in the classical pathway. Consequently, iron insertion follows and the decarboxylation of ring A and B, as performed at the level of coproporphyrinogen by two different enzymes (CpgC, CgdH) in the classical pathway, utilizes the iron containing Fe-coproheme III in the novel pathway. The first committed step of the novel pathway, the oxidation of coproporpyhrinogen III to coproporphyrin III is performed by a coproporpyhrinogen III oxidase (HemY, CgoX). It was originally described for Bacillus subtilis as protoporphyrinogen oxidase (Corrigall et al. 1998; Hansson and Hederstedt 1994; Qin et al. 2010). However, already then it was reported that the enzyme catalyzes the oxidation of coproporpyhrinogen III to coproporphyrin III at a higher rate as the proposed protoporpyhrinogen oxidation (Han et al. 2013; Hansson et al. 1997b). With the discovery of the novel pathway this observation makes sense. The mechanism of the oxygen-dependent six electron oxidation performed by the FAD enzyme is highly similar to the oxygen-dependent protoporphyrinogen oxidase (FgoX) mechanism with the abstraction of six protons from the porphyrinogen and

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the formation of three molecules H2O2. However, the active site pocket of CgoXs is with 1173 Å3 is much larger than those of FgoX protoporphyrinogen IX oxidases with 527 to 440 Å3 and contains more positively charged surface areas (Qin et al. 2010). One explanation is the accommodation of the remaining propionate side chain containing coproporpyhrinogen III by CgoX. The oxygen-independent enzyme is currently unknown. Next, coproporphyrinogen ferrochelatase (HemH, CpfC) catalyzes the insertion of ferrous iron into coproporpyhrin III to generate coproheme III. The best characterized enzyme is again from B. subtilis. It is water soluble and possesses a [2Fe-2S] cluster of unknown function. The mechanism is most likely highly similar to those of the ferrochelatase PpfC (Hansson et al. 2007; Karlberg et al. 2002; Lecerof et al. 2000, 2003; Olsson et al. 2002). The last step of the novel pathway is the decarboxylation of coproheme III to heme b and was named coproheme decarboxylase or heme synthase (ChdC, HemQ). An enzyme (AhbD) of identical catalysis is also part of the novel siroheme pathway for heme biosynthesis described below, but structurally not related to ChdC. While AhbD belongs to the family of “Radical SAM” enzymes, ChdC is a member of the chlorite dismutase family (Celis et al. 2015; Dailey et al. 2015; Pfanzagl et al. 2018). The structures of several incorrectly annotated as potential chlorite dismutase ChdCs were solved (PDB accession numbers 1T0T, 3DZT, 1VDH, 4WWS, SLOQ). The Listeria monocytogenes enzyme was co-crystallized with the product heme (Hofbauer et al. 2016). Two molecules of H2O2 are needed for catalysis (Celis et al. 2015; Hofbauer et al. 2014, 2016). During catalysis coproheme acts as substrate and cofactor. The coproheme ferric iron is coordinated by a conserved histidine residue. Under aerobic conditions O2 gets to the coproheme iron bound and oxidizes it to the ferric state. The subsequent second-order reaction between the ferric complex and H2O2 is slow and pH-dependent. First evidence for ferryl porphyrin cation radical was obtained (Streit et al. 2018). A tyrosine, hydrogen bonding to the propionate at ring A, is essential for decarboxylation. It is proposed that an oxidizing equivalent from the most likely radical allows for the formation of a tyrosine radical which abstracts the hydrogen from the propionate side chain. Migration of the unpaired propionyl electron back to the coproheme would yield ferric haderoheme and CO2. The propionate at ring B forms salt bridges to a lysine residue. Now, a similar pathway is proposed with this lysine as the essential proton shuttle for the second decarboxylation reaction (Celis et al. 2017). An alternative version of the ChdC protein called PitA, representing a fusion of ChdC with a monooxygenase domain, was described for the aerobic growth of Halferax volcanii (Kosugi et al. 2017).

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The Second Alternative Pathway via Siroheme

The second alternative pathway starts directly at uroporphyrinogen III and uses the initial steps of cobalamin, F430, heme d1, and the complete siroheme biosynthesis (Bali et al. 2014; Kuhner et al. 2014). The pathway was originally proposed for

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archaea but subsequently found in multiple bacteria (Storbeck et al. 2010). The three reactions of siroheme formation are the SAM-dependent methylation of uroporphyrinogen II at position C-2 and C-7 to form precorrin-2 via precorrin-1, the following NAD-dependent ring dehydrogenation of precorrin-2 to form sirohydrochlorin and NADH, and the final iron insertion into sirohydrochlorin to yield siroheme. These three catalytic challenging reactions can be performed by one multifunctional enzyme called siroheme synthase (CysG) (Spencer et al. 1993; Warren et al. 1990). The enzyme contains two independent enzymatic modules, one for the methyltransferase and the other for the combined dehydrogenaseferrochelatase function (Anderson et al. 2001; Lobo et al. 2009; Strey et al. 1999). Alternatively, the identical reactions are performed by three different enzymes, uroporphyrinogen-III C-methyltransferase (SUMT = NirE, CobA, SirA, Met1p) for the SAM-dependent methylation of uroporphyrinogen II at position C-2 and C-7 to form precorrin-2 via precorrin, the precorrin 2-dehydrogenase for the NAD-dependent ring dehydrogenation of precorrin-2 to form sirohydrochlorin, and the sirohydrochlorin ferrochelatase (SirB) for iron insertion. Interestingly, some bacteria carry a protein fusion of uroporphyrinogen III synthase and SUMT (Anderson et al. 2001; Lobo et al. 2009), most likely allowing direct channeling of the uroporphyrinogen III intermediate into the precorrin-2 pathway. Uroporphyrinogen-III C-methyltransferase (SUMT = NirE, CobA, SirA, Met1p) was first described for cobalamin biosynthesis in Pseudomonas denitrificans (Blanche et al. 1989). The crystal structures of SUMT involved in cobalamin and heme d1 biosynthesis were solved (Rehse et al. 2005; Storbeck et al. 2011; Vevodova et al. 2004). The NirE structure revealed the coordination of the tetrapyrrole by three arginines, a histidine, and a methionine residue. A mechanism induced by the arginine-mediated proton abstraction from the C-20 position was proposed (Storbeck et al. 2011). The subsequent movement of electrons facilitates the nucleophilic attack of C-2 at the methyl group of SAM. Upon proton abstraction from C-20 a new double bond between C-20 and C-1 is formed. After methyl transfer, rearrangements of double bonds within the macrocycle occur to form presorrin-1. After the first round of methylation the intermediate and side product SAH are released from the active site. Precorrin-1 and new SAM have to bind again to initiate a novel round of methylation at C-7 (Storbeck et al. 2011). Precorrin-2 dehydrogenase (SirC, Met8p) catalyzes the NAD+-dependent oxidation of precorrin-2 (dipyrrocorphin) to form sirohydrochlorin (Raux et al. 2003; Schubert et al. 2008). The identical reaction is catalyzed by domains of the multifunctional CysG and Met8p. The crystal structures of SirC and Met8p were solved. Both enzymes were found to bind metals including Co(II) and Cu(II). It was proposed that SirC evolved from a Met8p-type protein via loosing its chelatase domain (Raux et al. 2003; Schubert et al. 2008). Sirohydrochlorin ferrochelatase (SirB, Met8p) catalyzes the insertion of iron into sirohydrochlorin to form siroheme (Schubert et al. 2002). SirB solely catalyzes the iron chelation reaction. The multifunctional Met8p carries one active site with a catalytically important aspartate residue for both reactions, the dehydrogenase and chelatase reaction (Schubert et al. 2002).

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The heterodimeric Siroheme decarboxylase (NirDLGH, AhbAB) catalyzes the conversion of siroheme to didecarboxysiroheme (Palmer et al. 2014). The two acyl side chains attached to C-12 and C-18 are decarboxylated to the corresponding methyl groups. Surprisingly, the crystal structure of the Desulfovibrio desulfuricans protein was solved and revealed structural similarity to proteins of the Asn/Lrp transcriptional regulator family proteins. A enzyme mechanism was proposed (Palmer et al. 2014). The coproheme (Fe-coproporphyrin) synthase (AhbC) converts didecarboxysiroheme into coproheme (Fe-coproporphyrin). Recombinant AhbC protein from Methanosarcina barkeri was shown to transform in the presence of SAM and the reducing agent dithionite 12,18-didecarboxysiroheme into Fe-coproporphyrin III (coproheme). Thus, the enzyme catalyzes the loss of the acyl side chains at C2 and C7 (Bali et al. 2011). The exact enzymatic mechanism remains to be determined. Heme synthase (AhbD) catalyzes the oxidative decarboxylation of coproheme into heme b (Kuhner et al. 2016). The protein belongs to the “Radical SAM” family of enzymes. A close mechanistic relationship to the coproporphyrinogen III dehydrogenase reaction can be assumed. Interestingly, the protein contains two [4Fe-4S] clusters. Besides the “classical” [4Fe-4S] cluster I involved in coproporphyrinogen III dehydrogenase type catalysis, the second auxiliary [4Fe-4S] cluster was identified to be involved in the electron transfer to the final electron acceptor of the reaction (Kuhner et al. 2016).

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Heme b Insertion by the Heme Chaperone HemW

The “radical SAM” protein HemW revealed a high degree of amino acid sequence homology to coproporphyrinogen III dehydrogenases. However, the corresponding enzyme activity was never demonstrated for these proteins. A genetic investigation in Lactobacillus lactis indicated HemW’s participation in the generation of cytochromes. Subsequently, stable heme binding of the protein was demonstrated. Very recently the E. coli HemW was shown to be a heme chaperone involved in the active integration of heme b into the usually heme containing respiratory nitrate NarGHI which is involved in the anaerobic energy generation of the bacterium. The human counterpart RSAD1 was also shown to stably bind heme indicating its heme chaperone function (Abicht et al. 2012; Haskamp et al. 2018). Like other radical SAM proteins, HemW contains three cysteines and one SAM coordinating a [4Fe-4S] cluster. The intact iron-sulfur cluster is required for HemW dimerization, which in turn causes membrane localization. The intact iron-sulfur cluster is not required for stable covalent heme binding. Bacterioferritins and the heme-containing subunit NarI of the respiratory nitrate reductase NarGHI were shown to interact directly with HemW. Bacterioferritins might serve as heme donors for HemW, while the cytochrome subunit NarI of nitrate reductase represents the target of HemW. During contact heme covalently bound to HemW gets actively transferred to a heme-depleted, catalytically inactive nitrate reductase, restoring its nitrate-reducing enzyme activity (Fig. 8). For the transfer process, an intact a

BfrB

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HemW Heme

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+[4Fe-4S]

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HemW HemW

NarH NarG

HemW HemW

NO3-

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Periplasm

Fig. 8 Model of E. coli HemW activity. Heme from heme biosynthesis is most likely transferred via bacterioferritin (Bfr, light green) to the heme chaperone HemW (blue). The [4Fe-4S] cluster-containing HemW dimerizes and localizes to the membrane, where it interacts with its target protein NarI (yellow), a subunit of the respiratory nitrate reductase NarGHI. After heme incorporation into apo-NarI, the holo-NarGHI catalyzes the reduction of nitrate to nitrite

5-Aminolevulinic acid

Protoporphyrinogen IX

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[4Fe-4S] cluster is required. The exact mechanism of heme transfer remains to be determined (Abicht et al. 2012; Haskamp et al. 2018).

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Research Needs

Some of the enzyme activities of the various alternative pathways, including the oxygen-independent coproporphyrinogen oxidase for the formation of coproporphyrin III, are unknown. Moreover, the exact nature of various enzymatic mechanisms and enzyme structures require further investigations. Possibly, additional alternative pathways shall be discovered. Acknowledgments We thank Stefan Barthels for his excellent technical assistance and are indebted to the Deutsche Forschungsgemeinschaft (GRK 2223, PROCOMPAS) for funding.

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Part II Biochemistry of Aerobic Degradation of Lipids

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Membrane Lipid Degradation and Lipid Cycles in Microbes Diana X. Sahonero-Canavesi, Isabel M. López-Lara, and Otto Geiger

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Key Enzymes for Membrane Lipid Degradation and Rounding Off Lipid Cycles . . . . . . . . 2.1 Phospholipases, Lipases, and Transferases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Acylating Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Diacylglycerol Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Phospholipid Turnover and Lipid Cycles in Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Role of DAG and Lands’ Cycle in Eukaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Lipid Cycles in Escherichia coli and Other Enterobactericeae . . . . . . . . . . . . . . . . . . . . . . 3.3 Complex DAG Cycles and Metabolism in Sinorhizobium meliloti . . . . . . . . . . . . . . . . . . 3.4 Diacylglycerol Cycles in Gram-Positive Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Emerging Amino Acyl-Phosphatidylglycerol Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Lipid Cycles Involving Undecaprenyl Phosphate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Physiological Implications of Membrane Lipid Turnover and Lipid Cycles . . . . . . . . . . . . . . 6 Conclusion and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

All living cells are delimited from the exterior world by a membrane, and membrane-forming lipids are the structural determinants for membrane assembly and maintenance. Although biosynthesis of membrane-forming lipids is well understood in many organisms, turnover, degradation, and remodeling of these lipids are less studied. An initial degradation of glycerol-containing membrane lipids may occur by (phospho)lipases or transferases which remove distinct groups from the membrane lipid converting it into a lysolipid or diacylglycerol. These degradation intermediates can either be totally degraded into low-molecular-weight D. X. Sahonero-Canavesi · I. M. López-Lara · O. Geiger (*) Centro de Ciencias Genómicas, Universidad Nacional Autónoma de México, Cuernavaca, Morelos, Mexico e-mail: [email protected]; [email protected]; [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_38

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metabolites or missing groups can be reintroduced onto the intermediates to convert them into fully functional membrane lipids again, thereby completing a lipid cycle. Classic examples in Escherichia coli are the lyso-phosphatidylethanolamine cycle, the diacylglycerol cycle, or cycles involving the isoprenoid undecaprenol. It is evident that many more lipid cycles exist in other proteobacteria and in grampositive bacteria and that these cycles play major roles in decorating biomolecules located outside the cytoplasmic compartment.

1

Introduction

In Bacteria and Eukarya, sn-glycerol-3-phosphate forms the backbone of all glycerophospholipid (PL) molecules, whereas the backbone in Archaea is formed by sn-glycerol-1-phosphate. In bacteria, acylations at the 1-position and subsequently at the 2-position lead to the formation of phosphatidic acid (PA) (Fig. 1). Then PA is converted to CDP-diacylglycerol (CDP-DAG), the central activated intermediate which can be modified with distinct headgroups, leading to the formation of phosphatidylethanolamine (PE), phosphatidylglycerol (PG), and cardiolipin (CL) in Escherichia coli (Fig. 1), plus phosphatidylcholine (PC), phosphatidylinositol (PI), or aminoacylated versions of PG or CL in some other bacteria (López-Lara and Geiger 2016). Turnover of membrane phospholipids can be achieved by phospholipases or transferases leading to a partial degradation of PLs and the formation of lysophospholipids (lyso-PLs), i.e., lyso-PE, diacylglycerol (DAG), or PA (Fig. 1). Fig. 1 Scheme on glycerophospholipid (PL) formation, turnover, and cycles in the bacterium Escherichia coli (for details see text)

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While PA is rapidly reintegrated into de novo phospholipid biosynthesis, cells need to prevent the accumulation of harmful lyso-PLs or DAG in its membrane in order to avoid lysis. The accumulation can be avoided by either totally degrading lyso-PLs (by lysophospholipases) or DAG (by DAG lipase) or by reconverting them into fully functional PLs employing acyltransferases in the case of lyso-PLs or a DAG kinase in the case of DAG (Fig. 1), thereby creating metabolic lipid cycles. In the well-studied model bacterium E. coli, partial degradation of membrane phospholipids followed by their reformation gives rise to cycles which have been known as the diacylglycerol (DAG) cycle and the lyso-PE (2-acylglycerolphosphoethanolamine) cycle (Rock 2008) for some time. Here, we present an overview on membrane lipid degradation and metabolic lipid cycles in microbes keeping the main focus on bacteria.

2

Key Enzymes for Membrane Lipid Degradation and Rounding Off Lipid Cycles

2.1

Phospholipases, Lipases, and Transferases

Many bacterial sphingomyelinases and phospholipases are secreted, function as important virulence factors of pathogenic bacteria, and usually degrade membrane lipids of their hosts (Flores-Díaz et al. 2016). However, other intrinsic phospholipases or transferases act on the bacterium’s own membrane phospholipids turning them over into partial breakdown products (Figs. 1 and 2). For example, phospholipase (PLA1) and (PLA2) hydrolyze the ester bonds of intact glycerophospholipids at C1 or C2 of the glycerol moiety, respectively, leading to the release of a free fatty acid and the corresponding formation of a 1-lyso-phospholipid or a 2-lysophospholipid (Fig. 2). If one of the fatty acids has been removed by a type A phospholipase, the second fatty acid can be removed by a lysophospholipase generating another fatty acid and a glycerophosphoalcohol. Phospholipases C and D each split one of the phosphodiester bonds, forming a free alcohol and phosphatidic acid in the case of phospholipase D (PLD) or a free phosphoalcohol and diacylglycerol in the case of phospholipase C (PLC). In E. coli, the detergent-resistant outer membrane PLA1 PldA (OMPLA) degrades phospholipids by rapidly removing fatty acyl residues from the sn-1 position and subsequently from the sn-2 position (Zhang and Rock 2016). It is thought that PldA acts only on glycerol-based lipids mislocalized to the outer layer of the outer membrane, thereby contributing to stability and rigidity of the outer membrane in proteobacteria. Inner membrane lysophospholipases in E. coli include lysophospholipase L2 PldB, which hydrolyzes 2-acylglycerophosphoethanolamine efficiently but is barely active on the 1-acyl isomer (Zhang and Rock 2016). PldB also catalyzes the transfer of fatty acids from 2-acylglycerophosphoethanolamine to the terminal hydroxyl of the headgroup of PG to form acyl-PG. Recent evidence suggests that also DAG can be degraded by DAG lipase DglA to monoacylglycerol and a fatty acid and further to glycerol and another fatty acid at least in some

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Fig. 2 Partial glycerophospholipid (PL) degradation by distinct phospholipases (for details see text)

bacteria (Sahonero-Canavesi et al. 2015). While the cleavage of phospholipid ester bonds by phospholipases is achieved through a nucleophilic attack by water, transferases facilitate similar nucleophilic attacks, however, by alcohols forming again one of the products obtained by partial hydrolytic phospholipid degradation, but also a new compound in which one of the functional groups of a PL has been attached to a target molecule via a newly formed ester bond. Therefore, transferases are important players in lipid turnover, degradation, and reacylation of PLs, especially in extracytoplasmatic compartments. For example, transfer of phospho-alcohol head groups by transferases occurs during the formation of osmoregulated periplasmic glucans or lipoteichoic acids (see below). Transfer of acyl residues can occur from or to a glycerol-based lipid as outlined in the following paragraph.

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Acylating Enzymes

Enzymes involved in acylating glycerol derivatives usually employ activated fatty acyl compounds either in their thioester forms, such as CoA or acyl carrier protein (ACP) derivatives, or as mixed anhydrides, such as phosphate or AMP derivatives. During de novo PL biosynthesis in bacteria (Geiger et al. 2018), the initial acylation of glycerol-3-phosphate may be performed by the PlsB transferase (employing acyl-CoA or ACP thioester derivatives) or by PlsY (employing acylphosphate). The second acylation, performed on lyso-PA, is achieved by the action of PlsC (employing acyl-CoA or ACP thioesters) leading to the formation of PA. Many other important acylating enzymes are involved in reacylating previously formed lyso-PLs, thereby completing the so-called Lands’ cycle (see below) in eukaryotes and mostly they consume acyl-CoA derivatives (Shindou and Shimizu 2009). In E. coli, an acyl-ACP synthetase (Aas) has been initially described which can acylate ACP in vitro with the consumption of ATP. However, the physiological function of this enzyme is to reacylate lyso-PLs that had previously been formed during the lyso-PE cycle. In this case, the PlsC domain of the Aas-PlsC fusion protein transfers the acyl residue from acyl-ACP to the lyso-PL converting it into a complete PL again. Although some acyl-ACP derivatives can be formed with Aas from E. coli, the Aas from Vibrio harveyi has broader substrate specificities and seems to be a much more widely applicable tool for generating distinct acyl-ACP derivatives (Beld et al. 2014). Acylation reactions during de novo biosynthesis of lipid A consume acyl-ACP derivatives and are occurring on the inner leaflet of the cytoplasmic membrane (Raetz et al. 2007). In contrast, acylations of lipid A in the outer leaflet of the outer membrane employ acyl transferases which transfer ester-linked fatty acyl groups from PLs to the 3-hydroxy group of a lipid A-linked fatty acid, thereby forming a new ester bond. An example for this latter reaction is the PagP transferase in the outer membrane of E. coli.

2.3

Diacylglycerol Kinase

Diacylglycerol (DAG) kinase phosphorylates DAG to yield PA. In yeast, DAG kinase Dgk1 uses CTP for PA formation and, together with PA phosphatase Pah1, forms a lipid cycle that controls the PA/DAG balance with implications on numerous phenotypes (Qiu et al. 2016). Obviously, at a given physiological situation, either PA or DAG is formed whereas the other is consumed, shifting relative amounts of both. In bacteria, DAG kinase uses ATP to phosphorylate DAG. DAG kinase DgkA of proteobacteria, such as E. coli or S. meliloti, is an integral membrane protein and surprisingly the closest homologs of DgkA in gram-positive bacteria have the function of an undecaprenol kinase. In contrast, the soluble DAG kinase DgkB is

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present in Bacillus subtilis which attaches to the membrane in order to phosphorylate DAG and DgkB is required for the production of lipoteichoic acid (Jerga et al. 2007).

3

Phospholipid Turnover and Lipid Cycles in Microbes

3.1

Role of DAG and Lands’ Cycle in Eukaryotes

It is known for some time that in eukaryotes, PLs are formed by a de novo biosynthesis pathway (Kennedy pathway) (Nelson and Cox 2017) and modified by a remodeling pathway (Lands’ cycle) (Lands 1958) to generate membrane asymmetry and diversity (Shindou and Shimizu 2009). In the de novo pathway, glycerol-3phosphate is converted to lyso-PA by glycerol-phosphate acyltransferase and further to PA by lyso-PA acyltransferase, using acyl-CoAs as donors in both cases. In eukaryotes, PA can be converted into DAG or CDP-DAG and DAG is substrate in reactions that lead to the formation of triacylglycerol, PC, or PE. In contrast, PI, PS, PG, and CL can be formed directly from CDP-DAG in reactions reminiscent of bacterial PL biosynthesis. After phospholipids have been synthesized in the de novo pathway, their fatty acid composition at the sn-2 position can be altered by the remodeling pathway (Lands’ cycle) through the concerted actions of PLA2s and lyso-PL acyltransferases. One of the prominent examples of partial PL degradation occurs in animals, when the action of a PLA2 liberates arachidonic acid in the initial step of eicosanoid formation and signaling (Nelson and Cox 2017). Lyso-PLs can be reacylated by lyso-PL acyltransferases completing the cycle now known as the Lands’ cycle (Lands 1958). Usually, several PLA2s and an even larger number of lyso-PL acyltransferases, displaying distinct substrate specificities, are encountered in each eukaryote. As detailed descriptions go beyond the scope of this chapter, we refer the reader to recent reviews presenting the knowledge on acyl-CoA-consuming lyso-PL acyltransferases (Shindou and Shimizu 2009) and on lipid acyl chain remodeling in yeast (Renne et al. 2015) in some more detail. When G protein-coupled receptors are activated by hormone ligands, some receptors activate a phosphatidylinositol-4,5-bisphosphate-specific phospholipase C producing two potent second messengers, DAG and inositol-1,4,5-trisphosphate. While inositol-1,4,5-trisphosphate provokes release of Ca2+ from the endoplasmic reticulum to the cytosol, DAG, in cooperation with Ca2+, activates protein kinase C leading to some of the cellular hormone responses (Nelson and Cox 2017). As mentioned above, even in yeast, a lipid cycle exists that controls the PA/DAG balance with implications on numerous phenotypes (Qiu et al. 2016).

3.2

Lipid Cycles in Escherichia coli and Other Enterobactericeae

Besides cycles of metabolism of lipids, other cycles of lipid transport might exist and, in some cases, they might even be interconnected. For example, the gastrointestinal pathogen Salmonella typhimurium, responds to acidic pH and cationic

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antimicrobial peptides (CAMP) through the PhoPQ two-component regulatory system, which stimulates PagP-catalyzed lipid A remodeling as well as conversion of PG to palmitoyl-PG in the outer membrane, CAMP resistance, and intracellular survival within the acidified phagosome (Dalebroux et al. 2014). Notably, PhoPQ controls also the association of the periplasmic domain-containing inner transmembrane protein PbgA to the outer membrane and thereby the transport of CL from the inner to the outer membrane, which might occur via a periplasmic PbgA bridge (Dalebroux et al. 2015). Another example for trafficking between membranes is provided by the retrograde transport of PLs from the outer to the inner membrane. This transport is initiated by sorting machines, such as the MlaA lipoprotein, that removes phospholipids from the outer leaflet of the outer membrane in order to maintain lipid asymmetry of the outer membrane (Sutterlin et al. 2016), followed by retrograde transport of PLs from the outer back to the inner membrane. However, in this chapter, we will not engage in covering transport of lipids between distinct membranes but we will focus on metabolic cycles of membrane lipids. In E. coli, partial degradation of membrane phospholipids followed by their resynthesis gives rise to cycles known as the DAG cycle and the lyso-PE (2-acylglycerolphosphoethanolamine) cycle (Rock 2008).

Lyso-PE Cycle in E. coli and Its Role in Lipoprotein Maturation and Processing Lipoproteins (Lpps) are synthesized in the cytoplasm as protein precursors with an N-terminal signal sequence (Konovalova and Silhavy 2015). In E. coli, lipoproteins are translocated across the inner membrane by the Sec translocon and maturation and processing takes place on the periplasmic side of the cytoplasmic membrane (Fig. 3). In an initial step, a DAG residue is transferred from PG to the sulfhydryl group of a conserved cysteine residue in Lpps by PG/prolipoprotein diacylglyceryl transferase Lgt forming a thioether linkage. This modification is a prerequisite for the removal of the signal sequence by the lipoprotein signal peptidase Lsp, generating the DAG-substituted apolipoprotein with a new N-terminus provided by the conserved Cys. The final acyl transfer is achieved by the apolipoprotein N-acyltransferase Lnt which transfers an acyl residue from the sn-1-position of PE to the N-terminus of the apolipoprotein, i.e., the major outer membrane lipoprotein Lpp (Braun’s lipoprotein), resulting in the formation of lyso-PE and of the mature triacylated lipoprotein (Fig. 3). The mature triacylated Lpps are transported to the inner leaflet of the outer membrane by the “localization of lipoproteins” (LOL) export pathway, where they are destined to perform distinct functions (Pailler et al. 2012). For example, Braun’s lipoprotein Lpp is connected to the peptidoglycan cell wall and important for the maintenance of cell envelope architecture and stability. The lyso-PL 2-acylglycerolphosphoethanolamine (2-acyl-GPE) is a minor lipid in E. coli and is rapidly transferred back to the cytoplasmic surface of the inner membrane by the lysophospholipid transporter LplT (Harvat et al. 2005) (Fig. 3). The acyltransferase PlsC/acyl-ACP synthetase Aas reacylates lyso-PE using acylACP as the acyl donor to regenerate PE. However, not only PE can be turned over by 3.2.1

Fig. 3 Lyso-PE (2-acylglycerophosphoethanolamine) cycle in E. coli (for details see text). Red bars indicate the peptide membrane anchor formed by the N-terminal signal sequence of Lpp. Blue arrows indicate transport across the membrane in the case of lipids by flippases

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this lipid cycle; PG or CL can serve as acyl donors in the Lnt reaction as well (Gupta et al. 1991) and deacylated versions of PG or CL can be transported back to the cytoplasmic face of the inner membrane, where they can be reacylated by Aas (Lin et al. 2016). PE molecules are flipped from the cytoplasmic side of the inner membrane to the periplasmic side by the MsbA flippase, thereby completing the so-called lyso-PE cycle (Fig. 3). Homologues of Lnt are widespread in bacteria and the LplT, PlsC, and Aas domains (often in fused versions) are found throughout the proteobacteria and chlamydia (Lin et al. 2016). However, although the lyso-PE cycle is well documented in E. coli, due to a lack of experimental evidence, it remains an open question whether this cycle is complete in other bacteria or whether in some of them lyso-PE is further degraded instead of being reacylated. Even E. coli possesses the inner membrane lysophospholipase PldB, which hydrolyzes 2-acyl-GPE providing an alternative route how to deal with lyso-PE, which is by total degradation. Another source for the formation of lyso-PLs is provided upon acyltransfer from a phospholipid to PE generating N-acylphosphatidylethanolamine in E. coli (Mileykovskaya et al. 2009).

Classic Diacylglycerol (DAG) Cycles in E. coli upon Formation of Osmoregulated Periplasmic Glucans The periplasm contains the murein (peptidoglycan) sacculus, a variety of proteins found exclusively in this compartment, and osmoregulated periplasmic glucans (OPGs) present in the α-, β-, and γ-proteobacteria (Bontemps-Gallo and Lacroix 2015). Although OPGs exhibit quite different structures among various species, they are all glucose-derived oligosaccharides, comprise 5–24 glucose units, and are increasingly formed as medium osmolarity decreases. OPGs in E. coli were described initially by Kennedy’s group (van Golde et al. 1973) as a result of studies on PL turnover. In this bacterium, the rapid PG turnover is associated with the transfer of sn-1-phosphoglycerol headgroups to a class of oligosaccharides, named as a consequence “membrane-derived oligosaccharides” (MDOs) (Bohin 2000). MDOs from E. coli are linear glucans containing 5–12 glucose units joined by β-1,2 linkages and branched by β-1,6 linkages. Biosynthesis of MDOs is initiated by the transmembrane glucosyl transferase OpgH, with ACP as a cofactor and UDP-glucose as a substrate, and results in the formation of linear β-1,2-linked glucose units (Fig. 4). Although polyprenol phosphate is required for glucosyl transferase activity (Weissborn et al. 1991), the mechanistic role of the isoprenoid in this reaction remains obscure. OpgG is a periplasmic glucosyl transferase branching glucose units on this linear backbone by β-1,6 linkages (Bontemps-Gallo et al. 2013) (Fig. 4). OpgD is a paralog of the OpgG transferase affecting the degree of OPG polymerization. While OpgG appears to be secreted via the Sec system to the periplasm, OpgD exhibits a Tat signal sequence. The phosphoglycerol transferase OpgB (formerly known as MdoB) catalyzes the transfer of sn-1-phosphoglycerol from PG to the β-glucan forming phosphoglycerol-substituted OPGs and DAG as a second product. In turn, DAG is flipped to the inner surface of the cytoplasmic membrane, phosphorylated by 3.2.2

Fig. 4 Classic DAG cycles in E. coli during the formation of OPGs (for details see text). Blue arrows indicate transport across the membrane in the case of lipids by flippases. CdsA: CDP-diglyceride synthase; PgsA: phosphatidylglycerol-phosphate synthase; Pgp: phosphatidylglycerol-phosphate phosphatase; PssA: phosphatidylserine synthase; Psd: phosphatidylserine decarboxylase

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DAG kinase DgkA and converted into PA, thereby entering de novo PL biosynthesis again (Rock 2008) (Fig. 4). CdsA, PgsA, and Pgp covert PA to PG and flipping of PG to the outer surface of the cytoplasmic membrane completes the “classic” DAG cycle. An alternative way to provide OPGs with negative charges consists in the transfer of succinyl residues from succinyl-CoA to OPGs by the OpgC transferase. In addition to sn-1-glycerol and succinate residues, OPG can also be decorated to a minor extent with phosphoethanolamine (PEA) residues. These PEA residues are derived from PE and are transferred by the phosphoethanolamine transferase OpgE (Bontemps-Gallo et al. 2013) generating phosphoethanolamine-substituted OPGs and DAG as second product. Again, DAG can be recycled to PE by the previously described steps and the action of PssA and Psd (Fig. 4).

3.2.3 Lipid Cycles Modifying Enterobacterial Lipid A Lipid cycles may also be initiated when PEA headgroups are transferred from PE or acyl groups from PLs to lipid A (Henderson et al. 2016). Ethanolamine transferases that move PEA from PE to lipopolysaccharides or lipooligosaccharides in gram-negative bacteria usually consist of a transmembrane domain anchoring the enzyme to the periplasmic face of the cytoplasmic membrane linked to a catalytic domain on the periplasmic side of the membrane. Members of this family include E. coli EptB (PEA transferase B) which catalyzes the transfer of PEA from PE to the outer 2-keto-3-deoxyoctanate (Kdo) residue of LPS (Fig. 5) and EptA (formerly PmrC) which catalyzes the transfer of PEA to the 1-phosphate headgroup of lipid A in Salmonella enterica (Fig. 5). Also, when PEA residues are transferred from PE to lipopolysaccharide in such PEA transferase reactions, DAG is formed as the second product (Reynolds et al. 2005) and needs to be recycled to PA. Bivalent cations such as Mg2+ are important for maintaining the outer membrane permeability barrier. Low Mg2+ concentrations are sensed by the PhoP/PhoQ signal transduction system triggering the expression of the outer membrane PagP acyltransferase. PagP transfers a palmitate chain from a PL to the hydroxyl group of the N-linked 3-OH-14:0 chain on the proximal glucosamine unit of lipid A (Fig. 5) forming a lyso-PL as the second product. This lyso-PL is transported to the inner membrane and reacylated (Hsu et al. 1989). PagP-modified lipid A provides increased resistance to CAMP. Other modifications of lipid A in the outer membrane include Salmonella lipid A deacylases PagL and LpxR which remove fatty acids from specific positions of the lipid A molecule (Fig. 5). LpxT transfers a phosphate residue from undecaprenyl pyrophosphate (undecaprenyl diphosphate) to the 1-position of lipid A converting lipid A 1-phosphate to lipid A 1-diphosphate and forming undecaprenyl phosphate as the second product (Touze et al. 2008). LpxT is thought to be important for regenerating undecaprenyl phosphate from undecaprenyl pyrophosphate in order to permit ongoing murein biosynthesis (see below). Remarkably, LpxT breaks an old and forms a new phosphoanhydride bond and thereby provides lipid A 1-diphosphate with a “high-energy” bond.

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Fig. 5 Lipid cycle-induced modifications of the Kdo2-lipid A domain of Salmonella enterica (for details see text)

3.3

Complex DAG Cycles and Metabolism in Sinorhizobium meliloti

S. meliloti usually synthesizes PG, CL, PE, monomethyl-PE, and PC as its major membrane lipids when grown in culture media rich in phosphate (Geiger et al. 1999). However, under phosphorus-limiting conditions, S. meliloti replaces most of its phospholipids by membrane-forming lipids that do not contain phosphorus, such as sulfoquinovosyl diacylgycerol (SQDG), ornithine-containing lipid, and diacylglyceryl-N,N,N-trimethylhomoserine (DGTS) (Geiger et al. 1999). These phosphorus-free membrane lipids are unimportant for the symbiotic life style of S. meliloti, but they are required for optimal growth under phosphorus-limiting conditions (López-Lara et al. 2005). Upon phosphorus limitation, PC and PE are degraded in S. meliloti by a phospholipase C (PlcP) (Zavaleta-Pastor et al. 2010) which converts PC to

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phosphocholine and DAG (Fig. 6). DAG, in turn, is the lipid anchor from which DGTS and maybe SQDG biosyntheses are initiated as mentioned previously (LópezLara and Geiger 2016). During adaptation to low osmolarity conditions, S. meliloti synthesizes its version of periplasmic OPGs - denominated “cyclic β-1,2-glucans” which are synthesized by a nonhomologous set of proteins, NdvAB/ChvAB (Bontemps-Gallo and Lacroix 2015). Also, cyclic glucans can be substituted with sn-1-phosphoglycerol moieties derived from PG (Miller et al. 1988). The phosphoglycerol transferase CgmB of S. meliloti (Wang et al. 1999) catalyzes a reaction analogous to the sn-1-phosphoglycerol transferase OpgB from E. coli, as it converts neutral cyclic glucans into anionic cyclic glucans producing DAG as the second product (Fig. 6). One of the genes induced in early symbiosis is dgkA (Zhang and Cheng 2006), suggesting that in symbiotic conditions, DAG is phosphorylated to PA thereby re-entering the biosynthesis pathway for phospholipids (Fig. 6). Bacteria, such as S. meliloti, have at least two totally different life styles, one as a soil bacterium exposed to phosphorus limitation stress and another as a symbiotic bacterium in association with the legume host plant where, at least for the bacterium, phosphorus supplies seem to be abundant. Between the two different life styles of S. meliloti, membrane lipid composition is distinctly different (Geiger et al. 1999). In symbiotic conditions, the membrane is composed by phospholipids, whereas upon phosphorus limitation, phospholipids are largely replaced by membrane lipids that do not contain any phosphorus in their structure. Membrane lipid turnover in S. meliloti is not limited to the headgroups but involves acyl chains as well. In E. coli as well as in S. meliloti, mutants deficient in fadD (acyl-CoA synthetase) cannot consume free fatty acids and therefore accumulate them during the stationary phase of growth. Notably, these free fatty acids seem to be derived from bacterial membrane lipids (Pech-Canul et al. 2011). A major contributing activity for the release of these free fatty acids is the predicted patatinlike phospholipase A SMc01003 (Sahonero-Canavesi et al. 2015). Surprisingly, SMc01003 is not a phospholipase A but a DAG lipase which can degrade DAG to monoacylglycerol (MAG) and a fatty acid and then MAG further to another fatty acid and glycerol (Sahonero-Canavesi et al. 2015). Presently, it is not clear why the DAG lipase DglA is present in the Rhizobiaceae and what its physiological function might be. The OPGs of the soybean symbiont Bradyrhizobium japonicum are β-1,3;1,6linked cyclic glucans which can be substituted with phosphocholine residues (Rolin et al. 1992) and it is likely that this phosphocholine headgroup is derived from phosphatidylcholine.

3.4

Diacylglycerol Cycles in Gram-Positive Bacteria

Teichoic acids of gram-positive bacteria are important cell wall polymers of repeating phosphodiester-linked polyols and are either anchored to the membrane in the

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Fig. 6 Membrane lipid formation, turnover, and recycling in Sinorhizobium meliloti. The major membrane phospholipids of S. meliloti, PE, PC, PG, and CL, are formed by well-known pathways (Geiger et al. 2018) (for further details see text). CdsA:CDP-diacylglycerol synthase; Pcs: phosphatidylcholine synthase; PssA: phosphatidylserine synthase; Psd: phosphatidylserine decarboxylase; PmtA: phospholipid N-methyltransferase; PgsA: phosphatidylglycerol-phosphate synthase; ClsB: cardiolipin synthase, bacterial type; DgkA: diacylglycerol kinase; SqdB: UDP-sulfoquinovose synthase; BtaA: S-adenosylmethionine:diacylglycerol 3-amino-3carboxypropyl transferase; BtaB: diacylglyceryl homoserine N-methyltransferase; CgmB: cyclic glucan-modifying phosphoglycerol transferase; PlcP: phospholipase C; DglA: diacylglycerol lipase. Steps increased under phosphorus limitation (red) or at low osmolarity (blue) are highlighted. In symbiosis, dgkA is induced suggesting an increased recycling of DAG to phosphatidic acid and thereby a reintroduction of the DAG lipid anchor into phospholipid biosynthesis (purple). Also, the degradation of diacylglycerol to monoacylglycerol and further to glycerol is indicated (green). Reproduced from López-Lara and Geiger (2016) with permission from Elsevier

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case of lipoteichoic acid (LTA) or are bound to peptidoglycan in the case of wall teichoic acid (WTA) (see below). Although four distinct types of LTA are known to date (Percy and Gründling 2014), only LTA type I biosynthesis, found in Bacillus subtilis, Staphylococcus aureus, or Listeria monocytogenes, provides clear examples for the involvement of lipid cycles. Type I LTA has an unbranched 1,3-linked glycerol-phosphate backbone and is usually connected to the bacterial membrane via a glycolipid anchor (Fig. 7a). The hydroxyl groups at the C2 position of the glycerol-phosphate repeating units may be modified with D-alanyl or glycosyl groups (Fig. 7a). In S. aureus, the cytoplasmic glycosyltransferase YpfP successively transfers two glucose units from UDP-glucose substrates to the glycolipid anchor DAG converting it into diglucosyl-DAG and this lipid is then moved to the outside of the membrane, presumably by the flippase LtaA (Fig. 7b). A glycerol-phosphate chain is then formed on the glycolipid anchor by repeated addition of glycerolphosphate units derived from the headgroup of the membrane lipid PG by the action of the LTA synthase LtaS. The concomitantly formed DAG is recycled in the cytoplasm to PG in reactions catalyzed by DgkB, CdsA, PgsA, and PG phosphate phosphatase (Fig. 7b). The LTA molecule contains an average of 25 glycerolphosphate units, and as it is an abundant lipid in the outer leaflet of the membrane, one can estimate that the PG pool needs to be turned over more than twice per generation in order to support LTA synthesis. Like OPGs in proteobacteria, LTA are important for Firmicutes to resist low osmolarity conditions (Percy and Gründling 2014).

3.5

Emerging Amino Acyl-Phosphatidylglycerol Cycle

Not in E. coli, but in many other bacteria, the anionic lipid PG can be converted into the cationic lipid lysyl-PG (LPG) (Slavetinsky et al. 2016). In this reaction, lysine is transferred from a charged tRNALys to the glycerol headgroup of PG, as initially described with cell-free Staphylococcus aureus extracts. The mprF gene responsible for LPG formation was initially identified in S. aureus during a screen for mutants more susceptible than the wild type to CAMP. Remarkably, the C-terminal domain of MprF is sufficient for full-level LPG production at the inner leaflet of the cytoplasmic membrane, whereas the N-terminal MprF domain translocates LPG from the inner to the outer leaflet and therefore acts as a flippase (Slavetinsky et al. 2016). MprF homologues are also present in some gram-negative bacteria and are sometimes termed “low pH-inducible” gene encoding for LpiA (Vinuesa et al. 2003). LpiA homologues cause indeed the formation of LPG or alanyl-PG (APG). The lpiA genes are often in an operon with a downstream atvA gene conferring acid tolerance and virulence in some bacteria. AtvA is predicted to be lipase (Vinuesa et al. 2003) and apparently this protein can release the amino acid alanine from alanyl-PG and re-form PG (Arendt et al. 2013) thereby closing a lipid

Fig. 7 (a) Type I lipoteichoic acid structure as found in Bacillus subtilis and (b) scheme on PG turnover and DAG recycling during type I LTA biosynthesis in Staphylococcus aureus (for details see text). Blue arrows indicate transport of lipids across the membrane by flippases. DgkB: diacylglycerol kinase; CdsA: CDP-diglyceride synthase; PgsA: phosphatidylglycerol-phosphate synthase; ?: predicted phosphatidylglycerol-phosphate phosphatase (structural gene in grampositives not known to date)

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cycle. However, the question remains open whether such an energy-costly system is needed to “transport” alanine from the cytoplasm to the periplasm or whether in an in vivo context alanine rather would be transferred to another target molecule than water.

4

Lipid Cycles Involving Undecaprenyl Phosphate

In all three domains of life, Archaea, Eukarya, and Bacteria, long- and linear-chain polyprenyl-phosphate lipids are employed to facilitate the translocation of a sugar or glycan strand across biological membranes. They are involved in protein glycosylation as well as the biogenesis of bacterial cell-wall polysaccharides such as peptidoglycan, lipopolysaccharide O-antigen, wall teichoic acids, capsular polysaccharides, common enterobacterial antigen, OPGs, and exopolysaccharides (Manat et al. 2014). Synthesis of peptidoglycan precursors initiates in the cytoplasm where nucleotide-linked precursors UDP-N-acetylglucosamine and UDP-N-acetylmuramic acid pentapeptide are made (Fig. 8). The latter precursor is linked to undecaprenyl phosphate by MraY to generate lipid I. Then, MurG utilizes lipid I and UDP-Nacetylglucosamine to synthesize lipid II. In some bacteria, lipid II can be additionally modified by FemXAB and GatT. A lipid II flippase (possibly FtsW) translocates lipid II across the cytoplasmic membrane so that transglycosylase can polymerize the disaccharide-pentapeptide into glycan chains, liberating undecaprenyl pyrophosphate. In parallel, transpeptidases catalyze peptide bond formation between stem peptides in adjacent glycan chains. Phosphate transferases, such as LpxT, and phosphatases convert undecaprenyl pyrophosphate to undecaprenyl phosphate which is flipped back to the inner surface of the cytoplasmic membrane, ready to initiate another undecaprenyl phosphate-dependent lipid cycle (Fig. 8). Also for wall teichoic acid (WTA), at least four structurally distinct classes are known (van der Es et al. 2016) and biosynthesis has mainly been studied in type I WTA. The common linkage unit that connects the poly(alditol phosphate) chain to the peptidoglycan is a β-N-acetyl-mannosamine-(1,4)-α-N-acetyl-glucosamine-1phosphate disaccharide that is connected through a phosphodiester linkage to a muramic acid moiety in the peptidoglycan. The synthesis of this linker is performed on an undecaprenyl phosphate carrier by the consecutive action of TarO and TarA (Fig. 8). Next, a glycerol phosphate residue is transferred to the linker disaccharide by TarB using CDP-glycerol as the glycerol phosphate donor. In S. aureus, another glycerol phosphate moiety is attached by TarF, after which TarL generates the ribitol phosphate polymer using CDP-ribitol as a substrate. Finally, the poly(alditol phosphate) WTA chain is decorated with sugar residues by various enzymes (TarM, TarS, not shown) and subsequently the complete polymer is transported by the membranebound TarG/TarH complex to the external layer of the cytoplasmic membrane. The machine performing the transfer from the undecaprenyl support to peptidoglycan is not well understood but probably involves the enzymes TarTUV, producing undecaprenyl phosphate as the second product, which is flipped back to the inner surface of the cytoplasmic membrane, ready to initiate another undecaprenyl

Fig. 8 Undecaprenyl phosphate cycles for peptidoglycan and WTA biosynthesis in S. aureus (for details see text). Blue arrows indicate transport of lipids across the membrane by flippases

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phosphate-dependent lipid cycle (Fig. 8). Interestingly, the stereochemistry of the glycerol phosphate that is transferred during WTA synthesis (sn-glycerol-3-phosphate) differs from the stereochemistry of the glycerol phosphate building blocks in LTA synthesis (sn-glycerol-1-phosphate). An example worth mentioning is provided by type IV LTA biosynthesis in Streptococcus pneumoniae (Percy and Gründling 2014). Remarkably, type IV LTA from Streptococcus pneumoniae can be substituted with phosphocholine residues. In this case, the phosphocholine residues are transferred from CDP-choline by LicD1 and LicD2 to the O-6 position of N-acetyl-galactosamine (GalNAc) residues still within the cytoplasmic compartment.

5

Physiological Implications of Membrane Lipid Turnover and Lipid Cycles

Replacement of phospholipids by phosphorus-free membrane lipids, under phosphorus-limiting conditions of growth (Zavaleta-Pastor et al. 2010), is not limited to S. meliloti in soils lacking phosphorus, or to laboratory studies. Upon phosphorus deficiency, lipid remodeling is a widespread strategy in marine heterotrophic bacteria (α-proteobacteria, Flavobacteria) employing phospholipase C PlcP (Sebastián et al. 2016). Also, bacteria of the SAR11 clade renovate their lipids in response to phosphate starvation (Carini et al. 2015). Other environmental stresses, such as low or high pH, temperature, medium osmolarity, etc., provoke changes in the lipid composition of individual bacteria as well (Sohlenkamp and Geiger 2016), emphasizing that potential lipid diversity is enormous even in individual bacterial cells. Obviously all differentiation processes occurring in bacteria, i.e., endospore formation in Bacillus or Clostridium species or fruiting body and spore formation in Myxobacteria, are expected to involve intensive membrane remodeling. Upon formation of desiccation-resistant cysts by Azotobacter vinelandii, it is even known that membrane phospholipids are largely replaced by phenolic lipids (López-Lara and Geiger 2016). However, it is unknown whether upon dedifferentiation, i.e., upon germination of the spore/cyst and their development into a vegetative cell, there is a closure of lipid cycles in the form of an active degradation of special spore/cystassociated lipids.

6

Conclusion and Research Needs

In the case of lipid cycles, as with any metabolic cycle in biochemistry, the question arises what they are good for and obviously many of them are important for biosynthesis purposes outside the cytoplasmic compartment. Cycles with no net yield would go around in circles, just consuming energy and therefore would be futile cycles. However, if no net compound is synthesized, it still might be important to drive forward one half of the cycle under a certain physiological condition and the

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other half under another condition, like, for example, the DAG/PA cycle in yeast or the lipid cycles observed in S. meliloti, which are driven by stress (phosphate limitation or low osmotic conditions) into one direction (toward DAG) and by normal, replete conditions back to PLs. Although membrane lipid composition has long been considered to be a near constant for a given cell, this idea needs to be modified in the face of contrasting findings. The rates of de novo biosynthesis of the distinct membrane lipids certainly affect membrane lipid composition in an important way. However, membrane lipid turnover, degradation, and recycling are other major processes that impact membrane lipid composition. Plants and bacteria cannot easily move away from the given macroenvironment and have to adapt their physiology to the existing environmental conditions which might even change considerably over time. In a given physiological situation, contributing vectors for synthesis and turnover of lipids might remain pretty much constant which should lead to a certain steady-state condition with a definable membrane lipid composition. However, a distinct physiological condition might alter synthesis or turnover or both resulting in a different steady-state condition with a different membrane lipid composition. Therefore, in plants and environmental microbes, extensive remodeling of membrane lipids can be expected. Adaptation to distinct stress conditions may go along with partial degradation of PLs and the formation of different membrane lipids. For example, upon phosphoruslimiting conditions, phosphorus-free membrane lipids (SL, OL, DGTS, and glycolipids) are formed and largely replace PLs. It is unclear to date, to what extent the lipid status is reversed when an organism is cultivated in culture media with abundant phosphorus again and whether SL, OL, DGTS, or glycolipids are actively degraded in high-phosphate conditions. Membrane lipid turnover in bacteria is a scarcely studied area, as there are considerable challenges in facile assignments of physiological substrates/products for intrinsic phospholipase or transferases that remove groups from membrane lipids. Defining substrate specificities and therefore the function of a newly discovered enzyme is often a multivariable problem and if no activity is detected, it is often not clear whether the assay condition are inadequate for the enzyme to work or whether the correct combinations of substrates were not provided. In some cases, it might be advisable to separate the optimization of enzyme activity from the search for the physiological substrate for the enzyme (Sahonero-Canavesi et al. 2016). Another important role for lipid cycles might consist in the fact that they enable biosynthesis pathways occurring outside the cytoplasm that otherwise would not be possible. High-energy compounds, which are able to release a large amount of free energy upon hydrolysis (Nelson and Cox 2017), i.e., anhydrides or thioesters, are usually limited to the cytosol of a cell. Therefore, within the cytosol, the transfer of a phosphoalcohol from an anhydride to an ester linkage is energetically favorable, ensuring that in such transferase reactions, most substrates are converted into products. An example is the transfer of phosphoglycerol from CDP-glycerol to a glycolipid during WTA synthesis. However, outside the cytoplasmic compartment, phosphoalcohol transferases usually remove an ester-linked phosphoalcohols from

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PLs in order to generate new ester-linked phosphoalcohols on target molecules such as OPGs, LTAs, or lipid A. The rule that high-energy compounds are used by transferases as substrates in the cytoplasm and low-energy compounds outside the cytoplasmic compartment applies in a similar way to acyl transferases. Acyl transferases acting in the cytoplasmic compartment usually use activated acyl derivatives (CoA or ACP thioesters, or acyl-phosphates), whereas acyl transferases acting outside the cytoplasmic compartment transfer acyl residues from PLs to other target molecules such as lipid A or prolipoprotein Lpp. The exception to this rule may be encountered in undecaprenyl cycles when undecaprenyl pyrophosphate derivatives are flipped across the cytoplasmic membrane and now provide a high energy compound due to the anhydride linkage in the pyrophosphate residue in a compartment outside the cytoplasm. Bacteria enjoy an infinite capacity for reproduction as long as they reside in an environment supporting growth. However, their rapid growth and efficient metabolism results in depletion of substrates and the cell population enters the stationary phase of growth in which cells are gradually degenerating. In stationary phase, a reduction of cell size can be observed, generally termed as “dwarfing,” and in parallel an active degradation of membrane lipids and fatty acids (Nyström 2004). However, the physiological relevance or the molecular mechanisms of these processes are not understood. Although membrane lipid degradation and cycles are evident in Eukarya and Bacteria, little is known on such processes in Archaea. It is obvious that protein glycosylation in Archaea requires the polyprenol dolichol and therefore it is likely that polyprenol-based lipid cycles must exist in Archaea (Meyer and Albers 2013). Archaeal membrane lipids show characteristic ether linkages between glycerol and isoprenol residues which are much more difficult to cleave than bacterial lipids in which fatty acyl residues are ester-linked to glycerol. Presently, it is unknown whether archaeal membrane-forming lipids are turned over or whether they might be members of lipid cycles. Acknowledgement Research in our lab was supported by grants from Consejo Nacional de Ciencia y Tecnología-México (CONACyT-Mexico) (178359 and 253549 in Investigación Científica Básica as well as 118 in Investigación en Fronteras de la Ciencia) and from Dirección General de Asuntos del Personal Académico-Universidad Nacional Autónoma de México (DGAPA-UNAM; PAPIIT IN202616, IN203612). We thank Lourdes Martínez-Aguilar for skillful technical assistance.

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Filip Kovacic, Nikolina Babic, Ulrich Krauss, and Karl-Erich Jaeger

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Structure–Function Relationships of Lipolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Family I: Triacylglycerol Lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Family II . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Family III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Family IV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Family V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Family VI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Family VII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Family VIII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Family IX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Family X . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Family XI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Family XII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13 Family XIII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.14 Family XIV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.15 Family XV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.16 Family XVI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.17 Family XVII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.18 Family XVIII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.19 Family XIX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Conclusions and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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F. Kovacic · N. Babic · U. Krauss Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Jülich, Germany e-mail: [email protected]; [email protected]; [email protected] K.-E. Jaeger (*) Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Jülich, Germany Institute of Bio- and Geosciences IBG-1: Biotechnology, Forschungszentrum Jülich GmbH, Jülich, Germany e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_39

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Abstract

Lipolytic enzymes including lipases and esterases comprise a versatile group of enzymes with diverse amino acid sequences but related three-dimensional structures. Despite the large number of bacterial lipolytic enzymes so far identified (~5000), only a small portion (C12) fatty acid esters; therefore, it can be classified as a typical esterase. The optimal temperature for EstA3-catalyzed ester hydrolysis at 70  C is consistent with the optimal temperature for growth of T. tengcongensis.

2.15

Family XV

Family XV of bacterial lipolytic enzymes contains the esterase EstGtA2 from Geobacillus thermodenitrificans (Charbonneau et al. 2010), the lipase LipS from a metagenome (Chow et al. 2012), and the monoacylglycerol lipase bMGL from Bacillus sp. H257 (Imamura and Kitaura 2000; Kitaura et al. 2001) with molecular weights ranging from 27 kDa to 30 kDa. Although LipS was identified by screening of a metagenomic library, its amino acid sequence is identical with the putative esterase from the thermophilic bacterium Symbiobacterium thermophilum. The intracellular lipase bMGL from the moderately thermophilic bacterium is homologs to esterases from Streptomyces coelicolor and B. stearothermophilus, yet it shows homology with human monoacylglycerol lipase, too (Imamura and Kitaura 2000; Kitaura et al. 2001). EstGtA2 shares 89% and 51% sequence identity with bMGL and LipS, respectively (Charbonneau and Beauregard 2013). As expected for enzymes isolated from thermophilic organisms, they were stable at temperatures

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above 60  C, and they optimally hydrolyzed the substrates at 70  C with the exception of EstGtA2 that has temperature optimum at 50  C (Kitaura et al. 2001; Charbonneau et al. 2010; Chow et al. 2012). Members of this family have a conserved SDH catalytic triad with a GLSMGG motif around the active-site serine. Despite similarity of ~30% of amino acid sequences among lipolytic enzymes from families XV and XIII, their threedimensional structures revealed important structural differences distinguishing these two families (Charbonneau and Beauregard 2013). Thus, the three α-helical cap structure of family XIII enzyme Est30 differs from the caps composed of an α-helix and two antiparallel β-strands identified in the X-ray structures of bMGL (PDB code: 3RM3) (Rengachari et al. 2012) and LipS (PDB code: 4FBL) (Chow et al. 2012). Furthermore, the structures of enzymes from families XIII and XV revealed salt bridges as hallmarks of these families. Of particular interest is the strongly conserved salt bridge which tethers the loops carrying the catalytic Asp and His residues. This interloop salt bridge is formed by residues located behind the catalytic Asp and His at positions 2 and 4, respectively. In most of the family XV enzymes, a negatively charged residue is conserved at position Asp-2 and a positively charged residue at position His-4, while the opposite organization was observed for family XIII enzymes (Charbonneau and Beauregard 2013). Despite similar sequences and structures, the substrate profiles of EstGTA2, bMGL, and LipS distinguish substantially from each other. While bMGL is a typical monoacylglycerol lipase without activity toward di- and triacylglycerols, EstGtA2 can hydrolyze triacylglycerides; acylglycerides were not tested as substrates for LipS. However, LipS showed a broad substrate specificity with a range of industrially relevant substrates, among them (R)-ibuprofen-phenyl ester that was hydrolyzed with an enantiomeric excess (ee) of 99%. All three enzymes showed specificity for p-nitrophenyl esters with fatty acids up to 8 carbon atoms; however, they showed activities with long-chain fatty acid esters, too (Imamura and Kitaura 2000; Kitaura et al. 2001; Charbonneau et al. 2010; Chow et al. 2012). Apparently, these enzymes are intracellular lipases; therefore, their catalytic activity is presumably related to a specific function in each source organism what might be an explanation for dissimilar substrate specificity.

2.16

Family XVI

Family XVI of lipolytic enzymes contains one experimentally studied enzyme, the cold-active alkaline lipase LipSM54 from Stenotrophomonas maltophilia (Li et al. 2016). This lipase with a molecular weight of 55 kDa is homologs to putative lipases from the genera Stenotrophomona, Xanthomonas, Saccharopolyspora, Saccharothrix, Actinosynnema, and Streptomyces. A sequence alignment of family XVI lipolytic enzymes revealed a conserved GISYGAG motif around catalytic Ser, as well as conservation of the catalytic triad residues Asp and His, that was proven to be essential for the catalysis by mutation. LipSM54 hydrolyzes p-NP and glycerol fatty acid esters with similar activities and specificities for C8 and C10 fatty acid

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esters, although esters with acyl chains up to 18 carbons were hydrolyzed by LipSM54, too. The optimal temperature for hydrolysis of p-NP esters by LipSM54 was 35  C, although the enzyme was highly active also at 5  C. Apparently, LipSM54 can be classified as a cold-active enzyme with low stability at temperatures above 40  C (Wu et al. 2015) and high activity at low temperatures. Characterization of other LipSM54 homologs will reveal if this is a general property of family XVI lipolytic enzymes.

2.17

Family XVII

The extracellular lipase LipJ2 from the halotolerant bacterium Janibacter sp. strain R02 with a molecular weight of 44 kDa was first identified as a member of family XVII of lipolytic enzymes homologs to hypothetical or putative lipases from Arsenicicoccus, Dermatophilus, and Janibacter genera (Castilla et al. 2017). These proteins, although classified as novel family XVII, show homology with members from family X. A sequence alignment of family XVII members revealed the presence of a SHD catalytic triad and a conserved GYSQG pentapeptide around the catalytic serine. The three-dimensional structures of LipJ2 and other members of family XVII are not known, but Q and Y were predicted as residues involved in forming the oxyanion hole (Castilla et al. 2017). Such a Y-type oxyanion hole with Y as the residue stabilizing the tetrahedral intermediate is untypical for bacterial lipases, but was found in the C. antarctica lipase A and several other fungal lipases. Biochemical characterization of LipJ2 showed preference for esters containing short- to medium (C4–C12)-chain fatty acids, while activity with the long-chain (>C16) substrates was significantly lower. LipJ2 shows thermophilic properties, optimal activity at 80  C, and no loss of activity after incubation for 1 h at a temperature close to 100  C. LipJ2 is stable under harsh pH conditions, as it retains more than 70% of activity even at pH 2, and no loss of activity was observed under alkaline conditions (up to pH 10). Interestingly, LipJ2 is strongly activated by a mixture of Na+ and K+ions (Castilla et al. 2017).

2.18

Family XVIII

Esterase EstUT1 with a molecular weight of 28 kDa from the thermophilic bacterium Ureibacillus thermosphaericus was classified into the new family XVIII of lipolytic enzymes (Samoylova et al. 2018). The closest homologs are putative esterases from Lysinibacillus, Bacillus, and Solibacillus genera. EstUT1 shares approximately 40% sequence identity with family VIII of the bacterial lipases, but the conserved pentapeptide motifs around the catalytic serine differ with GVSLG for family XVIII and GGSVG for family VIII. The typical SHD catalytic triad and an α/β-hydrolase fold were predicted for EstUT1. EstUT1 is highly specific for p-NP esters of short-chain fatty acids (C2, C4) with the highest activity for p-NP-acetate (C2) and almost no activity with esters containing C12 or longer fatty acids. EstUT1

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is thermostable with a temperature optimum between 70  C and 80  C, where it shows high stability with ~70% activity retained after incubation for 6 h at 70  C. Notably, this enzyme also exhibits high stability in many organic solvents even after incubation for 6 h at a concentration of 30% (v/v) and 50  C.

2.19

Family XIX

The discovery of the thermostable lipase LipSm from Stenotrophomonas maltophilia with a molecular weight of 42 kDa resulted in the definition of family XIX of lipolytic enzymes (Parapouli et al. 2018). LipSm shows >48% sequence identity with several uncharacterized putative lipolytic enzymes from Pseudomonas geniculata and P. aeruginosa, Stenotrophomonas pavanii, Gordonia sp., and Rhodococcus sp. The sequence alignment revealed a typical SHD catalytic triad with a GHSQGGA peptide surrounding the catalytic serine. Similar to family XVII LipJ2, residues Q and Y were predicted for LipSm as being involved in forming a Y-type oxyanion hole. Moreover, several AA motifs possibly related to the thermostability of LipSm were conserved among the members of family XIX. The optimal temperature for hydrolysis of p-NP esters by LipSm was 64  C, the enzyme was fully stable at 70  C for 30 min, and optimal pH conditions for catalysis were between pH 8 and 9.

3

Conclusions and Research Needs

In the preceding version of this chapter published in 2010, we reported a number of about 900 lipolytic enzymes originating from bacteria (Hausmann and Jaeger 2010). Since then, this number steadily increased now exceeding 5000 identified enzymes. Notably, less than 10% of them have been biochemically characterized, whereas more than 90% were identified only by sequence similarity. Following the initial classification published by Arpigny and Jaeger (1999), we have grouped these bacterial lipolytic enzymes into 19 families based on amino acid sequence, structure, and function. This classification should allow predicting functions and biochemical properties of enzymes which have newly been identified in databases by sequence similarity. Our results show that the overall amino acid sequences of bacterial lipolytic enzymes are very diverse; however, the conserved pentapeptide sequence surrounding the catalytic serine can efficiently be used to distinguish 19 families (Table 3) as suggested previously (Zarafeta et al. 2016). Small variations of the canonical GXSXG motif seem to provide a phylogenetic signature of lipases (Fig. 3). Apparently, non-catalytic residues contribute significantly to substrate specificity, protein activity, and stability by still unknown mechanisms; therefore, more structural data are needed to elucidate the structure–function relationships of lipolytic enzymes. Furthermore, many of the newly identified lipolytic enzyme families contain lipolytic enzymes originating from extremophilic microorganisms; consequently, these

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Table 3 Conserved sequence motifs in bacterial lipolytic enzymes around the catalytic serine Family I II III IV V VI VII VIII IX X XI XII XIII XIV XV XVI XVII XVIII XIX a

Representative enzymea PaLip SsEst SeLip AaEst Est2 PfEst PnbA EstB PhaZ7 EstD LipG LipEH166 Est30 EstA3 LipS LipSM54 LipJ2 EstUT1 LipSm

Consensus sequence GHSQG GDSL GXSMG GDSAGG GXSMGG GFSQG GESAG GGSVG AHSMG GHSLG GHSLGG GHSLG GLSLGG CHSMG GHSAG GISYGAG GYSQG GVSLG GHSQGGA

References for all enzymes are included in the text

enzymes were adapted to be active and stable at conditions optimal for growth of these organisms. Although hot- and cold-active lipolytic enzymes were recognized in families XV and XVI, respectively, more members from each family need to be biochemically characterized to assign extremophilic properties to the entire family. Lipolytic enzymes are ubiquitously distributed and have important cellular functions that are still unknown for most of them. The systematic analysis and classification of novel lipolytic enzymes provided within this chapter will help in understanding their biochemical and physiological functions. Acknowledgments NB is a recipient of a PhD grant from the Manchot Graduate School “Molecules of Infection” at Heinrich Heine University Düsseldorf, Germany.

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Pathways for the Degradation of Fatty Acids in Bacteria

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Lorena Jimenez-Diaz, Antonio Caballero, and Ana Segura

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Transportation of Exogenous Long- and Medium-Chain Fatty Acids . . . . . . . . . . . . . . . . . . . . . 3 Aerobic Degradation of Long- and Medium-Chain Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Acyl-CoA Synthetase (FadD) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Acyl-CoA Dehydrogenases (FadE) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Fatty Acid β-Oxidation Multienzyme Complex (FadAB) . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Aerobic Degradation of Acetoacetate and Short-Chain Fatty Acids . . . . . . . . . . . . . . . . . 3.5 Degradation of Unsaturated Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Anaerobic Degradation of Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7 The Glyoxylate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Biotechnological Applications of Fatty Acid Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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L. Jimenez-Diaz (*) Abengoa Research, Sevilla, Spain e-mail: [email protected] A. Caballero Abengoa Research, Sevilla, Spain Bacmine, Tres Cantos, Spain e-mail: [email protected] A. Segura Abengoa Research, Sevilla, Spain Department of Environmental Protection, Consejo Superior de Investigaciones Científicas, Estación Experimental del Zaidín, Granada, Spain e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_42

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Abstract

The metabolism of fatty acids is of central importance to wide range of microbes, and the related metabolic pathways have been intensely studied for decades. Fatty acid degradation occurs through the well-characterized β-oxidation cycle and yields acetyl-coenzyme A (CoA), which is further metabolized to obtain energy and precursors for cellular biosynthesis. As demand for sustainable biofuels and bioplastics grows, there is ever-increasing interest in fatty acid flux and related genomic bacterial diversity, which is opening avenues to exciting new biotechnological applications. In this chapter, we describe the steps involved in bacterial fatty acid degradation, with an emphasis on the latest advancements in the determination of enzymatic structures and characterization of novel fatty acid degradation enzymes. Finally, we briefly discuss the relevance of fatty acid degradation to several industrial applications.

1

Introduction

Fatty acids (FAs) are simple molecules that are composed of a carboxylic acid with a long aliphatic chain that is either saturated or unsaturated (IUPAC 1997). They are present in the membrane structures of all organisms and play essential roles in membrane architecture, homeostasis, and transport. FAs also constitute important sources of metabolic energy, while various derivative molecules are effectors that regulate several cellular processes. In bacteria, FAs are frequently found as components of phospholipids; two FAs are attached to a glycerol moiety to form a diacylglyceride, while the third carbon of the glycerol molecule is attached to a polar head group. Phospholipids in the membranes are constantly being synthesized, modified, and destroyed to maintain membrane homeostasis and to respond to environmental stressors. Free FAs are released during these processes, constituting important sources of metabolic energy. Bacteria can also take up external FAs that are transported across the membrane, activated to acyl-CoAs, and degraded. FA degradation occurs through the β-oxidation cycle, which yields acetylcoenzyme A (CoA) that is further metabolized to obtain energy and precursors for cellular biosynthesis. Because of the central role that FAs have in the cell, fatty acid degradation (FAD) has been exhaustively studied in several organisms (Schultz 1991), and the molecular mechanisms of FAD in Escherichia coli or Salmonella enterica have been well established (Nunn 1996; Heath et al. 2002). Although the E. coli FAD pathway was considered paradigmatic, genomic studies have revealed that there exists a wide variety of unique genes and enzymes that participate in FA metabolism in other microorganisms. For example, during long-chain FA degradation (i.e., growth on oleate), unlike S. enterica, where there is no accumulation of intermediates, E. coli accumulates short- and medium-chain acyl-CoA (Iram and Cronan 2006). This strategy is metabolically expensive; each acyl chain requires

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activation by ATP hydrolysis in order to enter the initial cycle of β-oxidation, whereas subsequent cycles require no additional activation. While S. enterica is able to obtain all the acetyl-CoA from long- and medium-chain FAs, E. coli extracts less acetyl-CoA equivalents. It has been suggested that the higher efficiency of the FadBA and FadE proteins of S. enterica account, at least partially, for these differences, although the amino acid sequence of the proteins in the two organisms shares almost 91% identity, and both strains occupy similar ecological niches (Iram and Cronan 2006). With the advent of the new sequencing technologies, information about genes participating in the degradation of FAs in other bacteria have been made available, and although the genetic determinants of the FA degradation in most of these sequenced microorganisms have not yet been studied in detail, a wide diversity in genes and substrate specificities can be inferred. Over the last 20 years, increased interest in the biotechnological synthesis of added-value compounds, mainly biofuels and bioplastics, has driven research on bacterial FA metabolism, revealing a wealth of information about the genetics, biochemistry, and bottlenecks of FA degradation in a variety of microorganisms. Despite the genetic diversity observed between different microorganisms, basic pathway components are required for growth on FAs. These include a specific transport system (for long-chain FAs) and the enzymes involved in FA degradation: an acyl-CoA synthethase and components of the β-oxidation cycle and glyoxylate shunt. In this chapter, we describe each of the microbial-based steps of FA degradation – from their transportation across the membrane to the degradation to acetyl-CoA, with emphasis on the most current advancements in the field. These advancements include new insights into the differences that exist in FA metabolism between different bacteria, as well as recently resolved structures of key enzymes, which have facilitated a deeper understanding of mechanisms of action and substrate specificity. Finally we will frame this knowledge within the context of several industrial applications of FA degradation.

2

Transportation of Exogenous Long- and Medium-Chain Fatty Acids

FA degradation can occur when FAs are released from membrane phospholipids or when free FAs are taken into the cell from the environment. While long-chain FAs (LCFAs, C18-C12) require facilitated transport to enter the cells, medium-chain FAs (C7-C11) can enter the cell by free diffusion or by an still unknown mechanism (Nunn et al. 1979; Maloy et al. 1981; Black and DiRusso 1994). Also, it has recently been proposed that the facilitated transport of short-chain fatty acids (SCFA, C6-C4) may be mediated by porin OmpF (Rodriguez-Moyá and Gonzalez 2015). FA transport into the cell is tightly linked to FA activation, which involves formation of a fatty acyl-CoA ester through a process named vectorial esterification (because of analogies with vectorial phosphorylation). The transportation of LCFAs into E. coli requires the long-chain fatty acid transport protein (FadL), which is located in the outer membrane and acts as a receptor of LCFAs, as well as the acyl-

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CoA synthetase (FadD), located in the inner membrane and the cytosol. FadD activates LCFAs by converting them into long-chain acyl-CoA thioesters. These thioesters subsequently activate the FadR regulator, which then represses expression of some FA biosynthesis genes and activates expression FA degradation genes (DiRusso and Black 2004). FadL specifically binds FAs (Black 1990) and does so with high affinity for LCFAs, displaying KT values of 4.1
105 M for myristate (C14:0), 6.7
105 M for palmitate (C16:0) and 13.2
105 M for oleate (C18:0) (Maloy et al. 1981). Furthermore, FadL is essential to the active transport of myristic (C14:0) and oleic (C18:1) acids (Nunn and Simons 1978; Maloy et al. 1981; Black 1990) in E. coli. It should be noted that the requirement for FadL does not hold for the E. coli strain MG1655, which is devoid of FadL (BW25113ΔfadL) but still able to produce C14-AFA (14-azidotetradecanoic acid) from externally provided C16-AFA (16-azidohexadecanoic acid), albeit with delayed production versus wild type (Perez and Bode 2015). Authors of the study suggested that this delay could be due to the absence of FadL, which would be expected to reduce FA membrane diffusion rates. Additionally, although FadL is not absolutely required for MCFA transport, the possibility exists that it may have overlapping specificity for MCFAs. The elucidated crystal structure of the E. coli FadL has provided insight into its mechanism of action (van den Berg et al. 2004). The current proposed transport mechanism starts when FA binds FadL at a hydrophobic groove between two extracellular loops; this binding results in high local concentrations of FAs and diffusion of the substrate into the high-affinity binding pocket. This binding provokes a conformational change at the N-terminal domain of the protein lowering the affinity for the substrate while opening the initial part of the channel that, with additional structural changes, opens the final part of the channel and facilitates the movement of the FA into the periplasm (van den Berg et al. 2004; Lepore et al. 2011). Given the low pH of the intermembrane space of Gram-negative bacteria, the carboxylate group of the FA is likely protonated; these uncharged periplasmic FAs can then flip into the cytosolic face of the membrane (Hamilton 2003). A variety of Gram-negative bacteria have been shown to have FadL homologues, for which some have been shown to participate in FA transport. Interestingly, a number of FadL-like proteins have also been shown to be involved in xenobiotic and alkane uptake (van den Berg et al. 2004; Grant et al. 2014; Call et al. 2016) or in the secretion of biologically active metabolites (Martínez et al. 2013). The second step in LCFA transport is carried out by FadD, which facilitates the ATP-dependent active transport of FAs from the periplasm across the inner membrane into the cytosol while, at the same time, activating the FA by adding a coenzyme A molecule to form a fatty acyl-CoA ester (Klein et al. 1971; Fig. 1). Due to the coupling between the transport and the activation of LCFAs, 98% of the transported fatty acids are in the form of fatty acyl-CoA thioesters in the cytoplasm, and they are mainly broken through the β-oxidation pathway (although they can also serve as substrates in phospholipid synthesis), while only 2% are converted to fatty acyl-ACPs (through a fatty acyl-ACP synthase) that are latter bound to phosphatidylethanolamine (Rock and Jackowski 1985).

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Long-chain fatty acid

Outer membrane

FadL

Periplasm

Inner membrane

FadD Fatty acid + ATP = fatty acyl-AMP + PPi Fatty acyl-AMP + CoA = fatty acyl-CoA + AMP ATP+CoA

AMP+PPi

FAD

Acetyl-CoA FADH2

CoASH

FadA

Fatty acyl-CoA

FadE

Iterative cycles

2-enoyl-CoA

3-ketoacyl-CoA

FadB

FadB

H2O

NADH NAD+

3-hydroxyacyl-CoA

Fig. 1 Schematic representation of the fatty acid degradation pathway. Each step in the β-oxidation cycle shortens the fatty acid molecule by two carbons

3

Aerobic Degradation of Long- and Medium-Chain Fatty Acids

3.1

Acyl-CoA Synthetase (FadD)

FadD, a fatty acyl coenzyme A (CoA) synthetase (FACS; fatty acid CoA ligase [AMP forming] [EC 6.2.1.3]), catalyzes the activation of the FAs that have been transported from the external media and also those that are cleaved from the

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cytoplasmatic membrane (Pech-Canul et al. 2011), converting them into fatty acylCoAs in a two-step reaction. The first step proceeds through the pyrophosphorolysis of ATP, where the carboxyl group of the FA is linked by an acyl bond to the phosphoryl group of AMP, while liberating a pyrophosphate from ATP (Fig. 1). In the second step, the fatty acyl moiety is transferred to the sulfhydryl group of coenzyme A with the release of AMP (Groot et al. 1976; Weimar et al. 2002). FadD was initially proposed to be soluble and loosely bound to the inner membrane (Samuel et al. 1970; Kameda and Nunn 1981), although a later model suggested that FadD moves between the cytosol and the inner membrane and is possibly only active in the membrane and remains inactive in the cytosol. It has also been suggested that the degree of membrane association depends on the state of energization of the cell membrane (Mangroo and Gerber 1993). Although E. coli FadD is active in vitro on FAs from C6 to C18, it has more specificity for LCFAs. Purified E. coli FadD has a Km and Vmax for dodecanoic acid (C12:0) of 1.6 μM and 2632 nmol/min/mg protein, respectively, while the Km and Vmax for hexanoic acid (C6:0) were 11.2 μM and 135 nmol/min/mg/protein (Kameda and Nunn 1981). The mechanisms behind FadD substrate specificity are not clearly understood, although the solution of the Thermus thermophilus FadD 3D structure has helped in this understanding (Hisanaga et al. 2004). The protein is a homodimer with each monomer consisting of a large N-terminal domain that contains a FA-binding tunnel which extends to the ATP-binding site located in the small C-terminal domain. The C-terminal domain presents a closed or open conformation depending on the presence of ligand. The FA-binding tunnel is formed by two paths (the “ATP path” and the “center path”) and pocket named “dead-end branch.” Upon ATP binding, the gated tunnel opens at the same time that the C-terminal domain adopts the closed conformation. The center path is the entrance for the FA, and the acyl chain of the FA-AMP enters into the dead-end branch. In the absence of ATP, the connection between the ATP path, the center path, and the dead-end branch is blocked, and the FAs cannot access the active site (Hisanaga et al. 2004). This structure and mechanism explains partially why strategies to obtain FadD mutants with an increased substrate range resulted in mutations that did not map in the active site but in a motif adjacent to a region of FadD involved in FA binding or in the C-terminal domain (Black et al. 1997; Ford and Way 2015). It was hypothesized that the depth of the tunnel (dead-end branch) determined the length of the acyl chain that FadD could accommodate, and the techniques used in these experiments made it difficult to modify this architecture. The C-terminal domain is responsible for the ATP binding and the release of reaction products (AMP and fatty-acyl-CoA). It is improbable that the bulky adenylate moieties in the resulting reaction products can pass back through the FA-binding tunnel to the central valley. Therefore, it is proposed that they are released from the ATP-binding site after opening of the C-terminal domain (Hisanaga et al. 2004). Modification of this domain could improve the affinity of FadD for FAs, although since no decrease in the Km of the enzyme for octanoate was detected while an increase in Vmax was, it was suggested that the liberation of AMP was a determinant for the improved growth of mutants in MCFAs (Ford and Way 2015).

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When the concentration of intracellular fatty-acyl-CoA thioesters is high, FadR, the transcriptional regulator that controls FA degradation pathways, derepresses the FA degradative genes ( fad). In E. coli, this derepression occurs only when long-chain fatty acyl-CoA are accumulated and not in the presence of MCFAs thioesters (Dirusso and Nunn 1985). Therefore, E. coli is unable to grow with octanoate (C8:0) and decanoate (C10:0) as the sole carbon source. In contrast, many other bacteria in which FAs of C6 or longer are effective inducers of the β-oxidation system are able to grow on FAs of different chain length (Sato et al. 1992; Hume et al. 2009; Zarzycki-Siek et al. 2013). Although based on genome sequence comparison, Salmonella enterica serovar Typhimurium LT2 is similar to that of E. coli K-12 they are functionally different. One of the differences is the ability of S. enterica to grow in MCFAs such as decanoic acid (C10:0). This growth can be attributed to higher levels of acyl-CoA pools in S. enterica than in E. coli. Enzymatic FadD activity in both strains is similar, so the differences in acyl-CoA pools could be due to differences in the fadD gene expression (Iram and Cronan 2006). In fact, a derivative E. coli strain overexpressing fadD was able to grow with these medium-chain FAs. The suggestion is that higher levels of acyl-CoA synthetase produce higher levels of MCFA thioesters that likely remove the inhibition that FadR exerts over the FA β-oxidation genes (Zhang et al. 2006). As in Salmonella, in Pseudomonas putida CA-3, there is a unique copy of an acyl-CoA synthetase which is essential for the metabolism and activation of FAs. This enzyme shows activity toward long-chain aromatic and aliphatic substrates as well as toward long-, medium-, and short-chain phenylalkanoic and alkanoic acids, although it exhibits a gradual increase in its activity as the chain length of the substrate increases (Hume et al. 2009). In E. coli there is only one copy of fadD, while in other organisms, such as the strain P. putida U, there are two copies, fadD1 and fadD2. FadD1 appears to perform the main activity while FadD2 is either expressed only when FadD1 is absent or it activates different substrates from FA (aromatic or aliphatic) (Olivera et al. 2001). Similarly, the strain P. putida Gpo1 has two identified acyl-CoA synthetases (ACS1 and ACS2); ACS1 is associated with polyhydroxyalkanoates (PHA) granules and is probably involved in PHA mobilization, while ACS2 is mainly located in the cell membrane (Ruth et al. 2008). Furthermore, in addition to fadD and fadD2, the strain P. putida KT2440 presents a third fadD gene ( fadDx), the function of which has not been investigated (Wang and Nomura 2010). Once the FAs have been activated, their degradation proceeds through the β-oxidation cycle. In each cycle, one molecule of fatty acyl-CoA loses two carbons in the form of acetyl-CoA, with the reduction of one molecule of flavin adenine dinucleotide (FAD) and one molecule of nicotinamide adenine dinucleotide (NAD). The released acetyl-CoA is further metabolized in the glyoxylate cycle, and the rest of the fatty acyl-CoA continues in the β-oxidation cycle without the need to be activated again (Clark and Cronan 2005) (Fig. 1). Only three enzymes are required to complete the β-oxidation cycle: an acyl-CoA dehydrogenase (encoded in E. coli by fadE), a 3-hydroxyacyl-CoA dehydrogenase/enoyl-CoA hydratase (encoded by fadB), and a β-ketothiolase (encoded by fadA). FAs with an unsaturation at the even-numbered carbon require an additional auxiliary enzyme, the dienoyl-CoA reductase (encoded by fadH).

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Acyl-CoA Dehydrogenases (FadE)

The acyl-CoA dehydrogenases (ACADs) catalyze the α, β-dehydrogenation of acyl-CoA esters in the catabolism of FAs and amino acids. The origin of the ACAD family can be traced to the common ancestor of Archaea, Bacteria, and Eukaryota, suggesting its essential role in metabolism of early life (Swigonová et al. 2009). ACAD subfamilies have been identified on the bases of the metabolic pathways in which they participate (β-oxidation of FAs or amino acid degradation) and by their substrate specificity. Those that participate in the β-oxidation pathway can be grouped into seven subfamilies depending on the substrate with which they display the optimal activity, short (ACADS), medium (ACADM), long (ACADL), or very long (ACADV and ACADV2) chain length FAs, plus two additional subfamilies reported in bacteria: FadE that degrades from short to long-chain acyl-CoAs, and FadE12 that prefers medium-chain length molecules (Shen et al. 2009). FadE is responsible for the oxidation of acyl-CoA to 2-enoyl-CoA, a step that involves the transfer of two electrons to the FAD cofactor, which is reduced to FADH2 (Black and DiRusso 1994) (Fig. 1). Although an initial report suggested the existence of three acyl-CoA dehydrogenases (FadF, FadG, and FadE) in E. coli with different substrate specificity, latter investigations showed that there is likely a single gene encoding an acyl-CoA dehydrogenase involved in FA degradation ( fadE) and that the mutants described for the three different acyl-CoA dehydrogenases were erroneously mapped (Campbell 2002). FadE is the best example to support the notion that FA metabolism might have certain differences among prokaryotes. Although E. coli seems to have a single fadE gene, most bacterial genomes encode several ACADs; it seems that only some of them participate in the β-oxidation cycle. The P. putida KT2440 genome, for example, has 19 predicted ORFs encoding acyl-CoA dehydrogenases (Winsor et al. 2016) although only a few of them have been experimentally assayed. The one encoded by the gene PP_2216 shows high specificity with short-chain substrates (McMahon et al. 2005); PP_0368 initiates the β-oxidation of the side chain of phenylacyl-CoA compounds – although some activity is also observed with aliphatic acyl-CoA substrates, it displays the highest activities with palmitoyl-CoA (C16) and stearoyl-CoA (C18) (McMahon and Mayhew 2007); and PP_2437 shows preferences toward medium- to longchain length FAs. Other ACADs from the 19 predicted in this organism are related to amino acid metabolism (glutaryl-CoA dehydrogenase encoded by PP_0158 and isovaleryl-CoA dehydrogenase encoded by PP_4064 involved in leucine catabolism) and organic sulfur metabolism (PP_3226 and PP_3259) (Guzik et al. 2014). The Mycobacterium tuberculosis genome is another example of apparent functional redundancy, encoding 35 putative ACADs. Interestingly, some of these dehydrogenases have implications in the degradation of the 5-carbon side chains of cholesterol and constitute an important advantage for the survival of this pathogen within the host macrophage (Wipperman et al. 2013; Ruprecht et al. 2015).

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3.3

Pathways for the Degradation of Fatty Acids in Bacteria

299

Fatty Acid b-Oxidation Multienzyme Complex (FadAB)

FadB carries out the hydration of the 2-enoyl-CoA to 3-hydroxyacyl-CoA (enoylCoA hydratase activity) with further dehydrogenation to produce 3-ketoacyl-CoA (hydroxyacyl-CoA dehydrogenase activity). In this process one molecule of NAD+ is reduced to NADH (Yang et al. 1988). The last step of the β-oxidation cycle is catalyzed by the β-ketothiolase FadA, which releases a molecule of acetyl-CoA and a two-carbon shortened molecule of fatty acyl-CoA (Feigenbaum and Schulz 1975). In E. coli and Pseudomonas spp., FadB and FadA (which are normally encoded in a single operon) form a complex located in the cytosol (DiRusso 1990; Fiedler et al. 2002). The FadBA complex consists of a α2β2 tetramer, composed of two monomers of the 78-kDa α-subunit (the product of fadB) and two monomers of the 48-kDa β-subunit (encoded by fadA) (Binstock et al. 1977). The formation of multienzymatic complexes allows catalytic reactions to proceed without diffusion of the intermediates into the bulk aqueous medium. The complex acts on substrates having 4–16 carbon atoms, although in E. coli the 2-enoyl-CoA hydratase exhibits more activity with SCFAs, while the dehydrogenase and the thiolase have more activity with long- and medium-chain substrates. Seven FadB homologous enzymes were identified in E. coli in silico: MaoC, YfcX, PaaG, PaaF, BhbD, SceH, and YdbU. Of these, MaoC, YfcX, PaaG, PaaF, or YdbU were able to enhance PHA production when overexpressed in an E. coli FadB mutant in which the pathway of PHA biosynthesis had been introduced. These results suggested that these enzymes were able to generate the R-3-hydroxy acyl-CoAs necessary for the PHA biosynthetic pathway in the absence of FadB activity (Snell et al. 2002; Park and Lee 2003; Park and Yup Lee 2004). YfcX together with YfcY were found to form part of the anaerobic β-oxidation pathway that uses nitrate as a terminal respiratory electron acceptor (Campbell et al. 2003; see below). In Pseudomonas putida KT2440, there are two sets of fadAB genes ( fadAB and fadAxBx). fadB is induced by lauric acid (C12) whereas fadBx is not. Consequently, the hypothesis is that FadB and FadA could be the main enzymes in the β-oxidation complex and that fadBx and fadAx might participate only in the absence of active fadB and fadA genes or that these genes are not involved in β-oxidation at all (Wang and Nomura 2010). In Ralstonia eutropha H16, there are two identified gene clusters coding for the β-oxidation of FAs. The two fadB genes ( fabB1 and fabB0 ) have different structural domains (Insomphun et al. 2014). The FabB1 presents a structure similar to that of E. coli FadB, with the enoyl-CoA hydratase (ECH, pfam00378) activity associated with the amino-terminal region and the L-3-hydroxyacyl-CoA dehydrogenase activity (3HCDH) with the central region, which is divided into a 3HCDH_N domain (pfam02737) and two 3HCDH domains (pfam00725). The second FadB homologue (FadB0 ) has an inverted structure of the N-terminal 3HCDH_N and two 3HCDH and C-terminal ECH domains. Outside of these clusters, a third FadB homologue (encoded by fabB2) was identified that had a similar structure to FadB1. According to Insomphun and colleagues, the disruption of fadB0 provokes a significant decrease in dry cell mass when the mutant is grown in soybean oil; this reduction is more significant in the fabB0 fabB1 double mutant. This

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result clearly demonstrates that at least these two genes were involved in β-oxidation and that one set of genes probably acts in the absence of the other. The structures of the FadAB protein complex have been studied in Pseudomonas fragi and E. coli (Ishikawa et al. 1997, 2004). The crystallographic analysis of the FadAB complex in E. coli indicated that the catalytic centers of the ECH (enoylCoA-hydratase), the 3-hydroxyacyl-CoA dehydrogenase (HACD), and the 3-ketoacylCoA thiolase (KACT) face toward the inner solvent region of the complex. However, there is a substantial separation between the three catalytic centers. The binding pocket of the 30 -phosphate ADP moiety is shared by the ECH and HACD but not by KACT. The substrate is anchored through the 30 -phosphate ADP moiety and allows the fatty acid tail to pivot from the ECH to the HACD active sites. Finally, through domain rearrangements, the acyl chain is incorporated into the KACT cavity, and the 30 phosphate ADP is relocated to the KACT active site. The α-helical linker within the structure contributes, through its deformation, to the pivoting center formation and the substrate transfer (Ishikawa et al. 2004). FadA belongs to the thiolase I family; in E. coli it is induced by oleic acid and is required to grow with LCFAs. On the other hand, thiolase II is induced by acetoacetate and is involved in the degradation of butyrate (Feigenbaum and Schulz 1975; Duncombe and Frerman 1976). The E. coli multifunctional complex also contains 3,2-enoyl-CoA isomerase and a 3-hydroxyacyl-CoA epimerase activities. Both activities were proposed to be located within the FadB protein. The epimerase activity allows E. coli to transform D-3-hydroxyacyl-CoA to 3-trans-enoyl-CoA, and the isomerase activity is involved in the degradation of unsaturated FAs with the double bond in an even number of carbon (Yang et al. 1988; Pramanik et al. 1979). The last cycle of the β-oxidation pathway produces a molecule of acetoacetylCoA, in addition to one molecule of acetyl-CoA. The enzyme that catalyzes the cleavage of this molecule is the thiolase II that is likely encoded by atoB in E. coli (Jenkins and Nunn 1987a). An alternative enzyme is the highly homologous protein YqeF which has been used to design a successful synthetic reverse β-oxidation route (Dellomonaco et al. 2011).

3.4

Aerobic Degradation of Acetoacetate and Short-Chain Fatty Acids

As described above, LCFA and MCFA are degraded through the enzymes encoded in the Fad regulon. The degradation of SCFAs requires other enzymes whose expression is not controlled by the FadR regulator. The proteins involved in the degradation of C4-C6 FAs are encoded by the atoDAEB operon (Pauli and Overath 1972; Jenkins and Nunn 1987a), which is induced by acetoacetate (3-oxo-butanoic) but not by saturated SCFAs such as butyrate (C4) and valerate (C5). Although SCFAs presumably cross the outer membrane via porin channels and diffuse across the cytoplasmic membrane in the non-ionized form (Clark and Cronan 2005), the product of atoE is a membrane protein belonging to the 2-hydroxycarboxylate transporter family that is probably involved in SCFA transport (Lolkema 2006).

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The atoD and atoA genes encode the α- and β-subunits of the acetyl-CoA: acetoacetyl-CoA transferase, which activates the acetoacetate to acetoacetyl-CoA (a step similar to that catalyzed by FadD in the degradation of LCFA). atoB encodes β-ketoacyl-CoA thiolase (thiolase II) which is responsible for the thiolytic cleavage of short-chain fatty acyl-CoA and production of acetyl-CoA. As mentioned above, there is another gene in the E. coli genome, yqeF, that encodes a β-ketoacyl-CoA thiolase, highly similar to AtoB but with unknown physiological function. atoC encodes a response regulator of a two-component system (TCS) that acts with the sensor histidine kinase (HK) encode by atoS (Jenkins and Nunn 1987b; Lioliou et al. 2005). Interestingly, AtoC has a dual function as both the transcriptional regulator of the atoDAEB operon and as an antizyme; the posttranslational inhibitor of polyamine biosynthetic enzymes (Fong et al. 1976). In addition to the ATO enzymes, SCFA degradation requires some of the Fad enzymes (FadD, FadE, and FadB; Fig. 2) (Jenkins and Nunn 1987a). For this reason, the degradation in E. coli requires the presence of acetoacetate and LCFA to degrade C4-C6 FAs (Clark and Cronan 2005).

Long-chain fatty acid

OM

SCFA (C6-C4)

Acetoacetate

AtoE?

FadL

P IM FadD

AtoDA

β-oxidation cycles

Short chain fatty acyl-CoA

Fatty acyl-CoA FadE FadB

Acetoacetyl-CoA AtoB

Acetyl-CoA

Fig. 2 Schematic representation of the degradation pathway for short-chain fatty acids (SCFAs). OM outer membrane, IM inner membrane, P periplasmic space

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Degradation of Unsaturated Fatty Acids

Unsaturated FAs constitute about one half of the fatty acid content of E. coli. The lipids contained in the membrane have palmitoleic (9-cis-16:1) and cis-vaccenic (11-cis-18:1) as the main unsaturated FAs, but not polyunsaturated FAs or cis-double bonds extending from even-numbered carbon atoms. For the degradation of FAs with double bonds located at the odd-numbered carbons, the Δ3-cis-Δ2-trans-enoylCoA isomerase activity of FadB carries out the needed transformation, allowing the unsaturated fatty acid to enter into the β-oxidation cycle (Pramanik et al. 1979; Clark and Cronan 2005; Fig. 3). Nevertheless, E. coli is also able to grow using FAs with the unsaturation at the even-numbered carbon atom, such as linoleic acid (9-cis, 12-cis-octadienoic acid). These cases require the 2,4 dienoyl-CoA reductase (FadH) (Fig. 3). This enzyme is Palmitoleic acid (9-cis-hexedecenoic acid) Transport and activation

Linoleic acid (9-cis,12-cis-octadienoic acid) Transport and activation

Fatty acyl-CoA

Fatty acyl-CoA 3 β-oxidation cycles

3 β-oxidation cycles

3-enoyl-CoA

FadB (3,2-enoyl-CoA isomerase activity)

FadB (3,2-enoyl-CoA isomerase activity)

FadBA

FadE

2-enoyl-CoA FadB

2,4-dienoyl-CoA FadH

3-hydroxyacyl-CoA FadB

2 enoyl-CoA

3-ketoacyl-CoA FadA

FadBA

Fatty acyl-CoA

Fatty acyl-CoA

2 β-oxidation cycles

2 β-oxidation cycle

acetyl-CoA

Fig. 3 Schematic representation of degradation of unsaturated fatty acids. Fatty acids with a double bond extending from odd-numbered carbon positions (left branch) require the 3,2-enoylCoA isomerase activity of FadB (enzymatic step shown in gray). Fatty acids with double bonds extending from even-numbered carbons positions (right branch) require the 2,4 dienoyl-CoA reductase activity encoded by fadH (step shown in the white circle)

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an iron-sulfur flavoenzyme that catalyzes the reduction of 2,4-dienoyl-CoA (derived from the oxidation of an unsaturated fatty acid) into 2-enoyl-CoA, which can be further metabolized by FadB. In this process, two reducing equivalents from NADPH are transferred via a direct hydride transfer mechanism to the FAD; this process turns on the transfer of electrons to FMN via the 4Fe-4S cluster. The fully reduced FMN provides a hydride ion to the C5 atom of the substrate, and the residues at the active center of the protein protonate the C4 atom to complete the reaction (Schulz and Kunau 1987; You et al. 1989; Hubbard et al. 2003). The crystal structure of the protein showed the large size of the substrate-binding pocket, which might explain the relative promiscuity of the enzyme that carries out the conversion of 2-trans, 4-cis-, and 2-trans-4-trans-dienoyl-CoA thioesters (Hubbard et al. 2003).

3.6

Anaerobic Degradation of Fatty Acids

Although the aerobic metabolism of FAs in E. coli has been studied profusely, this bacterium is naturally living in the mammalian gut, an environment with a wide variety of carbon and energy sources but severely limited in oxygen. For that reason, it seems unlikely that the FAs, which are abundant in the intestinal lumen, are being degraded by the aerobic pathway. In fact, although there is evidence that some fatty acid degradation activities, such as 3-hydroxyacyl-CoA dehydrogenase and acylCoA dehydrogenase are repressed under anaerobic conditions (Iuchi and Lin 1988), E. coli is able to grow on FAs as carbon source in an anaerobic environment using alternative electron acceptors such as nitrate, fumarate, or trimethylamine (Campbell et al. 2003). These data suggest the existence of an alternative pathway for the degradation of FAs under anaerobic conditions. This hypothesis is further supported by other evidence: (i) A mutant strain lacking FadD, FadE, and FadBA was able to grow using FAs under anaerobic conditions, (ii) E. coli is able to degrade mediumchain FAs such as octanoate and decanoate under anaerobic but not under aerobic conditions, and (iii) the FadR regulator which represses the aerobic growth on FAs failed to block the anaerobic pathway. The yfcX and yfcY genes, which have high sequence homology with fadB and fadA, were the first two genes described in this pathway; these genes form an operon that has since been renamed fadIJ. These genes not only have an important role in the anaerobic growth of E. coli on FAs, using preferably nitrate as electron acceptor, but they can also replace the FadBA under aerobic conditions when these genes are not functioning. YcfYX performs the cleavage of the carbon chain between the β and γ carbons (carbon atoms 2 and 3) as in the classical β-oxidation pathway producing acetyl-CoA (Campbell et al. 2003). In fact, YfcX possesses both enoyl-CoA hydratase and 3-hydroxyacyl-CoA dehydrogenase activities, and yfcY encodes the thiolase activity (Snell et al. 2002). As in the classical aerobic pathway, the pathway requires activation of fatty acid, and the enzyme responsible is YdiD (FadK). YdiD is an acyl-CoA synthetase with preference for SCFA substrates (C24 n-alkanes and pristane, but not towards C8, C12, or C16. The same work identified three outer membrane proteins, named OmpT-1, OmpT-2, and OmpT-3, required for the uptake of n-alkanes (Fig. 4). The transcription of ompT-1 was strongly induced in cells growing on C24–C34 n-alkanes or pristane but less so when C8–C16 n-alkanes were the carbon source used. The transcription of ompT-2 and ompT-3 was efficiently induced by all these

Fig. 4 Regulation of the genes involved in alkane degradation in A. dieselolei B5. Double grey lines depict the outer membrane (OM) and the cytoplasmic membrane (CM). The OM contains the OmpS sensor and the OmpT-1, OmpT-2, and OmpT-3 porins, while the CM includes the Cyo terminal oxidase and the AlkB1/AlkB2 terminal oxidases. The soluble P450 cytochrome is depicted associated with the CM. Rub, rubredoxin; Rbr, rubredoxin reductase; Fdx, ferredoxin; Fdr, ferredoxin reductase. Black lines indicate regulatory effects (+, activation;
, repression); continuous lines stand for direct effects, and discontinuous lines for indirect effects. See text for additional details

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hydrocarbons. Inactivation of the ompT-1 gene impaired growth on C28, C32, and C36 n-alkanes and pristane, although C8–C24 n-alkanes were still used. In contrast, the inactivation of ompT-3 impaired growth on C8–C12 n-alkanes but did not affect the assimilation of C16, C24 n-alkanes or pristane. A mutant strain lacking OmpT-2 did not grow on C16–C24 n-alkanes and grew slowly on shorter or longer n-alkanes. These observations, together with transport studies, suggest that OmpT-1 is used for the uptake of n-alkanes of over 28 carbon atoms and of pristane, while OmpT-2 is preferentially used for C16–C24 n-alkanes, and OmpT3 transports C8–C12 n-alkanes. Inactivation of the gene coding for the OmpT-1 porin abolished chemotaxis towards the hydrocarbons transported by this porin (C28, C32, C36 n-alkanes and pristane). Similarly, the absence of OmpT-2 or OmpT-3 abolishes chemotaxis towards the n-alkanes they transport (Wang and Shao 2014). It therefore seems that the sensing, chemotaxis, uptake, and assimilation of n-alkanes are interdependent processes in A. dieselolei, and probably in other Alcanivorax strains as well. Further, the same authors showed that the inactivation of the gene coding for the cyoD subunit of the Cyo (cytochrome bo) terminal oxidase impaired the use of C32, C36, and pristane as the carbon source, but stimulated growth on C8–C24 n-alkanes. In fact, the expression of cyoD in the wild type strain was lower on C8–C24 n-alkanes and higher on C28–C36 or pristane. Disruption of cyoD led to a strong and constitutive expression of AlmR, a transcriptional repressor that inhibits the expression of almA, the gene that codes for the alkane hydroxylase responsible for the terminal oxidation of C28–C36 n-alkanes and branched alkanes such as pristane (Fig. 4; Wang and Shao 2014). In addition, AlmR was found to inhibit the expression of the OmpT-1 porin, which is preferentially used for the uptake of C24–C36 n-alkanes and pristane but had no effect on OmpT-2 and OmpT-3. However, Cyo was found to inhibit the expression of OmpT-2 and OmpT-3. Therefore, the influence of Cyo on the assimilation of C32, C36, and pristane derives, in part, from its ability to control the expression of AlmR but also from its influence on the expression of the porins involved in the uptake of the different alkanes. As explained in Sect. 2.2, the Cyo terminal oxidase also modulates the expression of the n-alkane degradation genes specified in the P. putida OCT plasmid. The exact mechanism through which Cyo controls gene expression in P. putida and A. dieselolei is at present unknown. As a terminal oxidase of the electron transport chain, Cyo receives electrons from the reduced ubiquinone of the cytoplasmic membrane and transfers them to oxygen, pumping protons out of the cell and thus contributing to the proton gradient that allows the function of ATPase and of several membrane transporters (Anraku and Gennis 1987; Nakamura et al. 1997; Williams et al. 2007). Its influence on the expression of genes involved in the assimilation of alkanes suggests that Cyo might provide information on the redox status of the electron transport chain to an unknown sensor protein, which in turn influences the activity of, for example, a transcriptional regulator. However the elements connecting Cyo with the expression of the final target genes have not been identified.

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2.4

Genetic Features and Regulation of n-Alkane Metabolism in Bacteria

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Regulation of n-Alkanes Degradation Genes in Dietzia Species

In the Gram positive strain Dietzia sp. DQ12-45-1b, which can assimilate n-alkanes ranging from 6 to 36 carbon atoms in length, two different alkane hydroxylases are responsible for the initial oxidation of the n-alkanes: a P450 cytochrome of the CYP153-family and an AlkB-type alkane hydroxylase named AlkW1 (Nie et al. 2011, 2014). The P450 enzyme oxidizes C6-C10 n-alkanes, while AlkW1 is responsible for the oxidation of C14-C36 n-alkanes (Nie et al. 2011, 2014). The gene coding for the P450 enzyme is induced in the presence of C8–C14 n-alkanes, a control mechanism that relies on CypR, a transcriptional activator of the AraC-family (Liang et al. 2016a). The expression of AlkW1 is controlled by the AlkX transcriptional repressor, which binds to and represses the promoter of alkW1 (Liang et al. 2016b). The latter gene is efficiently expressed when cells grow at the expense of C8–C36 n-alkanes (Nie et al. 2011), although the n-alkanes are not effectors for the AlkX repressor; rather, repression is abrogated by the fatty acids derived from the oxidation of the nalkanes (Liang et al. 2016b). In fact, AlkX was shown by the latter authors to bind in vitro C10–C24 fatty acids, which inhibited binding of the repressor to the alkW1 promoter. The alkX gene maps immediately downstream of alkW1; both genes are co-transcribed and therefore co-regulated through a positive feedback mechanism governed by the concentration of fatty acids within the cell.

3

General Patterns in the Regulation of Alkane Degradation Genes

3.1

Specific Regulators: Are there Common Characteristics?

The identification of the specific regulators that induce the genes responsible for the assimilation of n-alkanes in response to the presence of these hydrocarbons 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 families, or to other nonrelated families of regulators and can act as transcriptional activators or repressors (see Table 1). Proteins homologous to the AlkS regulator of the OCT plasmid n-alkane degradation pathway (see Sect. 2.2) have been repeatedly found associated with genes coding for alkane hydroxylases of the AlkB family. AlkS belongs to the LuxR family of regulators, which 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, including AlkS, are grouped within the MalT subfamily and have an unusually long N-terminal domain that includes an ATP binding site. At least in the case of MalT, transcriptional

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Table 1 Transcriptional regulators known or presumed to control the expression of alkane degradation genes. Global regulators are not included Bacterium T. butanivorans

Gene bmoR

Effector C2–C8 n-alkanols

Evidence Direct

alkS

Family σ54dependent LuxR/MalT

P. putida GPo1

C6–C10 n-alkanes

Direct

P. putida P1

alkS

LuxR/MalT

Not tested

Similarity

A. borkumensis SK2

alkS

LuxR/MalT

C8–C20 n-alkanes

Direct

A. borkumensis AP1 A. borkumensis SK2 A. dieselolei B5

alkS

LuxR/MalT

Not tested

Similarity

cypR

AraC/XylS

C8–C18 n-alkanes

Direct

almR

AlmR

Direct

alkX

TetR

Expression of AlmR repressor induced by C8–C 24 n-alkanes C10–C24 fatty acids

cypR

AraC/XylS

C8–C 14 n-alkanes

Direct

alkR

AraC/XylS

Direct

alkRa

AraC/XylS

C7–C 18 n-alkanes >C22 n-alkanes

alkRb

OruR

C16–C22 n-alkanes

Indirect

Dietzia sp. DQ12-45-b1 Dietzia sp. DQ12-45-b1 Acinetobacter sp. ADP1 Acinetobacter sp. M1 Acinetobacter sp. M1

Direct

Indirect

Reference Kurth et al. (2008) Panke et al. (1999), Sticher et al. (1997) van Beilen et al. (2001) Schneiker et al. (2006); Kumari et al. (2011) van Beilen et al. (2004) Schneiker et al. (2006) Wang and Shao (2014)

Liang et al. (2016b) Liang et al. (2016a) Ratajczak et al. (1998) Tani et al. (2001) Tani et al. (2001)

activation requires ATP (but not ATP hydrolysis) and an effector, which triggers the multimerization of the protein (Schreiber and Richet 1999). Genes coding for AlkBlike alkane hydroxylases can, however, be controlled by regulators of other families, such as those of the AraC/XylS family (see, for example, Ratajczak et al. 1998). These regulators are widespread and have two domains, one of which is conserved and contains determinants for DNA binding and transcription activation, while the nonconserved domain is critical for signal recognition in those members of the family that are activated by the binding of an effector (Gallegos et al. 1997). For some of these regulators there is evidence supporting the idea that n-alkanes or n-alkanols act as effectors. The water solubility of alkanes with more than eight carbon atoms is below the micromolar range, thus these hydrocarbons most likely accumulate in the cytoplasmic membrane. Transcriptional regulators are normally cytoplasmic proteins which interact with DNA. The question arises as to how these

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regulators interact with the alkanes. The A. borkumensis AlkS transcriptional regulator activates the 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 with the membrane fraction rather than the cytoplasmic fraction (Sabirova et al. 2006). Although AlkS does not show the characteristics expected of a membrane protein, it may have affinity for the inner side of the cytoplasmic membrane, where it has easy access to the alkanes acting as effectors. Once bound to the alkane, AlkS must move and find its binding site on the DNA. The possible changes that the alkane effector can induce on AlkS are unknown. The membrane affinity of other alkaneresponsive regulators has not been analyzed either.

3.2

Alkanes Are Not Preferred Growth Substrates for Many of the Bacterial Species that Can Assimilate Them

Many hydrocarbon-degrading bacteria have a versatile metabolism and can use other carbon sources in addition to hydrocarbons. In most of these, hydrocarbons are not the preferred growth substrates, and their assimilation is inhibited and delayed until the preferred compounds have been consumed. Strains of P. putida, Pseudomonas aeruginosa, Burkholderia cepacia, T. butanivorans, or Acinetobacter sp. are well known examples of such behaviour (Doughty et al. 2006; Marín et al. 2001, 2003; Ratajczak et al. 1998; Staijen et al. 1999; Yuste et al. 1998). A detailed description of the mechanisms involved is provided above for the n-alkane degradation pathways present in T. butanivorans (Sect. 2.1) and in P. putida (Sect. 2.2). The preferential use of a carbon source over other potential substrates when these are all present in a growth medium is known as catabolite repression control (Magasanik 1970). However, the preferred compounds vary in different bacterial species and the mechanisms used to modulate the expression of catabolic pathways, where characterized, frequently differ as well (Rojo 2010). This process probably arises from a number of global regulatory mechanisms directed towards optimizing carbon metabolism and energy generation in response to different signals. What signals these are is not clear, but they may be related to the concentrations of key metabolites or molecules in the cell that in turn depend on the energy gain provided by different catabolic pathways. Alkanes are generally considered nontoxic compounds. However, there are several reports indicating that the use of alkanes as a carbon source can have harmful consequences for cell physiology. The growth of P. putida GPo1 on 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 first enzyme of the n-alkanes degradation pathway, the AlkB alkane hydroxylase, which is an integral membrane protein. Induction of the alkB gene by the gratuitous inducer dicyclopropylketone, which is not used as a carbon source, is harmful to cell physiology. The growth of cells in complete medium with dicyclopropylketone for several generations led to the isolation of mutant derivatives in which alkB could no longer be induced, an effect not observed if

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dicyclopropylketone was omitted (Chen et al. 1996). Apparently, AlkB alkane hydroxylase and AlkJ alcohol dehydrogenase are produced in very large amounts when expression of the pathway genes is fully induced, 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 exert on the expression of alkane degradation genes in several bacterial strains (Doughty et al. 2006; Marín et al. 2001, 2003; Ratajczak et al. 1998); this “product repression” effect coordinates the generation of these compounds with their onward metabolism, thereby avoiding their harmful accumulation in the cell membrane.

3.3

Differential Regulation of Multiple Alkane Hydroxylases

Alkanes are usually present in the environment as mixtures of molecules with different chain lengths and structures (linear or branched), which may in part explain why many bacterial strains have several alkane hydroxylases with substrate specificities that may frequently overlap and that are often regulated in a coordinated fashion. The example described in Sect. 2.3 for Alcanivorax species is particularly illuminating and complex, but other examples are available. For example, Acinetobacter sp. M-1 contains two AlkB-related alkane hydroxylases named AlkMa and AlkMb that are differentially regulated depending on the alkane used as carbon source, although it is not clear which are the enzymatic differences between the two proteins. The expression of AlkMa, which is controlled by the AlkRa regulator, is induced by n-alkanes with a very long chain length (>C22), while that of AlkMb is induced by AlkRb in the presence of C16–C22 n-alkanes (Tani et al. 2001). P. aeruginosa strains RR1 and PAO1 contain two alkane hydroxylases, AlkB1 and AlkB2. The substrate range of the two enzymes is very similar, with both able to oxidize C12–C16 n-alkanes (Smits et al. 2002). The regulation of the genes encoding AlkB1 and AlkB2 has been studied in strain RR1 (Marín et al. 2003). The expression of both is induced by C10–C22 n-alkanes, although alkB2 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 the stationary phase. The regulators responsible for this differential regulation have not been characterized. The alkB1 and alkB2 genes are also present in P. aeruginosa SJTD1. Although their individual inactivation was found to have no effect on the ability of this strain to grow on C12–C24 n-alkanes, the double mutant lacking both AlkB1 and AlkB2 hydroxylases failed to grow on C12–C16 n-alkanes, but it could still thrive at the expense of C18–C24 n-alkanes (Liu et al. 2014). Therefore, AlkB1 and AlkB2 are responsible for the assimilation of C12–C16 n-alkanes, but additional alkane hydroxylases able to oxidize C18–C24 n-alkanes must be present in this strain. Two genes coding for proteins homologous to the P450 cytochromes of the CYP-153 family and a gene coding for a putative AlmA alkane hydroxylase were

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detected in P. aeruginosa SJTD1, although their inactivation (individual or combined) had no effect on growth on C18–C24 n-alkanes (Liu et al. 2014). It therefore remains unclear which alkane hydroxylase is able to oxidize these high molecular weight n-alkanes. Although the expression of the different components of the alkane degradation pathways is frequently coordinated, this is not always the case. In P. putida GPo1, the expression of the genes coding for 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 n-alkanes (see Sect. 2.2; reviewed in van Beilen et al. 2001). However, there are 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 (Marín et al. 2003), while the expression of the genes coding for the corresponding AlkB-like alkane monooxygenases are inducible by n-alkanes. In these bacteria it is unclear whether the rubredoxin and rubredoxin reductase are shared with other monooxygenases, even serving purposes different to the oxidation of n-alkanes.

4

Research Needs

While it is clear that the expression of most alkane degradation genes is regulated, how regulation is accomplished is often unclear. The specific regulators responsible for the induction of these pathways have been identified in very few bacterial species. Their characterization is important, especially in those microorganisms with several alkane hydroxylation systems, each with its own (albeit sometimes overlapping) specificity. Such knowledge would be particularly important with respect to hydrocarbonoclastic bacteria such as Alcanivorax sp., which specialize in alkane metabolism and contain several independent alkane hydroxylation systems. An intriguing question is why the expression of the monooxygenase component of alkane hydroxylases is usually regulated while that of the auxiliary proteins rubredoxin and rubredoxin reductase is constitutive in many bacteria. They may serve other functions in addition to the transfer of electrons to alkane monooxygenases, but this is not known for sure. As explained in Sects. 2.1, 2.2, and 2.3, the expression of the alkane degradation pathways is frequently coordinated with other aspects of cell metabolism, including chemotaxis, transport, and catabolite repression control. Further work on different microorganisms is required, however, since the molecular mechanisms likely differ. Finally, there are indications that the expression of n-alkane degradation pathway components is coordinated with the induction of fatty acid metabolism and the generation of storage polymers, but the details on this are again missing for most bacteria. A better knowledge of these regulatory processes would allow bioremediation strategies to be optimized, as well as the construction of strains with different elements of alkane degradation pathways useful in biotransformations or in other processes. Indeed, different biosensors have already been developed based on

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regulator/promoters pairs derived from n-alkane degradation pathways. These allow the detection of low concentrations of n-alkanes in water samples (Sticher et al. 1997; Kumari et al. 2011; Sevilla et al. 2015) and have been found useful for the monitoring of crude oil spills in seawater (Brussaard et al. 2016). Other useful applications will surely follow. Acknowledgments Work in the author’s laboratory is funded by the Spanish Ministry of Economy and Competitiveness (grant BIO2015-66203-P; MINECO/FEDER) and the European Commission VII Framework Program (grant number 312139).

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Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

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Ryouichi Fukuda and Akinori Ohta

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Response to n-Alkanes in n-Alkane-Assimilating Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Mechanism of Transcriptional Activation of Genes Responsible for n-Alkane Degradation in Y. lipolytica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Promoter Elements of the CYP52-Family P450 Gene Involved in n-Alkane Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Transcription Activators Involved in n-Alkane Response . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Regulation of n-Alkane Metabolic Genes by the Opi1-Family Transcription Factor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Role of the Opi1-Family Proteins in Other Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Repression of Transcription of n-Alkane Metabolic Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

The yeasts Candida tropicalis, Candida maltosa, and Yarrowia lipolytica have an excellent ability to use n-alkanes as the sole carbon and energy source. Here, we summarize the current knowledge of the genetic features and regulation of n-alkane metabolism in these yeasts. The transcription of genes encoding the CYP52-family cytochromes P450 that catalyze the initial hydroxylation of n-alkanes has been shown to be activated when these yeasts are cultured in the presence of n-alkanes. In Y. lipolytica, the transcription of ALK1, the gene R. Fukuda (*) Department of Biotechnology, The University of Tokyo, Tokyo, Japan e-mail: [email protected] A. Ohta Department of Biological Chemistry, College of Bioscience and Biotechnology, Chubu University, Kasugai, Aichi, Japan e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_24

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encoding P450, is activated by a complex composed of two basic helix-loop-helix transcription activators Yas1p and Yas2p through a promoter element ARE1. This transcription is regulated by an Opi1-family transcriptional repressor Yas3p. In the absence of n-alkanes, Yas3p binds to Yas2p in the nucleus thereby repressing the transcription of ALK1. However, in the presence of n-alkanes, Yas3p is sequestered to the endoplasmic reticulum to derepress the transcription of the gene.

1

Introduction

A variety of microorganisms, including certain species of yeasts, has developed metabolic systems to assimilate n-alkanes containing 10–18 carbons as the sole carbon and energy source. The degradation pathway of n-alkanes in yeasts and the enzymes involved in it have been extensively studied in Candida tropicalis (Tanaka and Fukui 1989), Candida maltosa (Mauersberger et al. 1996), and Yarrowia lipolytica (Barth and Gaillardin 1996, 1997; Fickers et al. 2005; Fukuda 2013; Fukuda and Ohta 2013; Nicaud 2012) as described in Chap. 6, ▶ “Enzymes for Aerobic Degradation of Alkanes in Yeasts” by Fukuda and Ohta. In these yeasts, n-alkanes are first hydroxylated to fatty alcohols in the endoplasmic reticulum (ER) by cytochromes P450 belonging to the CYP52 family (Figs. 2 and 4 of the Chap. 6, ▶ “Enzymes for Aerobic Degradation of Alkanes in Yeasts” by Fukuda and Ohta). Fatty alcohols are then oxidized to fatty aldehydes by fatty alcohol dehydrogenase (FADH) in the ER or by fatty alcohol oxidase (FAO) in the peroxisome. Fatty aldehydes are further oxidized to fatty acids by fatty aldehyde dehydrogenase (FALDH) in the ER or the peroxisome, and then finally activated to acyl-CoAs by acyl-CoA synthetase (ACS). These activated fatty acids are then utilized for lipid synthesis, or degraded in the peroxisome via β-oxidation. It is crucial for an organism to respond to and adapt rapidly to environmental changes for survival. In C. tropicalis, C. maltosa, and Y. lipolytica, the metabolism of n-alkanes was found to be regulated at the transcriptional level. The transcription of the genes encoding enzymes involved in the metabolism of n-alkanes was found to be activated in the presence of n-alkanes. The mechanism of the transcription regulation in these yeasts is a very interesting subject, particularly because the regulation of transcription by hydrophobic compounds in lower eukaryotes remains largely elusive. This chapter will focus on the molecular mechanisms of the transcriptional regulation of genes involved in the degradation of n-alkanes in these yeasts.

2

Response to n-Alkanes in n-Alkane-Assimilating Yeasts

Experiments conducted in the 1970s showed that the production of cytochromes P450 was induced when C. tropicalis was cultured in a medium containing n-tetradecane (Lebeault et al. 1971). Cloning and subsequent characterization of

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Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

Fig. 1 Response to n-alkanes in n-alkane-assimilating yeasts. When n-alkaneassimilating yeasts are cultured in the medium containing n-alkane, the transcriptional induction of genes involved in the degradation of n-alkanes and the proliferation of the ER and the peroxisome, in which nalkanes are degraded, are observed

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the genes in C. tropicalis revealed that this yeast has at least eight genes encoding the CYP52-family P450s, (ALK1–ALK7 and CYP52D2) and that the transcription of four of them (ALK1–ALK3 and ALK5) is activated by n-alkanes (Fig. 1) (Craft et al. 2003; Nelson 2009; Sanglard et al. 1987;Seghezzi et al. 1991, 1992). In C. maltosa, of the eight genes (ALK1–ALK8) encoding the CYP52-family P450s, the transcription of all except ALK4 was induced by n-alkanes (Ohkuma et al. 1991, 1995a). Y. lipolytica has 12 genes (ALK1–ALK12) encoding the CYP52-family P450s, transcription of nine of which (ALK1–ALK6, ALK9, ALK11, and ALK12) was induced by n-alkanes (Hirakawa et al. 2009; Iida et al. 1998, 2000; Iwama et al. 2016; Takai et al. 2012). The transcriptional induction of the P450 genes by n-alkanes was also observed in sophorolipid-producing yeast Candida bombicola (Van Bogaert et al. 2009). The transcription of genes involved in the degradation of n-alkane metabolites was also found to be induced in the presence of n-alkanes in Y. lipolytica (Fig. 4 of the Chap. 6, ▶ “Enzymes for Aerobic Degradation of Alkanes in Yeasts” by Fukuda and Ohta). In Y. lipolytica, the transcription of ADH1 and ADH3, encoding alcohol dehydrogenases, and FAO1, encoding a fatty alcohol oxidase, is upregulated in the presence of n-alkanes (Iwama et al. 2015). In addition, transcription of three (HFD1–HFD3) of the four genes (HFD1–HFD4) encoding fatty aldehyde dehydrogenases and FAA1 and FAT1 encoding ACSs was increased in the presence of n-alkanes (Iwama et al. 2014; Tenagy et al. 2015). The transcription of PAT1 encoding a peroxisomal acetoacetyl-CoA thiolase involved in β-oxidation was also induced by n-alkanes (Yamagami et al. 2001). These results confirm that the transcription of genes important for the n-alkane degradation is activated in response to n-alkanes. Our transcriptome analysis in Y. lipolytica cells cultured in medium containing either glucose or n-decane suggested that the transcripts of approximately 500 genes were increased more than twofold in response to n-decane (our unpublished results). n-Alkanes have also been reported to induce the proliferation of the ER and peroxisome in the n-alkane-assimilating yeasts (Fig. 1) (Mauersberger et al. 1987;

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Osumi et al. 1974; Vogel et al. 1992). Proliferation of the peroxisome by fatty acids has been widely observed in various organisms, including C. tropicalis, Y. lipolytica, and Saccharomyces cerevisiae, and its mechanism has been extensively studied (Gurvitz and Rottensteiner 2006). In contrast, the mechanism underlying the proliferation of the ER by n-alkanes in n-alkane-assimilating yeasts remains poorly understood. Interestingly, overproduction of the CYP52-family P450 induced the proliferation of the ER in C. maltosa and S. cerevisiae (Ohkuma et al. 1995b; Schunck et al. 1991). Proliferation of the ER due to overproduction of various membrane proteins has been reported (Federovitch et al. 2005) and may be one of the quality control mechanisms of the ER to avoid over accumulation of proteins in the ER membrane.

3

Mechanism of Transcriptional Activation of Genes Responsible for n-Alkane Degradation in Y. lipolytica

The transcriptional induction of the P450 genes in response to n-alkane was first identified in yeasts of the Candida genus as described above. However, the molecular mechanism of its regulation remains unclear. This is largely due to the difficulty in obtaining mutant strains defective for transcriptional activation in the presence of n-alkanes, since C. tropicalis and C. maltosa are diploid or partial diploid yeasts, in which teleomorphs have not been found. Furthermore, the CUG codon has been shown to code for serine instead of leucine in C. tropicalis and C. maltosa, as well as in other n-alkane-assimilating yeasts phylogenetically close to them, including Candida albicans, Candida dubliniensis, Candida parapsilosis, Debaryomyces hansenii, Lodderomyces elongisporus, and Meyerozyma guilliermondii (Massey et al. 2003; Sugiyama et al. 1995; Ueda et al. 1994), and this poses an obstacle to the analysis of DNA-protein and/or protein-protein interaction using S. cerevisiae system. Y. lipolytica, on the other hand, has a teleomorph and a stable haploid and diploid life cycle, and genetic methods that permit isolation and characterization of mutants as well as molecular biology methods are well established in it (Barth and Gaillardin 1996, 1997). Among the yeasts that can assimilate n-alkanes, the genome sequences were determined initially in Y. lipolytica, D. hansenii (Dujon et al. 2004), and C. albicans (Jones et al. 2004), followed by C. tropicalis and others (Butler et al. 2009). As a result of these advantages, the mechanism of the transcriptional regulation by n-alkane has been elucidated in Y. lipolytica.

3.1

Promoter Elements of the CYP52-Family P450 Gene Involved in n-Alkane Response

The transcriptional regulation in response to n-alkanes has been investigated by studying ALK1 encoding the primary CYP52-family P450 in the assimilation of n-alkanes in Y. lipolytica (Iida et al. 1998, 2000; Iwama et al. 2016; Takai et al.

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2012). The transcription of ALK1 was highly activated in the presence of n-alkanes and the transcript level was found to be highest among the 12 ALK genes (Hirakawa et al. 2009). Promoter analysis of ALK1 led to the identification of a sequence named ARR1 (alkane responsive region 1), which is involved in the transcription activation (Sumita et al. 2002). ARR1 was found to contain two elements, ARE1 (alkane responsive element 1) and ARE2. In electrophoretic mobility shift assay, specific shift bands corresponding to both ARE1 and ARE2 could be identified with cellular extracts of Y. lipolytica cells cultured in n-alkane-containing medium. ARE1 contains a sequence similar to the E-box motif, the consensus sequence to which basic helix-loop-helix (bHLH) transcription factors interact (Murre et al. 1994), and was found to play a critical role in the transcriptional activation process (see below). Interestingly, ARE1-like sequences were found in the promoter regions of various genes involved in n-alkane metabolism (Yamagami et al. 2004). In contrast, the role of ARE2 is still unknown.

3.2

Transcription Activators Involved in n-Alkane Response

A gene YAS1 encoding a transcription factor that activates the ARE1-mediated transcription was identified by the analysis of a mutant defective in the transcriptional activation through ARE1 by n-alkanes and in the growth on n-alkanes (Yamagami et al. 2004). YAS1 encodes a bHLH transcription factor of 137 amino acids (Fig. 2). A deletion mutant of YAS1 showed defects in the transcription activation of ALK1 by n-alkanes. However, Yas1p did not bind to ARE1 in vitro. bHLH transcription factors generally form homo- or heterodimers through their HLH regions and interact with the E-box motif through the basic regions (Murre et al. 1994). Indeed, a gene named YAS2 encoding a 700-amino acid protein that contains a bHLH motif similar to that of Yas1 was identified from the genome database. This gene was found to be involved in the ARE1-mediated transcriptional activation (Fig. 2) (Endoh-Yamagami et al. 2007). The deletion mutant of YAS2 was also defective in the transcription induction of ALK1 by n-alkanes. Yas1p and Yas2p formed a complex in vitro and bound to ARE1 only when both proteins existed. Yas1p and Yas2p constitutively localized in the nucleus (Hirakawa et al. 2009; Yamagami et al. 2004). These results suggest that the complex of Yas1p and Yas2p binds to ARE1 and activates the transcription of ALK1 in response to n-alkane. Deletion mutants of YAS1 or YAS2 did not have the ability to grow on n-alkanes, indicating the importance of Yas1p-Yas2p complex in the ARE1mediated transcriptional activation.

3.3

Regulation of n-Alkane Metabolic Genes by the Opi1-Family Transcription Factor

The bHLH motifs of Yas1p and Yas2p show sequence similarities to Ino4p and Ino2p, respectively, of S. cerevisiae. In S. cerevisiae, these function as transcription

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Fig. 2 Schematic diagrams of bHLH transcription activators and Opi1-family proteins in S. cerevisiae and Y. lipolytica. The bHLH motifs, repressor interaction domains (RID), PA-binding domains, FFAT motifs, leucine zipper motifs, and activator interaction domains (AID) are indicated by grey boxes

activators regulating genes involved in phospholipid synthesis. In this yeast, transcription of phospholipid synthetic genes, including INO1 encoding inositol3-phosphate synthase, is activated in the absence of myo-inositol and repressed in the presence of it (Henry et al. 2012). A heterodimer of Ino2p and Ino4p constitutively binds to an upstream activating element, UASINO/ICRE (inositol choline responsive element), in the promoter regions of the target genes. The transcription repressor Opi1p binds through its activator interaction domain (AID) to the repressor interaction domain (RID) of Ino2p (Heyken et al. 2005). Loewen et al. proposed a model in which Opi1p is retained to the ER by binding to phosphatidic acid (PA) using its PA-binding domain and to an ER membrane-spanning protein Scs2p through its FFAT (two phenylalanine in an acidic tract) motif in the absence of myo-inositol. This leads to the activation of transcription of the phospholipid synthesis genes by the Ino2-Ino4 complex (Loewen et al. 2004). However, in the presence of myo-inositol, PA is utilized for the synthesis of phosphatidylinositol (PI). Opi1p is then released from the ER and transported to the nucleus where it binds to Ino2p to repress the transcription. Therefore, Opi1p is the key regulator controlling transcription of genes responsible for phospholipid biosynthesis in response to myo-inositol.

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In the Y. lipolytica genome database, an ortholog of OPI1 was identified and named as YAS3 (Hirakawa et al. 2009). Two different transcripts with different transcription initiation sites were obtained from YAS3. These transcripts were predicted to encode a long form of Yas3p (l-Yas3p of 727 amino acids) and a short form of Yas3p (s-Yas3p of 422 amino acids) (Fig. 2). The AID, PA-binding domain, and leucine zipper domain found in Opi1p were conserved in both forms of Yas3p, but the FFAT motif was absent in Yas3p (Fig. 2). Whether these two forms of Yas3p have different functions is still not clear, but it has been shown that the s-Yas3p is sufficient to regulate the transcription of ALK1. The deletion mutant of YAS3 accumulated much more transcripts of ALK1 than the wild-type strain, even when cells were cultured in the medium containing glucose or glycerol. Yas3p interacted with Yas2p, but not with Yas1p. In line with this observation, a RID-like sequence motif was found in Yas2p (Fig. 2). In addition, while Yas3p was localized in the nucleus when cultured in the medium containing glucose, it was sequestered to the ER in the presence of n-alkanes. Among the 12 ALK genes, the transcription of ALK1, ALK2, ALK4, ALK6, ALK9, and ALK11 appeared to be regulated by the Yas1p-Yas2p-Yas3p system. In contrast, Yas3p was not involved in the transcriptional regulation of INO1 by myo-inositol. A question remained as to why Yas3p remained localized to the ER in the presence of n-alkanes. Yas3p was found to bind to PA and phosphoinositides (PIPs), particularly to phosphatidylinositol 4-phosphate (PI(4)P), in vitro, but not to n-alkanes (Kobayashi et al. 2013). In addition, the ARE1-mediated transcription was upregulated in mutants defective for an ortholog of S. cerevisiae PAH1, encoding PA phosphatase and an ortholog of SAC1, encoding PIP phosphatase in the ER. These results suggest that Yas3p is localized to the ER by binding to PA and/or PIP in the ER membrane. In contrast to S. cerevisiae, an ortholog of SCS2 or its paralog, SCS22, was not required for the transcriptional activation of ALK1 by n-alkanes in line with the absence of FFAT motif-like sequence in Yas3p, although the deletion mutant of SCS2 exhibited a growth defect when cultured on n-decane (Kobayashi et al. 2008). Based on these results, a model of the transcriptional regulation of n-alkane metabolic genes was proposed (Fig. 3). A heterocomplex of the bHLH transcription activators, Yas1p and Yas2p, constitutively localizes in the nucleus and binds to ARE1 in the promoter regions of the genes involved in the n-alkane metabolism. In the absence of n-alkanes, Yas3p is transported to the nucleus and binds to Yas2p of the Yas1p-Yas2p complex, resulting in repression of the ARE1-dependent transcription. When the medium is supplemented with n-alkanes, Yas3p is retained to the ER by binding to PA and/or PIP, and the transcription is activated by Yas1p-Yas2p complex. It remains to be determined whether the amounts of PA and PIPs in the ER membrane increase in response to n-alkanes. Scs2p is not involved in this process, but it is possible that other ER-resident protein is involved in the localization of Yas3p to the ER. This is suggested by the fact that the C-terminal region of Yas3p was found to be localized to the ER in a PA- and PI(4)P-independent manner (Kobayashi et al. 2015).

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Fig. 3 Model of n-alkaneresponsive transcriptional regulation by the transcription factors Yas1p, Yas2p, and Yas3p in Y. lipolytica. In the absence of n-alkanes (upper), Opi1-family transcription repressor Yas3p binds to the complex composed of bHLH transcription activators Yas1p and Yas2p in the nucleus, and ARE1-dependent transcription is repressed. In the presence of n-alkanes (lower), Yas3p is sequestered to the ER membrane through the interaction with PA and/or PIP, and ARE1-dependent transcription is activated by Yas1p-Yas2p complex

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3.4

Role of the Opi1-Family Proteins in Other Yeasts

Orthologs of OPI1 exist in the genomes of a variety of yeasts (Fig. 4), although their functions are largely unknown. In Candida glabrata, a yeast closely related to S. cerevisiae phylogenetically, a homolog of Opi1p is involved in the transcriptional regulation of an ortholog of INO1 by myo-inositol (Bethea et al. 2010). In contrast, Opi1p homolog does not regulate INO1 expression in C. albicans, but it controls the expression of SAP2 encoding the secreted aspartyl protease and is involved in the filamentous growth and virulence (Chen et al. 2015). Therefore, Opi1-family proteins possibly regulate processes other than phospholipid synthesis. n-Alkane assimilating yeasts C. tropicalis, C. maltosa, C. dubliniensis, C. parapsilosis, D. hansenii, L. elongisporus, and M. guilliermondii all have Opi1-family proteins, and it would be of great interest to examine whether these orthologs are involved in the transcriptional regulation of n-alkane metabolism in these yeasts.

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Saccharomyces cerevisiae Candida glabrata

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Fig. 4 Phylogenetic tree of the Opi1-family proteins in yeasts. Phylogenetic tree of the Opi1family proteins of yeasts was constructed using ClustalW (DDBJ, v2.1) and drawn using Njplot. The scale bar indicates 0.1 substitutions per site. The bootstrap values by 1000 repetitions are indicated. The accession numbers of sequences from UniProtKB are as follows: Ashbya gossypii (Q75DH7), Candida albicans (Q5ALN4), Candida dubliniensis (B9WAR2), Candida glabrata (Q6FN27), C. maltosa (M3JFB2), Candida parapsilosis (G8BGL1), C. tropicalis (C5M6C2), Debaryomyces hansenii (Q6BJD0), Kluyveromyces lactis (Q6CIM8), Komagataella pastoris (A0A1B2J6H0), Lodderomyces elongisporus (A5DT94), Meyerozyma guilliermondii (A5DPS9), Ogataea polymorpha (A0A1B7SCJ0), Rhodosporidium toruloides (M7XL84), S. cerevisiae (P21957), Ustilago maydis (A0A0D1CM93), and Y. lipolytica (B9X0I4)

4

Repression of Transcription of n-Alkane Metabolic Genes

Glucose is the primary source of carbon and energy for most organisms, and the transcription of genes involved in other carbon and energy source metabolism remains repressed in the presence of glucose. Transcriptional repression of genes involved in n-alkane metabolism by glucose is observed in n-alkane-assimilating yeasts. In C. tropicalis and C. bombicola, expression of a subset of the CYP52family P450 genes induced by n-alkanes is repressed by glucose (Seghezzi et al. 1992; Van Bogaert et al. 2009). In C. maltosa, transcription of most of the ALK genes is severely repressed by glucose, but not by glycerol (Ohkuma et al. 1995a). The carbon catabolite repression by glucose has been well documented in S. cerevisiae (Conrad et al. 2014; Kayikci and Nielsen 2015), but it remains to be elucidated

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whether the transcription of the P450 genes is repressed by a similar mechanism in these n-alkane-assimilating yeasts. In marked contrast to these yeasts, in Y. lipolytica, the transcription of genes involved in n-alkane metabolism is strictly repressed by glycerol, but not so much by glucose (Iida et al. 1998, 2000; Mori et al. 2013). In line with this observation, glycerol is a preferred carbon and energy source for this organism, and it shows better growth on glycerol than on glucose (Mori et al. 2013). The transcriptional repression by glycerol was also observed in Kluyveromyces lactis, in which the transcription of KlICL1 encoding isocitrate lyase was repressed by glycerol (Rodicio et al. 2008). The molecular mechanisms underlying the transcriptional repression by glycerol is unclear, but it was shown that phosphorylation of glycerol is required for the glycerol repression in both these yeasts (Mori et al. 2013; Rodicio et al. 2008).

5

Research Needs

It has been revealed that, in Y. lipolytica, the Opi1-family protein Yas3p plays a pivotal role in the transcriptional regulation of genes involved in n-alkane metabolism by n-alkanes. However, it remains to be clarified how n-alkanes are recognized and how these signals are transduced retaining Yas3p to the ER. Transcriptional activation of the ARE1-containing promoter by n-alkane was also observed in the deletion mutant of the 12 ALK genes, which could not utilize n-alkanes owing to a defect in the hydroxylation of n-alkanes (Takai et al. 2012). This suggests that n-alkanes and not their metabolites activate the ARE1-mediated transcription. It is possible that there are proteins that sense n-alkanes and trigger the transcriptional response to n-alkanes. Alternatively, since n-alkanes are supposed to accumulate in the membranes, alterations in the membrane conditions may be perceived by a sensor protein. n-Alkane-assimilating yeasts have been shown to have great potential for production of single-cell protein (SCP) as well as various useful compounds, including long-chain dicarboxylic acids, by metabolizing n-alkanes (Barth and Gaillardin 1996, 1997; Fickers et al. 2005; Mauersberger et al. 1996; Tanaka and Fukui 1989). Elucidation of the mechanisms underlying the regulation of n-alkane metabolism will contribute to the construction of efficient bioconversion systems using these n-alkane-assimilating yeasts.

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Genetics and Ecology of Isoprene Degradation

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Andrew T. Crombie, Nasmille L. Mejia-Florez, Terry J. McGenity, and J. Colin Murrell

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Microbial Consumption of Isoprene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial Degradation of Isoprene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Ecology of Isoprene Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Approximately 550 million tonnes of the monoterpene, isoprene, are emitted to the atmosphere annually, principally from terrestrial plants. In contrast to methane, which is emitted in similar quantities, little is known about the biodegradation of isoprene. However, 30 years ago, bacteria capable of living on isoprene as a sole source of carbon and energy were described, although they were not investigated in detail. Recently there has been renewed interest in the potential of bacteria living in soils, marine sediments, and on the leaves of plants to degrade isoprene. Isolates capable of isoprene metabolism use a multicomponent soluble monooxygenase, which contains a diiron center at the active site, to oxidize isoprene to the epoxide, and all isolates described to date depend on glutathione for subsequent metabolic A. T. Crombie (*) School of Biological Sciences, University of East Anglia, Norwich, UK e-mail: [email protected] N. L. Mejia-Florez School of Environmental Sciences, University of East Anglia, Norwich, UK T. J. McGenity School of Biological Sciences, University of Essex, Colchester, UK J. C. Murrell School of Environmental Sciences, University of East Anglia, Norwich Research Park, Norwich, UK # Crown 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_27

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steps. The diversity of isoprene degraders has been investigated in terrestrial and marine environments using DNA-stable isotope probing (DNA-SIP), together with the use of gene probes targeting the monooxygenase active-site subunit. Gaps in our knowledge and future research directions are described.

1

Introduction

Isoprene (2-methyl-1,3-butadiene) is one of the most highly produced biogenic volatile organic compounds (BVOCs) emitted to the atmosphere, accounting for approximately 550 Tg C y 1, or 1/3 of total BVOCs (Guenther et al. 2006, 2012). This is similar in magnitude to the release of methane, with all other BVOCs comprising the remaining third. Isoprene is volatile, with a boiling point of 34  C and can be considered as a trace gas; its high reactivity in the atmosphere has a major influence on Earth’s climate. The effects of isoprene on climate are complex and not fully understood. It reacts with hydroxyl radicals, reducing the oxidative capacity of the atmosphere and resulting in a slower turnover of methane, a potent greenhouse gas, leading to global warming. Isoprene also reacts with oxides of nitrogen in the atmosphere, resulting in increased ozone levels, with effects on air quality and health (Sanderson et al. 2003; Pacifico et al. 2009). The oxidation products can result in secondary organic aerosols which promote increased cloud formation and global cooling (Carlton et al. 2009). Therefore under different circumstances, isoprene can act as both a global warming and a global cooling gas. Isoprene is a key building block for isoprenoids, which consist of two or more isoprene units and are produced by all free-living organisms. This large family of molecules includes, for example, carotenoids, sterols, chlorophyll, quinones, archaeal lipids, and hopanoids. The precursor molecules for isoprenoids are dimethylallyl diphosphate (DMAPP) and isopentenyl diphosphate (IPP), which are synthesized in animals, fungi, archaea, some bacteria, and the cytosol of plants using the mevalonate (MVA) pathway or by the methylerythritol phosphate (MEP) pathway in chloroplasts and most bacteria (Rohmer 1999). The vast majority of isoprene (approximately 90%) is produced globally by terrestrial plants by the action of isoprene synthase on DMAPP in chloroplasts (Sharkey et al. 2008). Interestingly, not all plants produce isoprene, with both high and low producers, even among closely related species – for example, all American oaks emit isoprene whereas many European oaks do not (Loreto et al. 1998). High isoprene-emitting trees typically divert 2% of fixed carbon to isoprene production, and in some cases considerably more (Sharkey et al. 1996), so it is striking that the reasons for this outlay of carbon and energy are not fully understood. There is good evidence that isoprene can alleviate heat and oxidative stress (Sharkey et al. 2008; Zeinali et al. 2016) and other proposed roles for isoprene biosynthesis include plant signaling, prevention from herbivory and as a way to dissipate excess energy from photosynthesis (Magel et al. 2006; Loivamäki et al. 2008). Certain emerging crop plants, for example palm oil, are high-isoprene

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emitters, and there is an increasing interest in the effects on air quality of the development of isoprene-emitting agroforestry (Hewitt et al. 2009). The remaining 10% of isoprene produced in the biosphere is attributed to bacteria, fungi, algae, and animals in both terrestrial and aquatic environments (Gelmont et al. 1981; Fares et al. 2008; Bäck et al. 2010; Exton et al. 2015). For example, Bacillus subtilis has been shown to produce isoprene, maybe as a consequence of stress in this bacterium (Kuzma et al. 1995). In the marine environment, macro- and microalgae are the major producers, responsible for a poorly constrained flux of 0.1–10 Tg C y 1 (Palmer and Shaw 2005; Luo and Yu 2010; Shaw et al. 2010; Srikanta Dani et al. 2017). As seen with plants, isoprene emissions by marine microalgae increase in response to higher temperature and light intensity and so may protect these organisms during periods of stress (Exton et al. 2013). Isoprene is also produced industrially (approximately 0.8 Tg y 1) and used primarily for polyisoprene elastomer (synthetic rubber) production (Morais et al. 2015). Since little is known about bacterial isoprene synthases, strategies have been developed to express isoprene synthase genes from plants in heterologous systems, including Escherichia coli, Saccharomyces, and Synechocystis (Marienhagen and Bott 2013; Lv et al. 2014). In comparison to the global methane cycle, information regarding the biogeochemical cycling of isoprene is rather sparse and there are many unknowns, particularly estimates of both production and consumption in the biosphere and the internal recycling of isoprene in soils, on the surface of plants, and in the marine environment. Microbial production and/or consumption of isoprene have previously been reviewed by Fall and Copley (2000), Shennan (2006), and McGenity et al. (2017). The global isoprene cycle is shown in Fig. 1. Here we review the mechanisms by which microbes degrade isoprene, and their diversity in the environment.

Fig. 1 Isoprene is emitted to the atmosphere by trees, plants, algae, animals, fungi, and bacteria in the terrestrial and marine environment (black arrows), where it is rapidly photochemically oxidized (blue arrows). Isoprene may also be taken up from the atmosphere by microbes in soils and aquatic ecosystems and sediments (grey arrows). Consumption of isoprene by microbes at the point of release is shown as red circular arrows

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Microbial Consumption of Isoprene

Soils have been recognized as a sink for isoprene for over 20 years. In the terrestrial environment, temperate, tropical, and boreal soils were shown to rapidly consume isoprene at concentrations of 500 ppbv in laboratory-based experiments. Field chamber experiments in temperate forests also revealed that soils could deplete isoprene added to chambers at ~400 ppbv down to below the detection limits of about 5 ppbv within 1 h (Cleveland and Yavitt 1997, 1998). Temperate agriforest and model tropical rainforest mesocosm experiments revealed rapid in situ consumption of isoprene (Pegoraro et al. 2005, 2006). More recently, continuous flow experiments with temperate forest soils revealed that these systems consume isoprene over a range of concentrations (2–200 ppbv), with substantial rates of isoprene removal even at low (20 ppbv) concentrations (Gray et al. 2015). The first demonstration of microbial consumption of isoprene in the marine environment was by Acuña Alvarez et al. (2009), who observed isoprene-degrading bacteria in estuarine, coastal, and open marine waters. They also demonstrated that isoprene produced by marine microalgae cultures could be consumed by isoprene-degrading bacteria, an important observation proving that marine microbes could benefit directly from isoprene, produced by microalgae and without using artificially high laboratory concentrations. This study also resulted in the isolation of a number of isoprenedegrading bacteria from these marine environments (see later).

3

Bacterial Degradation of Isoprene

The first reports of isolation of bacteria growing on isoprene as sole carbon and energy source were by van Ginkel et al. (1987a, b), Ewers et al. (1990), and Cleveland and Yavitt (1997). Soil enrichments with isoprene under aerobic conditions yielded bacteria assigned to the actinobacterial genera Rhodococcus, Nocardia, and Arthrobacter and to the proteobacterial genus Alcaligenes. More recently aerobic isoprene-degrading Pseudomonas, Klebsiella, and Alcaligenes strains have been reported (Srivastva et al. 2015), but, as with earlier studies, these have not been characterized in any detail. Our recent work has provided collections of aerobic isoprene-degrading bacteria for further study with a number of Rhodococcus species being isolated from soils and the leaves of isoprene-producing trees such as Poplar and Willow (El Khawand et al. 2016; Murphy 2017). Marine sediments have also yielded a variety of Gram-positive and Gram-negative aerobic isoprene degraders (Acuña Alvarez et al. 2009; Johnston et al. 2017) (Fig. 2). To our knowledge, no anaerobic bacteria, archaea, or fungi growing on isoprene have yet been reported. The most well-characterized isoprene-degrading bacterium described so far is Rhodococcus sp. AD45, isolated from freshwater sediment by Janssen and colleagues (van Hylckama Vlieg et al. 1998). In this aerobe, the initial oxidation of isoprene to 1,2-epoxyisoprene is carried out by the enzyme isoprene monooxygenase (IsoMO), a multicomponent soluble diiron monooxygenase (SDIMO) belonging to the same large family of enzymes as soluble methane monooxygenase,

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Fig. 2 The 16S rRNA gene-based phylogeny of isoprene degraders (shown in bold) together with other representative strains. The tree was drawn using the Neighbor Joining method in Mega6 (Tamura et al. 2013) with pairwise deletion, resulting in 1594 nucleotide positions in the analysis. Bootstrap values (1000 replications) greater than 75% are shown by black circles at the nodes. The scalebar shows base substitutions per site

toluene monooxygenase, and alkene monooxygenase (Leahy et al. 2003; Holmes and Coleman 2008). The reactive epoxide produced in the first step of isoprene metabolism is conjugated with glutathione by a glutathione-S-transferase to form

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Fig. 3 The isoprene degradation pathway. HGMB, 1-hydroxy-2-glutathionyl-2-methyl-3-butene; GMB, 2-glutathionyl-2-methyl-3-butenal; GMBA, 2-methyl-2-glutathionyl butenoic acid; GS, glutathione.

1-hydroxy-2-glutathionyl-2-methyl-3-butene (HGMB). HGMB is further metabolized by a dehydrogenase to glutathionyl-2-methyl-3-butenoate (GMBA) (van Hylckama Vlieg et al. 1998, 1999). The subsequent fate of GMBA remains to be elucidated but it can be assumed that the subsequent removal of the glutathione moiety and β-oxidation of the intermediates of isoprene metabolism allows Rhodococcus sp. AD45 to grow on isoprene as a sole carbon and energy source (Fig. 3). van Hycklama Vleig et al. (2000) showed that the isoprene monooxygenase is encoded by the genes isoABCDEF. Subsequent sequencing of the 6.8 Mbp genome of Rhodococcus sp. AD45 (Crombie et al. 2015) revealed that all of the genes necessary for the metabolism of isoprene are carried on a 300 kbp megaplasmid in this Rhodococcus strain (Fig. 4). This clustering of isoprene metabolic genes seems to be a common feature in all isoprene degraders studied to date, although in some other isoprene-degrading Rhodococcus strains (e.g., Rhodococcus opacus PD630) these isoprene-related genes appear to be chromosomally located rather than on a plasmid (Chen et al. 2014; Crombie et al. 2015). Genes isoA, isoB, and isoE encode

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Fig. 4 The isoprene metabolic gene cluster from representative terrestrial and marine strains. The isoprene monooxygenase genes are shown in red, and homologous genes are in corresponding colors

the diiron (α2β2γ2) oxygenase component; isoF encodes a flavoprotein NADH reductase; isoC encodes a Rieske-type ferredoxin; and isoD encodes a coupling protein, which together form the multicomponent isoprene monooxygenase. The four genes isoGHIJ, preceding the IsoMO genes, encode a putative coenzyme A transferase, a dehydrogenase, and two glutathione transferases (van Hylckama Vlieg et al. 1998, 1999, 2000). An additional copy of this isoGHIJ cluster is located on the Rhodococcus sp. AD45 megaplasmid (Crombie et al. 2015), and these and other isoprene metabolic genes are duplicated in many isoprene degraders (Fig. 4). Upstream of isoG, gshA encodes glutamate cysteine ligase, which catalyzes the first step of glutathione biosynthesis. Two copies of gshB, encoding glutathione synthetase, are also present. The cluster also contains genes predicted to encode two aldehyde dehydrogenases (aldh1 and ald2), both of which are present in the genomes of all other sequenced isoprene degraders. In Rhodococcus opacus PD630, the former of these, aldh1, was shown to have glyceraldehyde-3-phosphate and NADP+-dependent activity (MacEachran and Sinskey 2013), possibly suggestive of a requirement for NADPH in isoprene metabolism, although, to our knowledge, alternative substrates of this enzyme were not investigated in detail. Genes encoding a putative coenzyme A disulfide reductase and three putative transcriptional regulators (marR1, marR2, and gntR) are also present. These iso genes are essential for isoprene metabolism since removal of the megaplasmid of Rhodococcus sp. AD45 by “curing” resulted in loss of the ability to grow on isoprene, as did deletion of isoA by mutagenesis (unpublished observation; Crombie et al. 2015). Actinobacteria such as Rhodococcus are metabolically versatile and it appears that isoprene metabolism in Rhodococcus sp. AD45 and other isoprene degraders tested to date (Gordonia and Mycobacterium) is an inducible trait (Crombie et al. 2015; Johnston et al. 2017). Carbon sources such as glucose and succinate repress isoprene metabolism but isoprene monooxygenase polypeptides are readily observed in cell extracts of isoprene-grown Rhodococcus sp. AD45. The regulation of expression of isoprene metabolic genes in this strain has been studied in some detail. A replicated time course experiment was conducted in which succinate-grown cells were subcultured into medium containing isoprene, epoxyisoprene, glucose, succinate, or no substrate (Crombie et al. 2015).

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Fig. 5 Induction of representative isoprene-degradation genes in Rhodococcus sp. AD45 after incubation with isoprene, at six timepoints from zero to 25 h, quantified by RNAseq. The bar chart shows the fold change at each time point relative to controls incubated without carbon substrate (Data from Crombie et al. 2015)

Isoprene- and epoxyisoprene-induced expression of iso genes were examined by sequencing the transcriptome and comparison with cells grown on glucose and succinate (at 0, 19, 43, 75, 240 min and 25 h). Both isoprene and epoxyisoprene induced high levels of expression of isoABCDEF, isoGHJI, and all other putative isoprene metabolism genes present in the 22-gene cluster on the megaplasmid of Rhodococcus sp. AD45 (Fig. 5). Under isoprene-induced conditions, these genes represented over 25% of all transcripts observed. These results were subsequently confirmed by targeted RT-PCR. There was no significant over-expression of genes on the chromosome in response to isoprene or epoxyisoprene, suggesting that all isoprene metabolic genes reside on this plasmid (Crombie et al. 2015). The dynamics of gene transcription in response to isoprene or epoxyisoprene indicated that the inducer was not isoprene itself, but rather epoxyisoprene or a subsequent metabolic product of isoprene oxidation, subsequently confirmed by transcriptional analysis of a mutant with an inactivated IsoMO in which isoprene (which could not be metabolized) did not induce iso gene transcription. These data provide further targets for mutagenesis and expression studies in order to elucidate the full pathway of isoprene metabolism in Rhodococcus sp. AD45. The use of reporter strains to analyze transcriptional regulation and analysis of putative regulators encoded by marR and gntR may also be a fruitful approach.

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Ecology of Isoprene Degraders

Analysis of the genomes of a number of other isoprene-degrading Rhodococcus, Gordonia, Mycobacterium, and Variovorax strains has revealed that the clustering of isoABCDEF along with isoGHIJ and genes involved in glutathione biosynthesis seems to be a common feature in isoprene degradation (Fig. 4) (Crombie et al. 2015; El Khawand et al. 2016; Johnston et al. 2017). The genes encoding isoprene monooxygenase are often misannotated as toluene or alkene monooxygenases, other members of the SDIMO family, but the close proximity of isoGHIJ to isoABCDEF is a good indication of isoprene metabolism in newly isolated strains. The availability of the isoABCDEF sequence has also provided tools for cultivation-independent studies aimed at assessing the distribution, diversity, and activity of isoprene degraders in the environment. A fruitful approach in molecular ecology studies of methane-oxidizing bacteria has been to use methane monooxygenase marker genes (“functional genes”) encoding key components of methane monooxygenases (pmoA or mmoX) to examine different environments (Dumont and Murrell 2005b; McDonald et al. 2008). We have used a similar approach with isoprene monooxygenase. The homologue of mmoX (which encodes the large subunit of the oxygenase component of the soluble methane monooxygenase) in IsoMO is isoA, which encodes the putative active site component. This polypeptide appears to be highly conserved in all isoprene degraders studied to date. Phylogenetic analysis of IsoA of isoMO from known isoprene degraders and comparison with the corresponding components of SDIMO enzymes such as toluene monooxygenase and alkene monooxygenases (Fig. 6) has confirmed that isoA is a suitable marker gene for cultivation-independent studies and that derived IsoA homologs can be readily distinguished from those of non-isoprenedegraders containing SDIMOs other than IsoMO. Alignment of IsoA from bona fide isoprene degraders has allowed the design of isoA PCR primer sets targeting these bacteria. These primers did not amplify SDIMO genes from non-isoprene degraders but gave isoA gene products with DNA from a range of isoprene-degrading isolates and enrichment cultures originating from various soils, sediments, and leaf samples (El Khawand et al. 2016). Alignments of the IsoA sequences retrieved from environmental samples, with those of characterized isoprene degraders, showed that the IsoA sequences were relatively highly conserved (>86% identity) and could be broadly separated into two groups, those from marine environments and those from terrestrial environments, predominantly actinobacterial isoprene degraders (El Khawand et al. 2016). However, the subsequent isolation and analysis of more isoprene degraders suggests that there is variation in their isoA sequences which will, as has been the case with methane monooxygenase functional gene PCR primers, necessitate redesign of isoA PCR primer sets. This functional gene probing approach has extended knowledge of the diversity of isoprene-degraders in both terrestrial and marine environments (El Khawand et al. 2016; Johnston et al. 2017). In order to determine which isoprene degraders are active in the environment, other cultivation-independent techniques such as DNA Stable Isotope Probing

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Fig. 6 The relationship between the isoA gene sequences of isoprene degraders (shown in bold) and other homologous sequences. The tree was drawn in Mega6 (Tamura et al. 2013) using the Maximum Likelihood method. Sites with less than 95% coverage were deleted, resulting in 1005 nucleotide positions in the analysis. Bootstrap values (1000 replications) greater than 75% are shown by black circles at the nodes. The scalebar shows base substitutions per site. Based on the presence of adjacent isoGHIJ genes, we can predict that Rhodococcus sp. JVH1 can grow on isoprene, although this has not, to our knowledge, been tested

(DNA-SIP) (Dumont and Murrell 2005a) need to be used. DNA-SIP has been used to identify active isoprene degraders in both the terrestrial and marine environment. For example, surface soil samples from around Willow trees (Salix fragilis) were supplied with 13C–labelled isoprene in microcosms and isoprene uptake was monitored by gas chromatography. After sufficient “heavy” isoprene was incorporated into biomass, 13C–labelled DNA was isolated by buoyant density gradient centrifugation and used as template in PCR reactions with primers targeting 16S rRNA genes. The analysis revealed a considerable enrichment of several species of Rhodococcus in these microcosms, indicating that under the incubation conditions used, Rhodococcus represented the majority of active isoprene degraders in these soils (El Khawand et al. 2016). Interestingly, 16S rRNA gene sequences from the Betaproteobacteria Comamonas and Variovorax were also enriched in 13 C–labelled DNA. In addition to soils, DNA-SIP has also been used to identify active isoprene degraders on leaves of White Poplar (Populus alba). Microbes washed from the leaves of this tree were incubated in microcosms with

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C–labelled isoprene. After sufficient 13C–labelling, “heavy” DNA was isolated and used in shotgun metagenomics experiments. This enabled the reconstruction of a considerable proportion of the draft genome of an isoprene-degrading Variovorax species, including the iso metabolic gene clusters isoABCDEF and isoGHIJ (Crombie et al., manuscript in preparation). This information has subsequently been used in the targeted isolation of isoprene-degrading Variovorax species (Mejia-Florez et al., unpublished). The functionality of these Variovorax iso genes retrieved directly from the environment using DNA-SIP was confirmed by expression studies. The putative IsoMO genes isoABCDE from the reconstructed Variovorax genome were expressed in a heterologous expression system: a non-isoprene-degrading variant of Rhodococcus sp. AD45 without the 300 kbp megaplasmid carrying the iso genes (Crombie et al., manuscript in preparation). When expressed, the Variovorax isoABCDEF genes conferred the ability of the Rhodococcus strain to oxidize isoprene, thus proving that it is a bona fide isoprene monooxygenase. Metatranscriptome data obtained from the same isoprene incubation experiments used for DNA-SIP also confirmed that these Variovorax genes were expressed under the enrichment conditions. Targeted isolations have now yielded isoprenedegrading Variovorax strains from leaves, thus providing further isoA genes to refine PCR primer sets and a new model Gram-negative isoprene degrader to complement the Gram-positive strains available (Crombie et al. manuscript in preparation). The diversity of isoprene degraders in the marine environment has also been investigated using cultivation-independent methods. Surface estuarine sediments from the Colne Estuary (UK), incubated with 13C–labelled isoprene, yielded 13 C–DNA which when analyzed revealed the development of isoprene-degrading communities dominated by Actinobacteria including Gordonia, Mycobacterium, Microbacterium, and Rhodococcus (Johnston et al. 2017). Enrichments of similar environmental samples from the Colne Estuary yielded isolates Gordonia sp. i37 and Mycobacterium sp. AT1, which grew on isoprene as a sole source of carbon and energy. Analysis of their genomes revealed the same gene arrangements of isoABCDEF and isoGHIJ seen in Rhodococcus sp. AD45 (Fig. 4). As with R. sp. AD45, isoprene oxidation was inducible in the presence of isoprene (Johnston et al. 2017). A second SDIMO was identified in the genomes of both of these isoprene-degraders which had significant sequence identity to the propane monooxygenase from Gordonia TY-5 (Kotani et al. 2003). This second SDIMO system enabled these bacteria to grow on propane, which, interestingly, appears to be a common feature of many isoprene-degrading bacteria (although not Rhodococcus sp. AD45) (Acuña Alvarez et al. 2009; Johnston et al. 2017). These marine isolates, together with other isoprene degraders isolated and characterized from the Colne Estuary and other marine environments (Acuña Alvarez et al. 2009; Johnston et al. 2017), are yielding valuable genome sequence information to refine isoA PCR primers and confirming the presence of both isoABCDEF and isoGHIJ gene clusters in bona fide isoprene degraders. 13

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Conclusions and Research Needs

All isoprene-degrading bacteria studied so far possess an isoprene monooxygenase of the SDIMO family, which is required for the initial oxidation of isoprene. This enzyme is induced by isoprene or a further oxidation product of isoprene metabolism. It will be interesting to explore the structure and substrate range of this new class of enzymes in relation to others in the SDIMO family; for example, soluble methane monooxygenase found in some methanotrophs and alkene monooxygenases of propene degraders. These enzymes can also (co-)oxidize isoprene but the bacteria lack the additional metabolic machinery to allow growth on isoprene (Johnston et al. 2017). A second unifying feature of extant isoprene-degraders is the use of glutathione to detoxify epoxyisoprene, the first oxidation product of isoprene. This is in contrast to other alkene degraders, which often use coenzyme M as cofactor (Krishnakumar et al. 2008), and glutathione biosynthesis genes have so far always been found in close proximity to iso genes in isoprene degraders. The subsequent steps in isoprene metabolism require further study but the identification of iso genes in Rhodococcus sp. AD45 and the availability of a mutagenesis and expression system in this “workhorse” organism will now allow us to characterize the mechanisms by which bacteria regulate isoprene metabolism and subsequently incorporate carbon from isoprene into biomass. In some cases (Rhodococcus sp. AD45), all necessary iso genes reside on a megaplasmid, hinting at the possibility that they are transferred between bacteria by horizontal gene transfer. This notion is supported by the lack of congruence between the phylogeny derived from isoA and 16S rRNA gene sequences (Figs. 1 and 6). The studies summarized here clearly indicate that isoprene-degrading bacteria are widespread in the environment and that soils possess the capability to deplete isoprene at environmental concentrations. Cultivation-independent techniques, such as DNA-, RNA- or protein-SIP, or single cell technologies such as Raman microspectroscopy, will help reveal new isoprene degraders (Murrell and Whiteley 2011; Wang et al. 2016). Challenges for the future include conducting these sequence-independent experiments at conditions that mimic those in the environment, which will identify isoprene-degrading microbes which may exploit specific micro niches. Given that intercellular isoprene concentrations inside leaves may be up to three orders of magnitude higher than atmospheric (Fini et al. 2017), reaching the low ppmv range, the possibility of isoprene-degrading endophytes should be explored. It will also be necessary to increase the diversity of isoprene-degrading strains in cultivation. Characterization of isolates may reveal other pathways for metabolism of isoprene by bacteria and indeed may identify other isoprene-degrading microbes such as Archaea and fungi. To our knowledge there is so far no evidence for anaerobic degradation of isoprene but this possibility should also be borne in mind. To determine the impact that biological isoprene uptake has on global fluxes it will be necessary to quantify microbial activity in the environment. Approaches being pursued in our laboratory range from construction of biosensor strains to express reporter genes under conditions where isoprene-related genes are expressed, through to purification and characterization of isoprene metabolic enzymes, including IsoMO, to determine the affinity and kinetics of isoprene degradation in different isolates.

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Acknowledgments The authors acknowledge funding from the Earth and Life Systems Alliance (ELSA) at the Norwich Research Park, a Natural Environment Research Council (NERC) grant to JCM (NE/J009725/1) and TJM (NE/J009555/1), an ERC Advanced Grant to JCM (694578 – IsoMet), and Colciencias (Government of Colombia) and Newton Fund support for a studentship to NLMF.

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Current View of the Mechanisms Controlling the Transcription of the TOL Plasmid Aromatic Degradation Pathways

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Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 The Upper Pathway: Onset of the Regulatory Cascade . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 The Meta-Cleavage Pathway: Current Understanding of the Activation Mechanism . . . . . 3.1 New Insights into the Cross Regulation of the Meta- and Ortho-Cleavage Pathways for Benzoate Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 A Family of Anti-activators Accompanying XylS/AraC Regulators . . . . . . . . . . . . . . . . 4 Integration in the Cell Regulatory Networks: Toward Optimization of Expression . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

The TOL plasmid-encoded pathway for the degradation of toluene and derivatives is an archetype in bacterial transcription regulation. Six promoters belonging to different classes 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 σ54-dependent promoters of the upper pathway Pu and of xylS gene PS1, and the AraC family regulator XylS, which controls the σ32-σ38-dependent meta-cleavage pathway promoter Pm. Both regulators respond to the presence of a specific effector and activate transcription through different

P. Domínguez-Cuevas Department of Biology, University of Copenhagen, Copenhagen, Denmark S. Marqués (*) Department of Environmental Protection, CSIC, Estación Experimental del Zaidín, Granada, Spain e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_29

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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 and their interrelationships with the cell metabolism.

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 (Fig. 1a) (Greated et al. 2002). 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 σ54-dependent promoters, the upper pathway promoter Pu and the xylS gene promoter PS1 (reviewed in Ramos et al. 1997). Activation of PS1 leads to increased synthesis of XylS, which in these conditions 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 (Marqués 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 substituted derivatives that activate XylS, a regulatory pathway known as the metaloop (Ramos et al. 1986; Inouye et al. 1987). Furthermore, 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, and following a few minutes delay, this aromatic activates the XylS regulator to promote increased transcription from the meta-cleavage pathway promoter Pm (Marqués et al. 1994). A recent transcriptomic analysis of TOL pathway gene expression confirms the basic steps in this regulatory network and exposes previously overlooked subtleties of the pathway operon mRNA synthesis (Kim et al. 2016). The resulting synthesis of the pathway enzymes, however, takes several hours to reach maximum steady levels (Hugouvieux-Cotte-Pattat et al. 1990). The extensive analysis of these processes shows that a considerable number of chromosome- and plasmid-encoded proteins cooperate to maintain an optimal pathway expression level in every circumstance (Table 1). The general patterns of this network have already been revised (Marqués and Ramos 1993; Ramos et al. 1997; Ruíz et al. 2004; Daniels et al. 2008; Domínguez-Cuevas and Marqués 2010). The intricacy and refinement of the regulatory network allow for an appropriate synchronization with the global cell metabolism. Upon the well-established effector-dependent specific control mechanisms, increasing regulatory steps connecting the TOL pathway with the host metabolic

Current View of the Mechanisms Controlling the Transcription of the TOL. . .

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Fig. 1 (a) The pWW0 TOL plasmid aromatic degradation pathway is organized in two operons and two regulatory genes included in two transposable elements inserting one within another. (b) Schematic presentation of XylR modular organization into three functional domains: a sensor domain (A), connected by a linker (B) to the central or AAA+ domain (C), and a DNA-binding domain (D). (c) Schematic presentation of the XylS protein functional domains: effector binding and dimerization domain and DNA-binding domain containing two HTH motives. The residue numbers delimiting each domain are indicated in each case

network are continuously being uncovered, reflecting the complexity and fine control of the pathway. This makes of the TOL system an archetype in bacterial transcription regulation, which has been exploited in fundamental and applied research fields. 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.

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 σ54-dependent promoter activator family known as bacterial enhancer-binding proteins (bEBPs). These proteins consist of three main domains: an N-terminal sensor domain (A) sensing the regulatory signal (e.g., the presence of an effector in the case of XylR (Delgado and Ramos 1994), connected through a B-linker to a central AAA+ domain (C) with ATPase activity, and a C-terminal DNA-binding domain (D) (Fig. 1b) (Bush and Dixon 2012). The role of the three main domains has been well established in XylR. Genetic analysis located the mutations altering the effector profile in the N-terminal domain (NTD) of the protein (Delgado and Ramos 1994; Salto et al. 1998; Garmendia et al. 2001, 2008). As for other bEBPs where the N-terminal domain exerts a negative control, a XylR protein devoid of the NTD shows constitutive activity (Pérez-Martín and De Lorenzo 1995a; Garmendia and de Lorenzo 2000). This domain given in trans exerts

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specific intramolecular repression, inhibiting ATP-binding capacity of the C domain; repression is released in the presence of an effector (Fernández et al. 1995; PérezMartín and De Lorenzo 1995a). Although in XylR physical interaction with the effector has not been analyzed, effector binding analysis in the phenol-responsive XylR homologue DmpR revealed that actually the interaction of labeled phenol with the N-terminal domain released C-domain ATPase activity repression (Shingler and Pavel 1995). Direct interaction with different effectors was proven by isothermal titration calorimetry (ITC) in MopR, a phenol-responsive XylR-DmpR homologue (Ray et al. 2016). Early structure prediction based on the alignment of 11 XylR family proteins and homology modeling suggested a fold similar to V4R domain (Pfam 02830) (Devos et al. 2002). This predicted structure was recently endorsed by the crystal structure of MopR NTD, the first structure available for a sensor domain of the XylR-NtrC subfamily of bEBPs (Ray et al. 2016). The NTD of both XylR and DmpR can be modeled using MopR structure, showing that the actual XylR residues involved in direct interaction with the effector are different from the previously suggested from genetic analysis. The specific role of several previously mutated residues could be inferred from the protein structure. Unexpectedly, the MopR crystal structure revealed the presence of a Zn-binding domain that was conserved in XylR and DmpR. According to the proposed model, this Zn-binding domain, together with the B-linker, would be directly involved in opening and closing an effector gate to the binding site and in triggering communication with the central domain, as suggested for several bEBPs (Bose et al. 2008; Ray et al. 2016). This is consistent with the previous suggestion, based on mutant analysis, of XylR B-linker influencing effector binding (Garmendia and de Lorenzo 2000). Structural analyses of several bEBPs show that this domain also regulates multimerization and couples the A-domain input signal with the 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 this regulator. The AAA+ central domain (C) is the most conserved among bEBPs. It carries the ATP-binding motif, ATPase activity, and σ54 interaction determinants, features that are essential to σ54 promoter activation (Schumacher et al. 2006; Chen et al. 2008). 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 (Pérez-Martín and de Lorenzo 1996a). The recent availability of the crystal structure of several regulators of the family either devoid of their NTD or naturally lacking this domain has helped 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; reviewed in Bush and Dixon 2012). Although the mechanistic model has only been proposed for those members belonging to a two-component system or for those naturally lacking an NTD, XylR is likely to share many of their features. Proteins of this family generally bind DNA as dimers. After the A-domain repression has been released (in XylR,

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effector binding would trigger A-domain movement and initiate derepression), 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 binding and hydrolysis, thus explaining the previously observed ATPase activity dependence on protein oligomerization (Bordes et al. 2003; reviewed in Schumacher et al. 2006). Two C-domain 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 σ54 (Bose et al. 2008; Wigneshweraraj et al. 2008). Mutations in these loops in XylR abolish ATP binding and hydrolysis (Pérez-Martin and de Lorenzo 1996a). Contacts between the regulator and the transcriptional machinery require that the two protein complexes are brought into proximity by DNA bending, generally facilitated by the integration host factor (IHF). Finally, σ54 undergoes a structural remodeling, allowing the σ54-RNA polymerase (Eσ54) closed complex to isomerize to an open complex. As in all bEBPs, in XylR 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). Comparison of the structure of the HTH domains of a series of bEBPs shows that they all share a similar DNA recognition element formed by a three-helical bundle, where the C-terminal helix is the DNA-contacting element (Vidangos et al. 2013). This structure is conserved in XylR structure model. XylR dimers are always bound to the two upstream activation sequences (UASs) in Pu (Abril et al. 1991). As discussed above for other bEBPs, binding of ATP induces multimerization to a hexameric conformation, which is then able to hydrolyze ATP (Pérez-Martín and de Lorenzo 1996b). Binding of bEBPs to their UASs is thought to generate high local concentrations of activated regulator in the proximity of Eσ54-promoter complex. 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 (Velázquez 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 RNA polymerase (RNAP) machinery bound at
12/
24 (de Lorenzo et al. 1991). Early models explaining bEBP activation mechanism suggested direct interaction between the regulator and Eσ54, in most cases brought to proximity by IHF-assisted DNA looping. Interestingly, the regulator approaches the Eσ54 closed complex through the unbound face of Eσ54 binding site to contact σ54 and catalyze open complex formation, so that DNA appears sandwiched between Eσ54 and the regulator (Huo et al. 2006; Wigneshweraraj et al. 2008). Despite the profusion of bEBP structures and biochemical analysis available in the past years, the precise mechanism underlying energy coupling from regulator ATP hydrolysis to promoter melting and escape is not yet

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fully understood (see Bush and Dixon 2012 for a comprehensive review on bEBP structure and activation mechanism). The structure of the PS1 promoter slightly differs from the canonical σ54-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 Eσ70 binding sites of the two xylR promoters PR1 and PR2. Two consensus IHF binding sequences are found, overlapping the
12/
24 Eσ54 binding site and the UASs. As a consequence of this complex organization, expression of XylR regulator never achieves maximum levels (Marqués et al. 1998): when XylR binds its UASs to activate PS1, it represses its own synthesis (Holtel et al. 1992; Bertoni et al. 1998). Self-repression of XylR has been suggested to help buffering possible fluctuations in XylR synthesis levels in any condition (Koutinas et al. 2010); in fact, xylR gene expression levels were shown to be similar along the growth curve in the presence and absence of effector (Marqués et al. 1994). In contrast to Pu activation mechanism, the presence of IHF strongly represses PS1 activity (Marqués et al. 1998), which probably explains why PS1 expression levels have been estimated to be fourfold lower than Pu expression levels (Marqués et al. 1994). In PS1, the histonelike protein HU replaces the positive function of IHF by increasing a preexistent static bend in the DNA to bring into proximity the DNA-bound complexes of the regulator and Eσ54 (Pérez-Martín and de Lorenzo 1995b).

3

The Meta-Cleavage Pathway: Current Understanding of 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: σ32 (RpoH) in the early exponential phase and σ38 (RpoS) in the late exponential and stationary phases (Marqués et al. 1999). In fact, Pm
10/
35 RNAP binding sequence diverges considerably from the consensus defined for σ32, σ38, and σ70 factors (Domínguez-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 (González-Pérez et al. 2002); on the other hand, the
10 region of Pm must include the essential determinants for recognition by the two polymerases involved, Eσ32 and Eσ38. Unlike σ70, 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 σ70 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 (DomínguezCuevas et al. 2006). 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 (Domínguez-Cuevas et al. 2006). The increase in

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σ32 level during the heat shock response is strong but transient, so the Pm promoter sequence is optimized to accommodate the temporary Eσ32 RNAP. After this initial heat shock response, a second alternative RNAP associated with the stress/stationary sigma factor σ38 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 (Fig. 2). 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; González-Pérez et al. 2002). XylS belongs to the AraC family of transcription regulators (Gallegos et al. 1997; Tobes and Ramos 2002) and is composed of two separate functional domains; XylS mutants with altered effector specificity cluster in the N-terminal domain of the protein, indicating that this domain carries the effector recognition determinants (Ramos et al. 1990; Michán et al. 1992; Ruíz 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 (Ruíz 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 (Fig. 1c). It has been recently reported that in AraC the inter-domain linker plays an active role both in protein dimerization and affecting the interaction between the COOH

CH3 R

CH3

R

Pu

activation XylR

σ54

XylR

upper

Eσ54

Effector

P S1

xylS RNAP core

overproduction

CH3

XylS

activation

XylS

XylS

COOH

XylS

CH3

XylS

meta-cleavage

Pm

Toluene

XylS

COOH

Effector

activation

XylS

Permanent Transient

XylS

CH3 3-methylbenzoate

Stress

Fast Slow

TCA cycle

Heat Shock Response 38

↑σ

↑σ

32

Eσ32 Eσ38

Fig. 2 Effector dual function in Pm activation. Red arrows indicate a direct positive effect on transcription. Light-green arrows indicate activation of the regulator. Black upward arrows indicate accumulation of a sigma factor

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effector/dimerization and the DNA-binding domains (Seedorff and Schleif 2011). X-ray crystallographic structures of other AraC family proteins show that the DNA-binding domain is composed of seven α-helices folding in two HTH domains, which bind two adjacent segments of the major groove (Rhee et al. 1998; Kwon et al. 2000). 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 (Domínguez-Cuevas et al. 2008b). Interestingly, newly reported data have shown that another member of the AraC/XylS family, VirF, is synthesized from two alternative translational start sites. The two produced protein forms play different functional roles; the full-length form activates Shigella spp. virulence system, while the short form, which includes the DNA-binding motif, negatively autoregulates virF expression itself (Di Martino et al. 2016). Intriguingly, reexamination of XylS openreading frame disclosed the existence of two putative internal ATG start codons, which, hypothetically, could generate alterative shorter forms of the activator. Analysis of the XylS DNA-binding domain showed it binds DNA forming two complexes (CI and CII), corresponding to one or two XylS-C monomers bound to DNA, respectively (Domínguez-Cuevas et al. 2010). Affinity calculation, DNA bending angle estimation and footprinting assays of XylS C-terminal domain suggest the two monomers bind DNA sequentially and noncooperatively. The first XylS-C monomer binds Pm at the proximal site (closest to the RNAP binding site), raising Pm curvature 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 (DomínguezCuevas and Marqués 2010). Our results indicate that sugar-phosphate backbone contacts greatly contribute to XylS/Pm binding strength (Domínguez-Cuevas 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 (Domínguez-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 (Ruíz et al. 2003; Domínguez-Cuevas et al. 2008a). In fact, in the absence of inducer, expression from Pm increases as a function of XylS expression levels, reaching a maximum activity level that compares with the one under induced conditions (Zwick et al. 2013). These data support the proposal that XylS could exist in three states: monomers, dimers, and aggregates, being the dimeric form the only one active and able to induce transcription from the Pm promoter. This model suggests that XylS overproduction leads to the formation of inactive aggregates, which correlates with the absence of Pm induction at high intracellular XylS levels. As for the role of 3MB in XylS activation, data obtained with the two purified protein domains suggest intramolecular repression of XylS-NTD upon XylS-CTD DNA binding, which is released in the presence of 3MB (Domínguez-Cuevas et al. 2008a). Even though the mechanism for activation upon 3MB addition is not fully understood, these data are consistent with the role of arabinose during AraC

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activation. Upon arabinose binding, the AraC flexible arm restructures, which releases the DNA-binding domain from the dimerization domain, allowing transcription activation (Rodgers et al. 2009). 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 derepression 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 derepression is favored, unmasking dimerization surface and DNA-binding determinants, which leads to Pm recognition and further activation.

3.1

New Insights into the Cross Regulation of the Metaand Ortho-Cleavage Pathways for Benzoate Degradation

Pseudomonas putida mt-2 encodes two alternative pathways for benzoate metabolism, the aforedescribed meta pathway and the chromosomally encoded ortho-cleavage pathway. The ortho pathway operon is transcribed from the Pben promoter in response to the BenR regulator, member of the AraC-XylS family of activators. The TOL pathway allows mineralization of toluene and m-xylene through the upper pathway. The methyl group of toluene and m-xylene is sequentially oxidized to render, respectively, benzoate or 3-methylbenzoate. Non-substituted benzoates can be degraded productively both through the ortho- and meta-cleavage pathways; however, methylated benzoates channeled via the ortho-cleavage pathway generate dead-end metabolites known as methyl-2-enelactones. This metabolic conflict posed when P. putida mt-2 cells, encoding both pathways, degrade 3-methylbenzoate has been analyzed in depth to conclude that it is overcome through a rather simple and effective regulatory solution. The pWW0-encoded XylS regulator is highly homologous to the chromosomally encoded BenR protein, to the point that both regulators show a degree of cross regulation (Kessler et al. 1994; Cowles et al. 2000; Domínguez-Cuevas et al. 2006). The operator sequences found in Pm and Pben are highly similar, however, while Pm has two complete operator sequences for XylS, Pben apparently presents three out of the four DNA boxes required for physiological XylS binding (Pérez-Pantoja et al. 2015). The lack of two complete operator sequences in Pben is sufficient to prevent activation of transcription by physiological levels of XylS (Pérez-Pantoja et al. 2015; Tsipa et al. 2016). In fact, generation of Pben promoter variants with two complete operator sequences resulted in increased activation levels in response to both inducers (Silva-Rocha and de Lorenzo 2012), due to improved recognition of the Pben promoter by 3MB-activated XylS. In connection with the simultaneous presence of the ortho and meta pathways, recent published

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data suggest that P. putida mt-2 contains an additional metabolic safety valve, the chromosomal catA2 gene, which helps counteracting the toxicity of high catechol concentrations (Jiménez et al. 2014). However, the most unexpected finding of a recent transcriptomic analysis of TOL operons in cells growing with different inducers was that the unsubstituted benzoate produced from toluene metabolism through the upper pathway was not able to induce the ortho pathway in the chromosome (Kim et al. 2016). Among the hypothesis suggested to explain this striking finding, physical channeling of TOL pathway metabolites in the cytoplasm toward the TOL enzymatic machinery opens up new interesting perspectives to analyze the spatial organization (compartmentalization) of cellular metabolism (de Lorenzo et al. 2015).

3.2

A Family of Anti-activators Accompanying XylS/AraC Regulators

XylS belongs to the AraC/XylS family of transcriptional regulators. Several members of the AraC family involved in virulence gene regulation display an autoactivation mechanism (Martinez-Laguna et al. 1999; Porter et al. 2004; Morin et al. 2010) and, in contrast to XylS, AraC, or many other well-characterized members of the family, do not require effector binding for activation. Recently, a highly conserved family of small proteins named the AraC Negative Regulators (ANRs) has been shown to downregulate their AraC partners in pathogenic E. coli (Santiago et al. 2014, 2016). ANRs seem to play their regulatory roles by directly binding their cognate AraC partners, preventing DNA binding and activation of expression. Preliminary searches into the Pseudomonas genome and the TOL coding sequence did not render any putative candidates belonging to this new family of regulators. Nevertheless, we cannot discard the possibility that functional homologues of the ANR family members might exist and have not been identified to date.

4

Integration in the Cell Regulatory Networks: Toward Optimization of Expression

Table 1 lists the cell global regulators identified so far that are involved in TOL-mediated degradation of aromatic compounds. The abundance of host factors involved in TOL pathway regulation denotes a long coexistence of plasmid and host, which has led to the 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 σ54-dependent Pu and PS1 promoters and to a lesser extent Pm. Pu and PS1 respond to the availability of alternative carbon source, while Pm is optimized to maintain significant expression levels throughout the growth phase under induced conditions. The host factors affecting Pm expression have already been addressed above.

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Table 1 Protein factors involved in TOL pathway regulation Factor XylR

Gene xylR

XylS

xylS

Sigma-70

rpoD

Sigma-38

rpoS

Sigma-32

rpoH

Sigma-54

rpoN

IHF IIANtr Crc/Hfq

him, hip ptsN crc/hfq

General, nitrogen, other functions General General General

FtsH

ftsH

General

HU

hupA, hupB General

PprA TurA

pprA turA

CRP a

Category Pathway specific Pathway specific General, house keeping General, stress/stationary General, stress

General General, low temperature crp/cya (vfr) General

Target (role)a References Upper pathway (+), xylS (+) meta-cleavage pathway (+) xylR (+), xylS (+) meta-cleavage pathway (+)

Marqués et al. (1995) meta-cleavage pathway (+) Marqués et al. (1999) Upper pathway (+), xylS (+) Gallegos et al. (1996a), Marqués et al. (1998) Upper pathway (+), xylS (
) Holtel et al. (1995) Upper pathway, xylS Cases et al. (1999) Upper pathway (
), metaMoreno et al. (2010, cleavage pathway (
), xylR 2015) (
), xylS (
) Upper pathway (+) Carmona and de Lorenzo (1999), Sze et al. (2002) xylS (+) Pérez-Martín and de Lorenzo (1995b) Upper pathway (
) Vitale et al. (2008) Upper pathway (
) Rescalli et al. (2004) Upper pathway (
) in E. coli Zhang et al. (2014)

(+) positive effect; (
) negative effect

Influence of global physiological conditions on Pu expression is observed in rich medium as a delay in the time course induction of the pathway after effector addition, which has also been called “exponential silencing” (reviewed in Cases and de Lorenzo 2005). This delay is also observed in PS1 and reflects the catabolite inhibition exerted by LB components, since it can be reproduced by the addition of casamino acids and of certain mixtures of amino acids (Marqués et al. 1994). This inhibition is at least partially dependent on the Crc and Hfq proteins (see below). Catabolite repression is also observed in defined minimal medium as reduced activation levels when a carbon source such as glucose, gluconate, or 2-ketoglutarate is present in addition to the effector (reviewed in Ruíz et al. 2004). These effects are mimicked in continuous culture, in which growth under carbon excess leads to total repression of the pathway in the presence of effector regardless of the carbon source used and regardless of the limiting substrate selected, while growth in carbon-limited conditions allows substantial expression when the effector is present (Duetz et al. 1996). The mechanisms underlying this modulation have been analyzed for years,

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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, and recent findings suggest that regulation is more intricate than initially considered. In Pseudomonas, the classical CRP-dependent regulation present in Enterobacteriaceae appears not to be involved in catabolite repression, rather the Pseudomonas CRP orthologue Vfr regulates processes such as quorum sensing response and virulence (Suh et al. 2002; Rojo 2010; Coggan and Wolfgang 2012). Although in some P. putida strains vfr was required for the assimilation of certain amino acids, organic acids, urea, and ammonia as carbon and/or nitrogen source (Daniels et al. 2010; Herrera et al. 2012), the function of the Vfr/cAMP tandem in this species is still unclear, although clearly unrelated to metabolic functions (Milanesio et al. 2011). In this sense, inactivation of vfr in P. putida KT2440 was clearly shown to be irrelevant for TOL pathway expression regulation through Pu and PS1 promoters (Aranda-Olmedo et al. 2005). However, experiments in E. coli showed that Pu is sensitive to repression by CRP in a cAMP-dependent manner, probably by preventing direct contact between UAS-bound XylR with the promoter-bound RNA polymerase (Zhang et al. 2014). While it has been shown that P. putida Vfr maintains the capacity to bind DNA and cAMP and to contact RNAP, it seems to respond to different, so far unknown signals (Milanesio et al. 2011), which would explain the absence of effect of an vfr mutation on Pu and PS1 expression in its natural P. putida host. Three additional players in TOL expression have been considered as target of the global regulation. The first one was based on the regulatory network prevailing in Pseudomonas CF600 for the homologous system DmpR/Po, which involves the alarmone (p)ppGpp as main player (Sze and Shingler 1999). However, it seems this mechanism has only a marginal influence on XylR-mediated Pu expression (Carmona et al. 2000; Sze et al. 2002). The sigma factor σ54 was also explored as possible target for global regulation. In this sense, the detrimental effect of a mutation in the FtsH membrane protease gene on Pu activity suggested that the functionality of σ54, which is controlled by this protease, would influence Pu activity (Carmona and de Lorenzo 1999). However, comparative and combinatorial studies with the XylR/Pu system and the related DmpR/Po tandem regulating phenol degradation in Pseudomonas sp. strain CF600 showed that the actual target of FtsH in Pu expression was the XylR regulator. FtsH, which belongs to the AAA+ family of chaperone-like ATPases, would be required for XylR correct folding or multimerization to form the regulator-Eσ54 complex (Sze et al. 2002). The third player is the PtsN protein. In most Gram-negative bacteria where the rpoN gene coding for σ54 is found, it appears clustered with three other genes: ptsN, which codes for the phosphoenolpyruvate: sugar phosphotransferase system (PTS) component EIIANtr (Deutscher et al. 2006), ptsP coding for EINtr, and ptsO coding for NPr. The phosphorelay cascade of this PTS system flows from phosphoenolpyruvate (PEP) through EINtr and NPr to EIIANtr. Inactivation of the ptsN gene leads to a partial release of glucose or succinatedependent Pu repression (Cases et al. 1999; Aranda-Olmedo et al. 2006). PtsN repressing activity on Pu depends on its phosphorylation level, where the non-phosphorylated form of the protein is the one exerting the repressive effect.

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In Pseudomonas, the PTS system is suggested to control carbon/nitrogen balance in the cell (Cases et al. 2007). Analysis of mutants in the carbon metabolism of P. putida showed that routing of the carbon sources through the Entner-Doudoroff pathway was required for carbon catabolite repression (Velázquez et al. 2004). Furthermore, interrupting this pathway by knocking out the eda gene for 2-dehydro-3-deoxyphosphogluconate aldolase increased the repressive effect of glucose, suggesting that the glucose metabolite 2-keto-3-deoxy-6-phosphogluconate (KDPG) was the cellular signal triggering PTS phosphorelay for Pu regulation, probably by controlling PtsN phosphorylation state (Velázquez et al. 2004; del Castillo and Ramos 2007). KDPG was also shown to be the signal triggering glucose-dependent carbon catabolite repression of phenylacetic acid metabolism in this strain (Kim et al. 2009). In addition, in P. putida the PTS system is interconnected with the transport system for fructose, a non-repressive carbon source (Pflüger and de Lorenzo 2008), suggesting that carbon flux through the Entner-Doudoroff pathway would be translated to the C/N PTS system through FruB (EIIAFru). The analysis of a series of P. putida metabolic mutants 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 (Ruíz et al. 2004; Cases and de Lorenzo 2005). Interestingly, toluenemediated repression of glucose catabolism is mediated by the Crc protein, a global regulator that works together with the Hfq posttranscriptional regulator and controls carbon flow in Pseudomonas, playing a key role in catabolite repression (Collier et al. 1996; Rojo 2010). Although Crc was initially believed to directly bind RNA at a specific consensus sequence (termed the catabolite active motif), recent findings have shown that the actual role of Crc is to enhance and stabilize the complex formed between RNA and the RNA-binding protein Hfq. Hfq recognizes specific sites in the translation initiation region of target mRNAs, repressing translation (Sonnleitner and Blasi 2014). The effect of Crc stabilization of Hfq-RNA complexes at many target sites favors translational inhibition and explains the observed catabolite repression relief of crc mutants (Moreno et al. 2007, 2015; Madhusani et al. 2015). In P. putida, Crc, probably in cooperation with Hfq, has been shown to inhibit the assimilation of glucose and fructose in rich medium and to modulate the uptake of amino acids as well as their assimilation pathways, so that the use of the strain’s preferred amino acids is favored over the non-preferred ones (reviewed in Rojo 2010). Although a minor role of Crc in Pu repression in P. putida growing on rich medium had been described (Aranda-Olmedo et al. 2005), it was only after a thorough analysis of TOL pathway gene expression in a crc mutant that the role of this interesting protein in TOL pathway expression was elucidated (Moreno et al. 2010). In TOL, xylR and xylS mRNAs are targets for Crc/Hfq binding and repression of translation, which reduces the availability of these regulators in the cell, therefore tuning down overall expression of the pathway genes. Moreover, target sequences for Crc/Hfq were found overlapping the translation start site of several enzyme genes located in the upper and meta-cleavage polycistronic mRNAs, providing a mean to maintain an appropriate balance between the different pathway enzymes coded for in each messenger (Moreno et al. 2010).

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FtsH TurA IHF PprA

σ38

σ54 PtsN

σ32

Pu

upper

Pm

Crc/Hfq

HU σ70 σ54 PtsN IHF σ70

meta-cleavage

Crc/Hfq

xylS PS2 PS1

Crc/Hfq

PR1

PR2

xylR

Crc/Hfq

Fig. 3 Current knowledge on global regulatory proteins involved in the control of TOL system at either the transcriptional or the translational level. Green arrows indicate positive effect over transcription. Red blunt-headed lines indicate negative effect over transcription, except for Crc and Hfq that exert their control posttranscriptionally, inhibiting mRNA translation in response certain carbon sources. The mRNA transcripts are depicted as a gray curved line

Finally, by assaying P. putida crude extracts, two proteins with the capacity to bind Pu promoter were identified. The LytTR family two-component response regulator PprA, with no known function in P. putida, is able to bind Pu promoter at a site overlapping the UASs for XylR binding, thus competing with this regulator (Vitale et al. 2008). As expected, in vivo the presence of PprA had a repressive effect on Pu expression. The signals sensed by PprA to modulate Pu expression are unknown so far. TurA, a small protein with structural homology to the nucleoid protein H-NS, was also shown to bind Pu and repress expression from this promoter, an effect that was strengthened at low temperature (Rescalli et al. 2004). This repression is unrelated to the previously described exponential silencing or catabolite repression control of Pu promoter. The targets of the global regulators of xyl pathway expression discussed above are summarized in Fig. 3.

5

Research Needs

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 understand transcription regulation in bacteria. This will include the basic mechanisms leading to σ54 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 posttranscriptional regulation mechanisms (Velázquez et al. 2005). Understanding the processes underlying global control will definitely be the subject of thorough analysis in the future, especially to determine the role of the C/N PTS system, the final target of this phosphorelay and the possible

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additional elements involved. The behavior and performance of the system under natural conditions such as those found in real polluted environments will probably rely on the multiple signals that the bacteria encounter in these habitats, which are expected to be essentially different to those acting under the laboratory conditions analyzed so far. In this sense, our current knowledge of TOL pathway functioning is consistent enough to represent a perfect model to analyze the response of this biotechnologically relevant strain under real environment biodegradation conditions. Attempts in this sense are already underway (Svenningsen et al. 2016). A general issue barely addressed to date is how highly hydrophobic signal compounds such as hydrocarbons are spread in aqueous systems, how they approach and enter the cell, and how they find the regulator in the cytoplasm. In this sense, the physical channeling of TOL metabolites in the cytoplasm has been recently suggested (Kim et al. 2016) and deserves further analysis. Also, the role of biofilms and secreted proteins in the capacity of hydrocarbonoclastic bacteria to degrade hydrocarbons has been highlighted recently (Ennouri et al. 2016; Espinosa-Urgel and Marqués 2016), opening up new perspectives in the analysis of aromatic hydrocarbon degradation. As evidenced throughout this review, the regulatory network of the TOL plasmid aromatic degradation pathway has become a paradigm of transcriptional regulation in bacteria and a model of integration of a horizontally acquired pathway in the host metabolism. The TOL regulatory network as a whole, as well as independent parts, has been extensively exploited with biotechnological purposes, in particular in the design of efficient biosensors and expression systems (Marqués et al. 2006; de las Heras and de Lorenzo 2011; de las Heras et al. 2012, 2015; Balzer et al. 2013). Furthermore, it has proven to be an excellent tool to develop logic models to explain the network layout and dynamics and to predict the circuit’s behavior upon different metabolic challenges using system biology approaches (Silva-Rocha et al. 2011, 2013). Acknowledgments This work was supported by the European Regional Development Fund FEDER and grant from the Spanish Ministry of Economy and Competitiveness (BIO2014-54361-R).

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Genetics and Biochemistry of Biphenyl and PCB Biodegradation

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Loreine Agulló, Dietmar H. Pieper, and Michael Seeger

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Aerobic Metabolism of PCBs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Upper Pathway Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Archetype bph Gene Clusters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Genome Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Toxicity of PCBs and Their Metabolites and Bacterial Stress Response . . . . . . . . . . . . 2.5 Metabolic Versatility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Optimized Enzymes and PCB-Degrading Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Bioremediation of PCBs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Microorganisms are crucial for the removal of polychlorinated biphenyls (PCBs) from polluted environments. Microbial anaerobic dehalogenation of highly and moderately chlorinated biphenyls generates the subsequent less chlorinated congeners. Microbial aerobic degradation performed by enzymes of the biphenyl (bph) upper and lower pathways oxidizes moderately and low chlorinated biphenyls. These enzymes and their substrate specificities are discussed in Sect. 2.1. Biphenyl 2,3-dioxgenases (BDOs) are key enzymes of biphenyl pathways, which determine substrate range and extent of PCB degradation. In addition, the L. Agulló · M. Seeger (*) Laboratorio de Microbiología Molecular y Biotecnología Ambiental, Department of Chemistry and Center for Nanotechnology and Systems Biology and Centro de Biotecnología, Universidad Técnica Federico Santa María, Valparaíso, Chile e-mail: [email protected] D. H. Pieper Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_30

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specificity of subsequent enzymes is also crucial for productive metabolism. Specific native and engineered BDOs possess a wide range of substrates, which permit their application for synthesis of fine organic chemicals including novel bioactive compounds. The metabolism of PCBs is described in detail for some model organisms, and the genetic organization of gene clusters of model organisms is described in Sect. 2.2. The sequenced genomes of some PCB-metabolizing organisms including the model strains Burkholderia xenovorans LB400 and Rhodococcus jostii RHA1 improve the understanding of their overall metabolism, physiology, and evolution as described in Sect. 2.3. This has also allowed a better evaluation into genome and proteome-wide defenses against PCB toxicity, which is summarized in Sect. 2.4. However, our knowledge on enzymes and genes involved in PCB metabolism is still rather fragmentary and an overview of the diversity of enzymes reported and mosaic routes is given in Sect. 2.5. Finally, strategies to optimize microorganisms for improved PCB degradation and bioremediation processes are discussed in Sects. 2.6 and 2.7.

1

Introduction

Preserving the environment for future generations is a main aim for sustainable development. 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. PCBs, which are widely distributed in the environment, mainly in aquatic and soil ecosystems (Gomez-Gutiérrez et al. 2007; Palma-Fleming et al. 2008) were classified in the list of the 12 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 one to 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 the USA by Monsanto. These compounds were not only 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, USA, Canada, and UK), 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).

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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 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). In 2013 the International Agency for Research on Cancer (IARC) classified PCBs as human carcinogens (LaubySecretan et al. 2013). A study of a cohort of 24,865 capacitor-manufacturing workers exposed to polychlorinated biphenyls (PCBs) has associated PCB exposure with increased stomach, uterine, and prostate cancer and myeloma mortality (Ruder et al. 2014). Diverse full-scale applications of remediation technologies have been used for bioremediation of PCBs (Gomes et al. 2013). Technologies for the treatment of contaminated sites such as incineration, landfilling, thermal treatment (desorption, destruction, and vitrification), capping, and chemical dehalogenation are generally costly and usually involve dredging or excavation followed by disposal (Gomes et al. 2013; Fuentes et al. 2014). Microorganisms play a main role in the carbon cycle and in the removal of persistent organic pollutants from the environment. In situ and ex situ bioremediation has been applied successfully for the removal of petroleum contamination (Fuentes et al. 2014, 2015, 2016). For cleanup 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; Wu et al. 2002; Fennell et al. 2004). Dehalococcoides sp. strain CBDB1 is efficient for dechlorination of a wide range of PCB congeners in the environment (Adrian et al. 2009; Sowers and May 2013). Even though various reductive dehalogenases for dehalogenation of tetrachloroethene, vinyl chloride, or chlorobenzene (Neumann et al. 1996; Müller et al. 2004; Adrian et al. 2007) have been described, enzymes involved in reductive dehalogenation of PCBs remain to be identified. Lower and some moderately chlorinated PCBs are susceptible to aerobic metabolism.

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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, Arthrobacter, Acinetobacter, Corynebacterium, 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). Diverse in situ assays have shown the biotransformation of a broad range of PCB congeners in the environmental (Xu et al. 2010; Tu et al. 2011; Pentyala et al. 2011; Gomes et al. 2013). Novel PCB degraders have been reported. Sinorhizobium meliloti biotransforms a broad range of PCB congeners (Tu et al. 2011).

2.1

Upper Pathway Enzymes

The dioxygenation of the aromatic ring constitutes the first step in the aerobic bacterial catabolism of aromatic compounds. This reaction destroys the aromatic system and functionalizes the molecule for further degradation (Furukawa, 2000; Overwin et al. 2015b). 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).

2.1.1 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). Biphenyl 2,3-dioxygenases (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 α-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

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Fig. 1 Pathway for biphenyl degradation

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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 PCBs (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), ortho-brominated, and ortho-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 and flavonoids by BphA of strain LB400 has also been described (Seeger et al. 2003; Overwin et al. 2015a).

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-dihydro-2,3-dihydroxybiphenyls (biphenyl 2,3-dihydrodiol) to give (chlorinated) 2,3-dihydroxybiphenyl, is catalyzed by cis-2,3-dihydro-2,3dihydroxybiphenyl 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 (Rogers and Gibson 1977; Jouanneau and Meyer 2006). The cis-2,3-dihydro-2,3dihydroxybiphenyl 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). Interestingly, BphA from Burkholderia xenovorans LB400 catalyze dioxygenation of biphenyl 2,3-dihydrodiol (biphenyldienediol) in the nonoxidized ring to form biphenyl-bis-dienediol (Fig. 3) (Overwin et al. 2012). This metabolite is used as growth carbon source by B. xenovorans LB400 and Pseudomonas sp. strain B4-Magdeburg. BphB oxidizes both rings of the biphenyl-bis-dienediol in two successive steps into 2,3,20 ,30 -tetrahydroxybiphenyl (Overwin et al. 2012). In addition, BphA is able to oxidize 2,3-dihydroxybiphenyl produced in the biphenyl pathway to 2,3-dihydroxybiphenyl-4,6-diene-2,30 -diol, which is further transformed by BphB into 2,3,20 ,30 -tetrahydroxybiphenyl. 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. While intradiol dioxygenases, which cleave the aromatic nucleus between the hydroxyl 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

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Fig. 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 chlorosubstituent. From such an unstable vic-dihydrodiol, the chlorosubstituent is spontaneously eliminated. 2,5,20 -Trichlorobiphenyl is subject to both 20 ,30 dioxygenation and 3,4-dioxygenation (Haddock et al. 1995; Seeger et al. 1995a, 1999, 2001)

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Fig. 3 Conversion of biphenyl into 2,3,20 ,30 -tetrahydroxybiphenyl via two routes. Carbon numbering as used throughout this chapter indicated with compound 1. Compounds: 1, biphenyl; 2, biphenyldienediol [1-phenyl-cyclohexa-4,6-diene-cis-2,3-diol] (metabolite 1); 3a, 2,3-dihydroxybiphenyl; 3b, biphenyl-bisdienediol [1-(10 -cyclohexyl-40 ,60 -diene-20 ,30 -diol)cyclohexa-4,6-diene-2,3-diol] (metabolite 2); 4, 2,3-dihydroxybiphenyl-40 ,60 -diene-20 ,30 -diol [1-(10 -phenyl-20 ,30 -diol)-cyclohexa-4,6-diene-2,3-diol] (metabolite 3); 5, 2,3,20 ,30 -tetrahydroxybiphenyl (metabolite 4). The stereochemistry at carbons 20 and 30 of compounds 3b and 4 is proposed; it is based on the likely assumption that, for the second dioxygenation, the two rings simply exchange their positions within the BphA active site. Enzymes: BphA, biphenyl-2,3 dioxygenase (EC 1.14.12.18); BphB, cis-2,3-dihydrobiphenyl-2,3-diol dehydrogenase (EC 1.3.1.56) (Overwin et al. 2012)

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. 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 and 20 -chlorosubstituted 2,3-dihydroxybiphenyls strongly inhibit BphC enzymes

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(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-Dieneoate (HOPDA) Hydrolases The fourth step in the bph pathway is catalyzed by 2-hydroxy-6-phenyl-6-oxohexa2,4-dieneoate (HOPDA) hydrolase BphD, which hydrolyzes HOPDA to 2-hydroxypenta-2,4-dienoate and benzoate (Fig. 2). HOPDA hydrolases belong to the family of C–C hydrolase enzymes of the α/β-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 (Seeger et al. 1995b; Seah et al. 2000, 2001). 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. 4). 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). DxnB2 hydrolase from Sphingomonas wittichii RW1 catalyzes the hydrolysis of 3-Cl HOPDA more efficiently than BphD from B. xenovorans LB400 and Rhodococcus globerulus P6 (Seah et al. 2007; Ruzzini et al. 2013). Interestingly, DxnB2 is not inhibited by chlorinated HOPDAs, and the chlorine substituent is accommodated in the hydrophobic pocket (Ruzzini et al. 2013). 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); however, this enzyme can catalyze dehalogenation of 4-chlorobenzoate (Fig. 4), 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-2hydroxy-6-oxo-6-phenyl-2,4-dieneoates, compounds that are produced by the cometabolism of PCBs by BphA, BphB, and BphC (Fortin et al. 2006) and that inhibit BphD (Fig. 4), 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. A BphK modified at position 180 from an alanine to a proline showed an increased GST activity toward diverse chlorinated pesticides (McGuinness et al. 2007).

Fig. 4 Transformation of chlorosubstituted 2-hydroxy-6-oxo-6-phenyl-2,4-dieneoates (HOPDAs) by BphD and BphK gene products, exemplified by the metabolism of 4,40 -dichlorobiphenyl (Seah et al. 2000; Gilmartin et al. 2003; Fortin et al. 2006)

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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 acetylating acetaldehyde dehydrogenase (bphJ) to pyruvate and acetyl-CoA (Fig. 1), which then can enter the Krebs cycle. The aldolase type II BphI forms a tetrameric complex with the dehydrogenase BphJ, channeling the toxic aldehyde from BphI to BphJ (Baker et al. 2011; Carere et al. 2011). Molecular determinants for volatile aldehyde products channeling by the BphI-BphJ complex have been reported (Carere et al. 2011). BphH, BphI, and BphJ enzymes 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-2,4-dienoate can be transformed by BphH has yet to be elucidated. Besides 2-hydroxypenta-2,4-dienoates, benzoates are generated during BphDcatalyzed hydrolysis of HOPDAs (Figs. 1 and 4). 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-dihydroxydihydrobenzoyl-CoA and nonoxygenolytic cleavage of the aromatic ring. In contrast, chlorobenzoates, typically formed during metabolism of PCBs by the biphenyl upper pathway, are usually dead-end metabolites for PCB transforming bacteria.

2.2

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 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. 5). Regulation of these clusters is assumed to be mediated by an orf0 encoded GntR family transcriptional regulator (Watanabe et al. 2000). P. putida KF715 contains a bphABCD gene cluster (Hayase et al. 1990) (Fig. 5) 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). In Sphingobium fuliginis HC3, the bph genes are clustered in a plasmid (Hu et al. 2015). The presence of bph genes on mobile genetic elements indicate that

Fig. 5 Genetic organization of the bph gene clusters of B. xenovorans LB40, 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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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. 5). 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 genes are also located on a mobile genomic island (Toussaint et al. 2003). A third type bphAaAbAcAdCB gene cluster devoid of a gene encoding a HOPDA hydrolase was observed in R. jostii RHA1 (Masai et al. 1995) (Fig. 5) 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. Figure 5 illustrates a fourth type of bph cluster (bphBCA1A2A3A4D) found in Rhodococcus sp. K37 and Rhodococcus sp. strain R04 (Yang et al. 2007; Taguchi et al. 2007), which differs from the bph cluster from Rhodococcus jostii RHA1.

2.3

Genome Analyses

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 Mbp 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 Mbp, 0.44 Mbp, 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). 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

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for ring cleavage and for central routes channeling those intermediates into the Krebs cycle. Diverse active catabolic pathways for aromatic compounds from B. xenovorans LB400 have been reported (Méndez et al. 2011; Chirino et al., 2013; Romero-Silva et al. 2013; Agulló et al. 2017). 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). The genomes of two additional PCB-degrading bacteria have been sequenced. Pseudomonas pseudoalcaligenes KF707 genome possesses an important number of genes involved in catabolic pathways of biphenyl/PCBs, phenol, benzoate, and other chloroaromatic compounds (Triscari-Barberi et al. 2012). Rhodococcus sp. WB1 genome contains a biphenyl/PCBs bphBA3A2A1CDA4 gene cluster and additional catabolic pathways genes for the degradation of nicotinate, nicotinamide, polycyclic aromatic hydrocarbons, toluene, nitrotoluene, and atrazine (Xu et al. 2016).

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), and, in fact, PCBs decrease bacterial cell viability (Cámara 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 (Seeger et al. 1995a; Seah et al. 2000). Biotransformation of PCBs by BphA and BphB produces dihydrodiols and dihydroxybiphenyls, which are highly toxic for bacteria (Cámara 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 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 generates reactive products that can be cytotoxic (Fig. 6). 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

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Fig. 6 Biotransformation of PCBs into cytotoxic intermediates. Cytotoxic metabolites are boxed. Inhibition is indicated by a dashed arrow

reason for the poor survival of PCB cometabolizing organisms in soil microcosm studies (Blasco et al. 1997). Toxicity of PCBs as 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 straindependent differences were observed in cells metabolizing PCBs. Strain LB400 exhibited a high tolerance to PCB degradation-dependent toxicity, whereas RHA1 was highly sensitive. 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 (Agulló et al. 2007) and of DnaK and HtpG by 4-chlorobenzoate, a dead-end metabolite of the biphenyl upper pathway (Martínez et al. 2007), indicating that such exposure constitutes stressful conditions. The generation of reactive oxygen species (Chavez et al. 2004; Méndez 2017), probably resulting from the action of oxygenases in the metabolism, resulted in the induction of the alkyl hydroperoxide reductase AhpCF and other proteins indicating oxidative stress (Agulló et al. 2007; Méndez 2017). AhpCF detoxifies peroxides. The induction of a 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

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balance the activities of enzymes involved in the degradation to avoid accumulation of toxic metabolites. Approaches to overcome a metabolic dead-end step (Saavedra et al. 2010) and to increase tolerance to oxidative stress (Ponce et al. 2011) of PCB-degrading bacteria have been successfully applied to improve biodegradation and bioremediation of PCBs and will be presented in Sect. 2.7. The isolation of novel PCB-degrading bacteria that also degrade chlorobenzoates such as Sphingobium fuliginis HC3 (Hu et al. 2015) will also provide improved catalysts for PCB bioremediation.

2.5 2.5.1

Metabolic Versatility

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 (Mukerjee-Dhar et al. 2005) (Fig. 5) encoding enzymes only distantly related to enzymes of archetype Bph enzymes (Fig. 7), 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; Mukerjee-Dhar 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. 7). 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 α-subunit of R. jostii RHA1, only distantly related to previously characterized BphA1 proteins (see Fig. 7), 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 α-subunit has been discovered in Rhodococcus sp. strain K37 (Taguchi et al. 2007) (Fig. 7), 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

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Fig. 7 Dendrogram showing the relatedness of oxygenase α-subunits of Rieske non-heme iron oxygenases. α-Subunits of supposed or validated biphenyl 2,3-dioxygenases are indicated by a filled circle

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. 7) (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 remain to be elucidated (Taylor et al. 2002; Witzig et al. 2006), and 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

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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 (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 (Taguchi et al. 2004; McLeod et al. 2006). 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 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 (Cámara et al. 2007). Combination of both approaches, the broad natural diversity and methods of artificial evolution by family shuffling of soil DNA encoding BphA segments, generates 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 toward elucidation of pathway bottlenecks in downstream enzyme activities and in identifying optimized isoenzymes. As an example, a HOPDA hydrolase with novel specificities toward 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). On the other side, a modified BphK with increased GST activity toward a broad range of chlorinated

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organic substrates (McGuinness et al. 2007) may be useful to improve PCB mineralization. As described above, most currently available microorganisms are capable to mineralize biphenyl, but only cometabolize PCBs due to the absence of enzymes necessary for mineralization of chlorobenzoates, generated through metabolism by the biphenyl upper pathway. 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. 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). As mentioned above, the isolation of S. fuliginis strain HC3 that efficiently degrades PCBs without accumulation of dead-end intermediates was reported (Hu et al. 2015). Notably, the BphA-B4h hybrid enzyme based on biphenyl-dioxygenase from Pseudomonas sp. B4-Magdeburg and B. xenovorans LB400 is more efficient than BphA from strain LB400 for the dioxygenation of aromatic compounds (Overwin et al. 2015b). The BphA-B4h hybrid enzyme possesses a remarkable capacity for the double dioxygenation of various bicyclic aromatic compounds generating bis-DHDs (Fig. 3) (Overwin et al. 2016). The BphA-B4h hybrid dioxygenase is able to hydroxylate diverse flavonoids into novel products with interesting biological activities (Overwin et al. 2015a). The biotransformation of flavone, isoflavone, flavanone, and isoflavanol by the biphenyl dioxygenase from P. pseudoalcaligenes KF707 was reported (Seo et al. 2011).

2.7

Bioremediation of PCBs

Bioremediation of PCB-contaminated soils and sediments in microcosmos using aerobic and anaerobic bacteria has been reported (Singer et al. 2000; Bedard et al. 2007; Saavedra et al. 2010; Ponce et al. 2011; Sowers and May 2013; Payne et al. 2013). The genetically modified strain Cupriavidus pinatubonensis JMS34 containing the bph locus from strain LB400 is able to mineralize low chlorinated biphenyls in polluted soils (Saavedra et al. 2010). The addition of antioxidant compounds improved PCB degradation by B. xenovorans strain LB400 in soils (Ponce et al. 2011). Bioaugmentation of sediment microcosms and mesocosms indicates a high PCB degradation by aerobic B. xenovorans LB400 and anaerobic Dehalobium chlorocoercia DF1 (Payne et al. 2013). An increase of β-Proteobacteria and Actinobacteria in soils polluted with PCBs has been reported,

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which correlates with an increase of two bphA1 gene subgroups from β-Proteobacteria (Comamonas, Burkholderia) and Actinobacteria (Rhodococcus), indicating the selection of PCB-degrading bacteria (Correa et al. 2010). Phytoremediation and natural attenuation of PCBs has also been reported (Gomes et al. 2013). A transgenic Nicotiana tabacum plant that expresses the bphC gene from Pandoraea pnomenusa B-356 was applied for the removal of 2,3-dihydroxybiphenyl (Novakova et al. 2010). Bioaugmentation with B. xenovorans LB400 combined with phytoremediation using switchgrass increased the degradation of PCBs in soils (Liang et al. 2014). However, in situ PCB bioremediation approaches have to be further studied and developed.

3

Research Needs

The isolation of novel PCB-degrading bacteria with improved capabilities for the mineralization of PCBs is still a challenge in order to optimize bioremediation of PCBs. Genome studies of PCB degraders provide novel insights into the catabolic capabilities and the adaptation of bacteria to these pollutants. The sequencing of genomes of additional PCB-degrading strains will provide the basis for the understanding of the physiology and adaptation of bacteria dealing with PCBs and their toxic metabolic intermediates and related processes such as biofilm formation. Biofilm formation by bacteria is crucial toward their application for bioremediation. Different PCB-bioremediation processes have been reported in the last decades. Nevertheless, the field is still under development. A higher number of bioremediation trials in soils, sediments, and aquatic systems at diverse scale-up levels including microcosms, macrocosms, and field studies are still required to find out crucial aspects and variables to improve these complex processes. Bioaugmentation, biostimulation, and bioventing are three of the most relevant technologies for bioremediation that should be further studied. For bioaugmentation, bacterial consortia or a combination of bacteria and fungi are main catalysts that should be applied in novel cleanup processes. Dynamics of microbial communities during bioremediation of PCBs in anaerobic and aerobic conditions should be addressed for the knowledge of main bioremediation actors and the most favorable scenarios for the cleanup of PCB-polluted environments. The knowledge of PCB degradation will require integration of singlemicroorganism analyses with environmental studies to understand functioning of complex microbial communities during the cleanup processes and to design novel bioremediation strategies. Acknowledgments M.S. gratefully acknowledges support from the grants FONDECYT (1070507, 1020221, 1110992, 1151174, 7020221, 7070174, 7080148, 7090079, and 7100027), USM (130522, 130836, 130948, 131109, 131342, 131562), MILENIO P04/007-F (MIDEPLAN), and CONICYT-BMBF. D.P. gratefully acknowledges support from the grant EU GOCE 003998 (BIOTOOL) and BACSIN.

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An Update on the Genomic View of Mycobacterial High-Molecular-Weight Polycyclic Aromatic Hydrocarbon Degradation

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Ohgew Kweon, Seong-Jae Kim, and Carl E. Cerniglia

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Mycobacterial Metabolism of HMW PAHs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Metabolic Ambiguity of HMW PAHs as a Strong Selective Constraint . . . . . . . . . . . . 2.2 Metabolic Network-Centric View of Mycobacterial HMW PAH Biodegradation . . 3 Genomic View of HMW PAH Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 PAH-Degrading Genes in HMW PAH-Degrading Mycobacterial Genomes . . . . . . . . 3.2 PAH-Degrading Genes in M. vanbaalenii PYR-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Ring-Hydroxylating Oxygenases (RHOs) as Key Enzymes in the Degradation of HMW PAHs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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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 potentially toxic to living organisms. A phylogenetically diverse bacterial community has been identified and characterized from PAH-contaminated soils and sediments that have the ability to degrade these ubiquitous pollutants. Strains of mycobacteria have frequently been found to degrade HMW PAHs. There are significant differences between the genetic systems in mycobacteria and other bacterial strains that degrade monocyclic aromatic compounds, low-molecular-weight (LMW), and HMW PAHs. Recently, a series of interdisciplinary studies using systems biology and analytical O. Kweon · S.-J. Kim · C. E. Cerniglia (*) Division of Microbiology, National Center for Toxicological Research, Food and Drug Administration, Jefferson, AR, USA e-mail: [email protected]; [email protected]; [email protected] # This is a U.S. Government work and not under copyright protection in the US; foreign copyright protection may apply 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_31

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chemistry approaches has provided global insights into the structure, behavior, and evolution of mycobacterial HMW PAH degradation. In this review, we provide an update on genome-based systematic research with explanations and insights on the mycobacterial degradation of HMW PAHs.

1

Introduction

The fate of high-molecular-weight (HMW) PAHs with four or more fused aromatic rings in nature 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; Kweon et al. 2016). A number of environmental factors affect the microbial metabolism of HMW PAHs and lead to their persistence in soil. Inherent recalcitrance, due to their stable physicochemical structures, also hampers the microbial degradation of HMW PAHs (Cerniglia 1992; Ghosal et al. 2016; Haritash and Kaushik 2009; Kanaly and Harayama 2010; Kweon et al. 2016; Peng et al. 2008). Adaptation strategies and PAH bioavailability-promoting mechanisms exist to overcome these obstacles. These include low requirements of energy or oxygen 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. 2002), multiple substrate utilization (Wick et al. 2003), 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 PAH-directed motility (Fredslund et al. 2008). However, from a fundamental standpoint, HMW PAH degradation strategies involve the functions of versatile catabolic genes and enzymes as well as their efficient regulation (Kim et al. 2007, 2008, 2012, 2015; Kweon et al. 2007, 2010, 2011, 2014). Thorough reviews have been written on the environmental factors and biological strategies for the microbial degradation of PAHs (Brezna et al. 2006; Cerniglia 1992; Cerniglia and Sutherland 2006; Das and Mukherjee 2007; Ghosal et al. 2016; Haritash and Kaushik 2009; Johnsen et al. 2005; Juhasz and Naidu 2000; Kanaly and Harayama 2000, 2010; Kim et al. 2006; Kweon et al. 2016; Peng et al. 2008; 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; Diaz 2004; Habe and Omori 2003; Johnsen et al. 2005; Williams and Sayers 1994). However, studies on the enzymes and catabolic genes involved in HMW PAH degradation are few (Kanaly and Harayama 2000, 2010). Relatively few bacterial strains have been found to be HMW PAH-degraders, in comparison to a large number of bacteria capable of degrading LMW aromatic compounds. This is perhaps due to the lower bioavailability of HMW PAHs (Kanaly and Harayama 2000, 2010). Although several Gram-negative Pseudomonas and Sphingomonas strains have been reported to degrade HMW PAHs (Das and Mukherjee 2007; Gibson et al. 1975; Kanaly and Harayama 2000, 2010; Kazunga and Aitken 2000; Pinyakong et al. 2003), most organisms degrading HMW PAHs

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are nocardioform Gram-positive actinomycetes, especially members of the genera Mycobacterium (Boldrin et al. 1993; Dean-Ross and Cerniglia 1996; Grosser et al. 1991; Heitkamp et al. 1988; Kelley and Cerniglia 1995; López et al. 2005; Vila et al. 2001) 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, 2008; Krivobok et al. 2003; Kweon et al. 2010, 2014; 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 in 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 of mycobacteria or significant genetic divergence from those of Gram-negative bacteria. In recent years, the knowledge of bacterial HMW PAH metabolism has increased using advanced high-throughput molecular, analytical chemistry and next-generation sequencing techniques (Kim et al. 2012, 2015; Kweon et al. 2011). These studies have shown that the members of the genus Mycobacterium appear to be specialized for the degradation of HMW PAHs (Brezna et al. 2006; Cerniglia 2003; Heitkamp and Cerniglia 1988; Heitkamp et al. 1988; Kallimanis et al. 2011; Kanaly and Harayama 2010; Kim et al. 2003, 2006, 2007, 2008, 2009, 2012; Kweon et al. 2007, 2010, 2011, 2014, 2015, 2016; Miller et al. 2004; Moody et al. 2004). In this review, we provide a genome-based systematic explanation for the mycobacterial degradation of HMW PAHs.

2

Mycobacterial Metabolism of HMW PAHs

Since a large base of information currently exists on PAH metabolism in certain mycobacteria (Badejo et al. 2013; Kallimanis et al. 2011; Kanaly and Harayama 2010; Kelley and Cerniglia 1995), several generalizations are now possible about mycobacterial HMW PAH metabolism. They are: (i) from a genomic perspective, highly conserved genetic sources have similar gene cluster structures; (ii) from a biochemical perspective, there is a common functional responsibility for the key enzyme systems; and (iii) from a metabolic network perspective, there is a common metabolic logic with a strategy for PAH degradation. The generalizations also include the following: (i) apparent substrate preference for HMW PAHs with more than three benzene rings (e.g., pyrene and fluoranthene), (ii) the occurrence of multiple degradation routes or pathways, (iii) relatively high gene/function redundancy and functional complexity (i.e., pleiotropic and epistatic activities) of the ring-hydroxylating oxygenases (RHOs), and (iv) a conserved central aromatic degradation pathway (β-ketoadipate pathway). Recently, systematic approaches,

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which rely on a basic principle that explains the integration of different layers and types of information, including the genome, transcriptome, metabolome, and proteome of mycobacteria, have established these meaningful relationships. Such strong relationships and their systematic explanations not only further support the generalizations but also provide an opportunity to integrate the generalizations into a PAH-degrading phenotype, which has its own structural, behavioral, and evolutionary features.

2.1

Metabolic Ambiguity of HMW PAHs as a Strong Selective Constraint

In general, HMW PAHs are more toxic and difficult to degrade, persisting longer than LMW PAHs in the environment. From a metabolic standpoint, HMW PAHs are intriguing. As shown in Fig. 1, HMW PAHs serve not only as nutrients to be metabolized but also as potential toxicants. Some activation enzymes (e.g., cytochromes P450 [P450s] and quinone reductases) can convert PAHs or intermediates into mutagenic metabolites with increased reactivity, including diol epoxides, quinones, and radical PAH cations (Cerniglia 1992; Kweon et al. 2015). In addition, some metabolic intermediates of PAHs can trigger an accumulation of reactive oxygen species (ROS), which cause oxidative DNA damage and the formation of DNA double-strand breaks. Therefore, uncontrolled metabolic activities of mycobacteria for HMW PAHs can cause toxicological problems. There is ample evidence that metabolic ambiguity is an evolutionary constraint, which has governed the structure, behavior, and evolution of mycobacterial HMW PAH metabolism (Kim et al. 2012, 2015; Kweon et al. 2011, 2014, 2015). This could be a fundamental reason for the generalizations on mycobacterial HMW PAH metabolism, i.e., a common selective response to maximize nutritional benefits and minimize possible toxic side effects.

2.2

Metabolic Network-Centric View of Mycobacterial HMW PAH Biodegradation

Several bacterial aromatic hydrocarbon metabolic networks have been reconstructed, e.g., a global biodegradation network, regardless of the microbial hosts (Pazos et al. 2003); an HMW PAH metabolic network (PAH-MN) from M. vanbaalenii PYR-1 (Kweon et al. 2011); and a fluoranthene degradation network from a fluoranthene-degrading soil microbial community (Zhao et al. 2016). Studies of the characteristics of the metabolic networks have successfully provided unique network-centric insights into bacterial aromatic biodegration. Especially, the genome-scale PAH-MN from M. vanbaalenii PYR-1, structurally visualized from 10 aromatic hydrocarbon metabolic pathways with 183 nodes and 224 edges, has provided direct evidence that mycobacterial HMW PAH metabolism is neither simple nor random in terms of its structure, behavior, and evolution, which are essential to control the metabolic ambiguity of HMW PAHs.

CAP

RCP

HO

OH

Catechol Ring-cleavage dioxygenase

HO

Carbon & energy sources

Ring-cleavage products

O2–

trans-Dihydrodiol

HO H H OH

Dihydrodiol Epoxide

HO H H OH

O

O

–

O–

o-Semiquinone anion radical

H2O2

O2

O2–

O

Quinone

O

PAH-DNA adduct formation

Oxidative DNA damage

CYPs

Cytochrome P450s (CYPs) & Epoxide hydrolases

Dihydrodiol dehydrogenases

COOH

TCA Cycle

H OH

cis-Dihydrodiol

HO H

PAHs

An Update on the Genomic View of Mycobacterial High-Molecular-Weight. . .

Fig. 1 Metabolic ambiguity of PAHs as nutrients or toxicants and functional module-based general scheme of PAH metabolism in Mycobacterium vanbaalenii PYR-1. RCP ring cleavage process, SCP side chain process, CAP central aromatic process. Please refer to articles by Kweon et al. (2011), (2015), and (2016)

SCP

Cn

RCP

Ring-hydroxylating Oxygenases (RHOs)

HO

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Topological features of a metabolic network are central to an understanding of its behavior and evolution (Kim et al. 2015; Kweon et al. 2011, 2014, 2015). Structural analysis of the PAH-MN reveals not only its general topological characteristics, shared with other biological networks, but also unique structural features, specific to the aromatic biodegradation networks. The general topological features include scale-freeness and a few highly connected nodes (i.e., hubs). The degree of distribution of the PAH-MN follows a power law, P(k)
kγ, with γin or out = 2.8. Here the degree exponent (γ) ranging between 2.0 and 2.8 indicates that the PAH-MN is highly nonrandom (i.e., scale-free), and a few hub nodes dominate the overall connectivity in the network. On the other hand, a close structural examination of the PAH-MN has led to biodegradation-specific topological features showing that: (i) large and insoluble compounds tend to be far away from the central metabolism; (ii) there is an apparent directed repeating pattern in the chemical properties (e.g., molecular weight and hydrophobicity); (iii) several key compounds have larger in-degree than out-degree, resulting in its concentrating structure (e.g., a typical funnel shape). As revealed in the successful translation of topological features to behavior, the scale-free, funnel-like structure of the PAH-MN is intimately related to its functional module-based behavior and evolution. As shown in Fig. 1, in the PAH-MN, PAH substrates are degraded by functional interactions of a set of functional modules, termed ring-cleavage processes (RCPs), side chain processes (SCPs), and central aromatic processes (CAPs). The three functional modules-based metabolic behavior of the PAH-MN follow the common aromatic compound metabolic logic, i.e., the activation of the thermodynamically stable benzene rings, ring cleavage, side chain removal, and production of biological metabolic precursors. The repeated RCP-SCP processes transform diverse PAHs into the common metabolic intermediate, protocatechuate, which is then funneled into CAP. The CAP (i.e., the β-ketoadipate pathway) converts protocatechuate to the small aliphatic compounds, acetyl-CoA and succinyl-CoA, which can directly enter central metabolism. The funnel-like structure and module-based behavior explain its channel management to enhance the beneficial metabolic funnel effects, i.e., generating more productive biological precursors, including pyruvate and acetyl coenzyme A, and fewer toxic intermediates, such as o-PAH quinones.

3

Genomic View of HMW PAH Degradation

Bacterial genome sequencing is now a standard procedure with an acceptable costeffectiveness. According to the 2017 Integrated Microbial Genomes (IMG) statistics (https://img.jgi.doe.gov/cgi-bin/w/main.cgi), there are more than 50,000 sequenced bacterial genomes, including over 6,300 finished genomes, currently publicly available. Among them, around 190 mycobacterial genomes have been completely sequenced. Although many of the mycobacterial genomes are of medical interest, genomic information for about eight completely genome-sequenced environmental mycobacteria with biodegradation ability provides unparalleled opportunities for understanding the genetic and molecular bases of the degradation of aromatic

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compounds, including HMW PAHs. These include seven HMW PAH-degrading mycobacterial strains (M. vanbaalenii PYR-1, M. gilvum PYR-GCK, M. gilvum Spyr1, Mycobacterium sp. Epa45, Mycobacterium spp. KMS, JLS, MCS), and Mycobacterium sp. YC-RL4, which degrades diethylhexyl phthalate (DEHP). Systematic analyses of mycobacterial genomes have addressed several molecular genetic questions on the versatile bacterial PAH degradation, such as how many PAH-degrading genes do the degraders have and evolutionary questions on how, when, and where they have been obtained.

3.1

PAH-Degrading Genes in HMW PAH-Degrading Mycobacterial Genomes

In the mycobacterial phenotype network (MPN), which was recently reconstructed to represent genotype-phenotype relationships in the genus Mycobacterium (Kweon et al. 2015), the “PAH-degrading” phenotype shows the lowest connection degree and a biased connection toward the free-living phenotypic nodes, i.e., “fast-growing” and “nonpathogenic” nodes. This network feature of the PAH-degrading phenotype consists of phylogenetic and environmental isolation of PAH degraders (Kweon et al. 2015). In addition, the functional pan-genomic analysis (pan-genomic analysis and then functional genomic data-based filtration) of the mycobacterial strains of the “PAH-degrading” node, i.e., PAH degraders, revealed that about 130 PAH-degrading genes are in common and among them, ~110 genes are expressed as proteins when cells are treated with PAHs (Kweon et al. 2015). HMW PAH-degrading mycobacteria share similar phenotypic features, genetic repertories, and PAH metabolic spectra. The genome size of PAH-degrading mycobacteria, including M. vanbaalenii PYR-1, is between 5.7 and 6.5 Mb, with an average of 6 Mb, which is larger than that of strains of the pathogen M. tuberculosis, whose average genome size is ~4.4 Mb. In the HMW PAH-degrading mycobacterial genomes, the PAH-degrading genes are mostly clustered, forming one or more catabolic islands. The gene order is highly conserved within the catabolic islands, but the genomic position is not conserved between strains. For strains JLS, KMS, and MCS, the catabolic islands range from 75 to 78 kb in length. In strain PYR-GCK, they consist of two duplicated 80 kb regions (DeBruyn et al. 2012). On the other hand, in M. vanbaalenii, the catabolic island is approximately 150 kb due to inserted genes not present in the other strains, including DNA mobility, transport, and hypothetical genes (Kim et al. 2008). The catabolic islands on the plasmids of strain KMS are 47 and 32 kb, respectively, and that of strain MCS is 29 kb. All the catabolic islands contain RHO genes (Kim et al. 2008). The most basic evolutionary questions on mycobacterial HMW PAH-degradation are how, when, and where the catabolic PAH metabolic and genetic capabilities have been obtained. Genome-based evidence could help address fundamental questions at the single cell and genus levels (DeBruyn et al. 2012; Kim et al. 2008; Kweon et al. 2011, 2015). Interestingly, the catabolic regions, congested by PAH-degrading

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genes, show an apparently lower average %G + C, ranging from 62% to 63% G + C, compared to the whole genome average of 67% to 68% G + C (chromosomes) and 65% to 66% G + C (plasmids) (DeBruyn et al. 2012). These deviations in GC content indicate that the catabolic regions could be clusters of horizontally acquired genes, i.e., “genomic islands (GIs).” The standard deviations (σ%G + C), the degree to which recent horizontal gene transfer (HGT) shapes a genome, of 23 aromatic degrading bacterial genomes, including those of the HMW PAH-degraders, further support the HGT-centric gene gain (Kweon et al. 2011). HMW PAH-degrading mycobacterial genomes show values of σ%G + C ranging from 2.99 to 3.17, suggesting that the mycobacterial PAH-degraders may have acquired the gene cluster by an ancient HGT event from a common ancestor rather than by vertical descent. This evolution pathway likely conferred immediate selective advantages to the recipients (Kweon et al. 2011). In addition, as revealed by analysis of the ~150 kb region A of M. vanbaalenii, the PAH-degrading genes seem to have been recruited from separate HGT events from neighbors with close physiological, biochemical, and phylogenetic relationships. The GIs in catalytic region A from M. vanbaalenii occupy 57% (85 kb) of the 150 kb and show a high sequence similarity to the catabolic genes of nocardioform actinomycetes, such as Terrabacter and Nocardioides. Another interesting observation was a strong relationship between the evolutionary modules (the GIs) of the genome and the functional modules of the PAH-MN (Kweon et al. 2011). Based on such a strong GIs-functional modules connection, a probable evolutionary pathway with a fundamental building principle, i.e., a patchwork assembly of the phylogenetic modules with a backward functional direction from central to peripheral reactions, has been introduced (Kweon et al. 2011, 2015).

3.2

PAH-Degrading Genes in M. vanbaalenii PYR-1

A comprehensive picture of the overall molecular basis for the bacterial metabolism of HMW PAHs has originated from analysis of the complete genome sequence of M. vanbaalenii (Kim et al. 2007, 2008; Kweon et al. 2007, 2011, 2015). 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 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 M. vanbaalenii genome has a 150 kb major catabolic island at positions 494–643 kb (region A) with an additional 31 kb catabolic island at positions 4711–4741 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 β-ketoadipate pathways (Fig. 2) (Kim et al. 2008). Significant differences in 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

0478 0506 0507

0476

631

0505

0502 0503 0504

0501

0500

0474 0475

phdF phdG 0472

phdJ

0493 0494 0495 0496 0497 0498 0499

0492

0491

0490

phdI

phtAb 0465 phtB phtAc phtAd

phtAa

0489

nidB

nidA

0460

0461 0462

nidD

0485

0484

0480 0481 0482

nidB2

494420 (bp) 0479

0459

0458

0456

0453 0454

a

0455

An Update on the Genomic View of Mycobacterial High-Molecular-Weight. . .

24

0529 0530 0531 0532

0528

0526 0527

0525

0524

0523

0522

0521

0519 0520

0518

0517

0516

0515

0513

0512

0511

0510

0508 0509

520905

0558

0579

0557

0553

0578

0555

0551 0552

0550

0547 0548 0549

0546

0545

0544

0543

0542

0541

0540

0539

0538

0534 0535 0536 0537

0533

547480

0585

0581 0582 0583

0580

0575 0576

0574

0573

0572

0571

0567 0568 0569 0570

0564 0565

0563

0562

0559 0560 0561

574710

0598 0599 0600

0597

0593

0594 0595 0596

0592

0591

0590

0587 R0007 0588 0589

0586

601556

627495

642917

b S22 H OH

S23

RHO (NidAB2/PhtAcAd)

S1

COOH OH

Dihydrodiol dehydrogenase (0544) OH

S2

H

OH

H OH

S24

O-CH3

S7

COOH

OH

S3

OH

O-CH3

Decarboxylase (0543)

OH

S10

S25

COOH

S5

O-CH3 O-CH3

RHO (0546/0547/PhtAcAd)

OH OH COOH

S6

Dihydrodiol dehydrogenase (0544)

COOH

S12

β-Ketoadipyl CoA thiolase (4589)

Succinyl-CoA + Acetyl-CoA

HO

HOOC

COOH COOH

trans-2’-Carboxybenzalpyruvate hydratase- S17 aldolase (0469)

CHO

H

S21

COOH

COOH

S11

O

OH 1-Hydroxy-2-naphthoate Protocatechuate-3,4dioxygenase (0468) S16 dioxygenase (0560/0561) O COOH

Catechol O-methyl transferase (3280)

Decarboxylase (0543)

S15

β-Ketoadipate succinyl CoA transferase (0564/0565)

CSCoA COOH

COOH OH

S20 O

COOH HO

COOH

S4

OH

Phthalate-3,4-dihydrodiol dehydrogenase (PhtB)

CHO

S9

COOH COOH

+

OH

β-ketoadipate enollactone hydrolase (0563)

S19

O

Hydratase-aldolase S14 (0472)

Aldehyde dehydrogenase (0486)

O

COOH COOH

COOH

HO

OH

OH

Ring-cleavage dioxygenase (0470)

COOH

OH

S8

O HOOC

Phthalate-3,4-dioxygenase (0463/0463/ PhtAcAd)

Ring-cleavage dioxygenase (0470) S13 O

Dihydrodiol dehydrogenase (0544)

OH

Catechol O-methyl transferase (3280)

COOH

OH

OH H

HOOC

β-Carboxy-cis,cis-muconate cycloisomerase (0562) O

O

TCA Cycle

HOOC

2’-Carboxybenzaldehyde dehydrogenase (0522)

S18

γ-Carboxymuconolactone decarboxylase (0563)

Fig. 2 (a) Graphical map showing genetic organization of catabolic region A of Mycobacterium vanbaalenii PYR-1. Genes and open reading frames (ORFs) are indicated by arrows. Numbers above the arrows indicate the locus tag numbers and genes involved in the degradation of aromatic hydrocarbons are indicated by boldface numbers. Arrows in color represent genes whose products are involved in the pyrene degradation pathway; blue for RCP, orange for SCP, and green for CAP. (b) Complete pyrene degradation pathway in Mycobacterium vanbaalenii PYR-1. The numbers in parentheses indicate the ORF locus tag numbers. The letter “S” denotes enzymatic reaction steps (Table 1). Arrows in color represent functional modules in the PAH-MN; blue for RCP, orange for SCP, and green for CAP. Readers are referred to the original papers (Kim et al. 2007, 2008, 2012; Kweon et al. 2007, 2011, 2014, 2015).

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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 (Fig. 2 and Table 1) (Kim et al. 2008). For example, genes Mvan_0487/0488 (RHO), Mvan_0544 (cis-dihydrodiol dehydrogenase), and Mvan_0470 (ring-cleavage dioxygenase), encoding the first three steps of pyrene degradation (i.e., responsible for RCP), are located in three different clusters over 70 kb of region A (Fig. 2). 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 (Fig. 2a). Many identified genes are enriched with multiple paralogs, showing a remarkable range of diversity (Kim et al. 2008).

3.3

Ring-Hydroxylating Oxygenases (RHOs) as Key Enzymes in the Degradation of HMW PAHs

The metabolism of HMW PAHs proceeds by mono- and dioxygenation reactions in bacterial systems. Either one or two atoms of oxygen are incorporated into substrates, forming dihydrodiol compounds with trans and cis configurations, respectively (Heitkamp and Cerniglia 1988; Kanaly and Harayama 2010; Kelley et al. 1990). Two groups of enzymes, multicomponent Rieske-type nonheme monoiron ring-hydroxylating oxygenases (RHOs) and heme cytochrome P450 monooxygenases (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. A significantly high number of gene copies encoding RHOs (21 copies) and CYPs (50 copies) have been identified in the genome of M. vanbaalenii (Kim et al. 2008). 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, 2008, 2012, 2015; Kweon et al. 2007, 2011, 2014). Recently, a series of the forward genetics-based functional studies for several RHO genes in M. vanbaalenii PYR-1 provided direct evidence for their pleiotropic and epistatic functional responsibilities in the PAH-MN (Kim et al. 2012; Kweon et al. 2014). Among the 21 genes, due mainly to the apparent selective functional benefit of the type V electron transfer chain (ETC), the type V RHOs are main oxygenases responsible for dioxygenation in the PAH-MN; (i) NidAB and NidA3B3 for HMW PAH-centric hydroxylation, such as that of pyrene and fluoranthene; (ii) PdoAB for LMW PAH-centric hydroxylation, including phenanthrene and fluorene; and (iii) PhtAB for phthalate (Kim et al. 2008, 2012; Kweon et al. 2010, 2011, 2014). The RHO-centric functional maps, which provide numerical scores of the relative contribution of each RHO enzyme in the PAH-MN (Kim et al. 2012; Kweon et al. 2014), prove the type V RHO enzymes’ dynamic pleiotropic and epistatic functional interactions. Considering the

Gene/(Mvan ID)b nidA/0488

nidB/0483

0544 phdF/0470 0543

0546

0547

phdG/0472 nidD/0486 phdI/0468 phdJ/0469 0522 phtAa/0463 phtAb/0464 phtAc

phtAd/0467

0466 0561 0560 0562

Metabolic enzymatic reactiona S1

S1

S2/6/23 S3/7 S4/15

S5

S5

S8 S9 S10 S11 S12 S13 S13 S1/5/13/22

S1/5/13/22

S14 S16 S16 S17

PdoB2

Phenanthrene ring-hydroxylating oxygenase, β subunit

Phthalate 3,4-dihydrodiol dehydrogenase Protocatechuate 3,4-dioxygenase, α subunit Protocatechuate 3,4-dioxygenase, β subunit β-Carboxy-cis,cis-muconate cycloisomerase

Oxygenase reductase component

PhtB PcaG PcaH PcaB

PhtAd

PhdG PhdH PhdI PhdJ PhdK PhtA1 PhtA2 PhtAc

PdoA2

Phenanthrene ring-hydroxylating oxygenase, α subunit

Hydratase-aldolase Aldehyde dehydrogenase 1-Hydroxy-2-naphthoate dioxygenase Trans-20 -carboxybenzalpyruvate hydratase-aldolase 2-Carboxylbenzaldehyde dehydrogenase Phthalate 3,4-dioxygenase, α subunit Phthalate 3,4-dioxygenase, β subunit Oxygenase ferredoxin component

PhdE PhdF PhtC

Dihydrodiol dehydrogenase Ring-cleavage dioxygenase Decarboxylase

64 45 58 44

59

84 85 46 55 60 74 68 69

99

99

76 83 74

98

Pyrene ring-hydroxylating oxygenase, β subunit PdoB1

Matching protein Proteinc Similarityd PdoA1 98

Enzyme Pyrene ring-hydroxylating oxygenase, α subunit

An Update on the Genomic View of Mycobacterial High-Molecular-Weight. . . (continued)

Source organism Mycobacterium sp. 6PY1 Mycobacterium sp. 6PY1 Nocardioides sp. KP7 Nocardioides sp. KP7 Arthrobacter keyseri 12B Mycobacterium sp. 6PY1 Mycobacterium sp. 6PY1 Nocardioides sp. KP7 Nocardioides sp. KP7 Nocardioides sp. KP7 Nocardioides sp. KP7 Nocardioides sp. KP7 Terrabacter sp. DBF63 Terrabacter sp. DBF63 Arthrobacter keyseri 12B Arthrobacter keyseri 12B Terrabacter sp. DBF63 Streptomyces sp. 2065 Streptomyces sp. 2065 Terrabacter sp. DBF63

Table 1 Proteins identified in the whole-cell proteome of M. vanbaalenii PYR-1 grown in the presence of pyrene (Kim et al. 2007, 2008) 24 633

Ephx1

Cytochrome P450 monooxygenase

Cytochrome P450 monooxygenase

Cytochrome P450 monooxygenase

Cytochrome P450 monooxygenase

Cytochrome P450 monooxygenase

Epoxide hydrolase I

0600

3012

3029

3108

CYP51/ 5161 0521

SpiL

MonD

MonD

Ema7

MonD

36

66

33

45

45

66

45

64 60 36 97 98 72

Oryctolagus Cuniculus

Source organism Rhodococcus opacus 1CP P. putida PRS2000 P. putida PRS2000 Terrabacter sp. DBF63 Terrabacter sp. HH4 Terrabacter sp. HH4 M. tuberculosis CDC1551 S. cinnamonensis ATCC15413 Streptomyces sp. HIS-0435 S. cinnamonensis ATCC15413 S. cinnamonensis ATCC15413 Sorangium cellulsum Soce90 Solanum chacoense

Indicates pyrene metabolic steps in Fig. 2 b 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 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

CYP51

Quinone reductase

PQR/2039

PcaI PcaJ PcaF PdoA PdoB MT1743

0564 0565 4589 nidA3/0525 nidB3/0526 COMT/3280

S20 S20 S21 S22 S22 S24/25

Matching protein Proteinc Similarityd PcaL 40

Enzyme γ-Carboxymuconolactone decarboxylase/ β-ketoadipate enol-lactone hydrolase β-Ketoadipate succinyl CoA transferase, α subunit β-Ketoadipate succinyl CoA transferase, β subunit β-Ketoadipyl CoA thiolase Ring-hydroxylating oxygenase, α subunit Ring-hydroxylating oxygenase, β subunit Catechol O-methyltransferase

Gene/(Mvan ID)b 0563

Metabolic enzymatic reactiona S18/19

Table 1 (continued)

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apparent substrate interaction effects in mixed PAHs, the pleiotropic and epistatic functional complexity of catabolic enzymes should be important key factors for a deeper understanding of mycobacterial HMW PAH degradation. Consistent with their functional responsibility in the HMW PAH metabolism, the type V RHO systems also are a key target for the pleiotropic-dependent abandonment of the “PAH-degrading” phenotype in a possible evolutionary trajectory of the genus Mycobacterium (Kweon et al. 2015).

4

Research Needs

Studies during the last three decades have proved that mycobacterial HMW PAH biodegradation is a well-organized metabolic endeavor by which these microorganisms obtain the energy and carbon sources to live and reproduce. Isolation of HMW PAH-degrading mycobacterial strains from the environment and a subsequent series of systematic studies, of molecular genetics, biochemistry, genomics, metabolomics, functional genomics, and bioinformatics, allowed us to characterize the components, including PAH metabolic intermediates, genes, and enzymes, and to understand the structure, behavior, and evolution of HMW PAH degradation. Despite this detailed knowledge, we are still a long way from understanding the cellular dynamic context of HMW PAH in vitro, in vivo, in situ, and ex situ biodegradation. In particular, knowledge gaps between a single compound PAH degradation and a PAH complex mixture degradation, single cell versus a microbial community, differences between in vitro and in vivo, and in situ and ex situ seriously hinder efforts toward practical bioremediation applications to reduce PAH concentration to nontoxic levels set by national and international regulatory authorities. More systematic integrated interdisciplinary research, assisted by application of advanced high-throughput omics and meta-omics techniques, should be conducted to bridge the laboratory and the field studies in the biodegradation of HMW PAHs. Disclaimer The views presented in this report are not necessarily those of the FDA.

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Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Aromatic Catabolic Pathways in Pseudomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 The β-Ketoadipate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Aerobic Hybrid Pathways: The Phenylacetyl-CoA Pathway . . . . . . . . . . . . . . . . . . . . . . . . 2.3 The Homogentisate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 The Gentisate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 The Homoprotocatechuate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 The Nicotinate and Nicotine Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 The meta-Cleavage Pathways that Generate 2-Hydroxypentadienoate . . . . . . . . . . . . . . 2.8 The Hydroquinone and Hydroxyhydroquinol Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 The Protocatechuate and Gallate meta-Cleavage Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . 3 System-Level Response to Aromatic Compounds in Pseudomonas . . . . . . . . . . . . . . . . . . . . . . . 4 Biotechnological Applications of the Metabolism of Aromatic Compounds in Pseudomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

The increased use of the “omic” techniques, e.g., genomics, proteomics, metabolomics, and fluxomics, as well as the systems biology approaches for addressing biological complexity from a holistic perspective, has contributed significantly to accelerate and complete our understanding on different aspects of the physiology, ecology, biochemistry, and regulatory mechanisms underlying the catabolism of aromatic compounds in bacteria of the Pseudomonas genus. Toxic aromatic compounds simultaneously serve as potential nutrients to be J. Nogales · J. L. García · E. Díaz (*) Department of Environmental Biology, Centro de Investigaciones Biológicas, Consejo Superior de Investigaciones Científicas, Madrid, Spain e-mail: [email protected]; [email protected]; [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_32

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metabolized by bacteria but also as cellular stressors. When bacteria are exposed to these compounds they exhibit a multifactorial response that comprises three major intimately connected programs: (i) metabolic programs that involve not only the compound-specific pathways but also their integration within the global metabolism of the host cell; (ii) stress-response programs, e.g., changes in lipid metabolism, efflux pumps, or molecular chaperones, for adaptation to sub-optimal growth conditions; and (iii) a social program, including cell motility and chemotaxis, reorganization of the cell envelope, biofilm formation, and cell-to-cell interactions. As individual cells rarely metabolize a wide range of substrates, metabolic specialization within the bacterial population becomes a relevant trait in the assembly of efficient microbial biodegrader communities. Genome-scale metabolic models of several Pseudomonas strains have been performed. These models, when combined with the emergent synthetic biology approaches, can be used to explore the potential of Pseudomonas as cell factories in different biotechnological applications. Therefore, Pseudomonas becomes a paradigmatic bacterial genus both for increasing basic knowledge on the catabolism of aromatic compounds and for the bioremediation and/or biosensing of toxic pollutants and the valorization of aromatic compounds present in biowaste toward a sustainable knowledge-based bioeconomy with social and environmental rewards.

1

Introduction

The γ-Proteobacteria of the genus Pseudomonas are widespread colonizers of soil and water, plant leaves and roots, animals, and humans. Thus, P. putida, P. fluorescens, P. protegens, P. stutzeri, P. azelaica, P. knackmussi, P. pseudoalcaligenes, 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. The genomes of many Pseudomonas strains belonging to different species, e.g., P. putida (e.g., strains KT2440, GB1, F1, W619, CSV86, S16, DOT-T1E, SF1, DLL-E4, B6-2), P. entomophila (e.g., strain L48), P. stutzeri (e.g., strains A1501, KF716, CCUG29243), P. fluorescens (strains SBW25, Pf-01), P. protegens (strain Pf-5), P. aeruginosa (e.g., strains PAO1, PA7, PA14, PACS2, and 2192), P. knackmussi (strain B13), P. azelaica (strains HBP1, Aramco J), P. pseudoalcaligenes (strain KF707), 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 is reflected by the sizes of their genomes (usually >6 Mb), which contain large sets of genes involved in carbon source utilization and adaptation, and explains the ubiquity of this genus (dos Santos et al. 2004). 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

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toluene/xylene-degrading strain, that is mainly known by 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 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 promoter strains (Belda et al. 2016). 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 acids (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 some of the available complete chromosomes of Pseudomonas strains and/or those pathways that have been identified in previous genome research work. Our genomic survey revealed, at least, 14 different central pathways to which many different peripheral pathways converge (Fig. 1). Most these pathways have been only experimentally confirmed in some Pseudomonas strains mainly P. putida and P. aeruginosa strains. Among the 11 Pseudomonas species analyzed, P. putida appears to be the most versatile regarding the ability to degrade aromatic compounds since it contains 12 out of the 14 identified central pathways, which is in agreement with the wide range of niches that this species can colonize (Udaondo et al. 2016). Nevertheless, the ability to degrade aromatic compounds is a strain-specific feature and several pathways present in 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 β-ketoadipate pathway are the most widespread in Pseudomonas, probably reflecting that their substrates (aromatic amino acids, phenylpropanoids, and several

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Fig. 1 Aromatic catabolic pathways and their genetic determinants in different Pseudomonas genomes. The different peripheral pathways (color lines) that converge in the same central intermediate (color circle) are indicated with the same color code. The name of the genes coding for the peripheral and central pathways is also shown. Continuous and discontinuous circles indicate central intermediates that are subject of meta-cleavage and ortho-cleavage, respectively. The aerobic hybrid pathway paa is indicated in green. The central pathways shown in blue (cat and pca) and violet (tod, amn, and cmt) tones mean convergent routes at a nonaromatic compound, i.e., β-ketoadipate enol-lactone and 2-hydroxypentadienoate, respectively. The chlorocatechol, hydroxyhydroquinol, and hydroquinone intermediates converge with the standard ortho-cleavage pathway at the level of β-ketoadipate. R in alkylcatechols can be H (catechol), CH3 (3-methylcatechol), or CH2CH3 (isopropylcatechol). The abbreviation of the different Pseudomonas species is as follows: ae, P. aeruginosa; az, P. azelaica; en, P. entomophila; fl, P. fluorescens; kn, P. knackmussi; me, P. mendocina; po, P. protegens; ps, P. pseudoalcaligenes; 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

aromatic acids and phenols) are common carbon sources in the environment, and suggesting that their assembly was an old evolutionary event. Whereas meta-cleavage pathways are highly diverse, ortho-cleavage is restricted to the central β-ketoadipate pathway, the modified β-ketoadipate pathway for chlorocatechols and the hydroxyhydroquinol pathway (Fig. 1). 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 both via ortho and meta cleavage (Fig. 1).

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Usually, the aromatic catabolic genes are arranged in gene clusters that contain also dedicated regulatory and transport genes, and, in some cases, some efflux pumps- and methyl-accepting chemotaxis proteins encoding genes. The detection of the compound in the environment, the uptake of the compound inside the cell, and the inducible expression of the catabolic genes are important issues that need to be fine-tuned for an efficient degradation of the aromatic compound. Methylaccepting chemotaxis proteins (MCPs), such as PcaY, NbaY, McpT, and CtpL, are involved in the detection and chemotaxis toward aromatic compounds (Luu et al. 2015). In P. putida F1, PcaY is a component of the pca regulon and it is involved in the metabolism-independent chemotaxis toward many different aromatic compounds that funnel into the β-ketoadipate pathway. Expression of pcaY is regulated by the levels of the inducer β-ketoadipate, which depends on the efficient uptake of 4-hydroxybenzoate by the specific PcaK transporter, thus explaining the intimate physiological links between transport, chemotaxis, and metabolism (Luu et al. 2015). Most of the known or 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 (Parales et al. 2008, Eren et al. 2012; Luu et al. 2015). 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. The effector-specific regulation can be tightly fine-tuned by the action of certain modulators and is, in turn, under control of overimposed mechanisms that connect the metabolic and energetic status of the cell to the activity of the individual catabolic clusters, leading to complex regulatory networks (Díaz et al. 2013; Durante-Rodríguez et al. 2017). It should be noted, however, that the catabolic potential in Pseudomonas is much wider than that summarized in Fig. 1 since many strains whose genome has not been yet 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; Marín et al. 2012). Moreover, a significant fraction of the degradation genes within the Pseudomonas genus are located in catabolic plasmids (sometimes within mobile genetic elements such as transposons that are present within the plasmids), some of which are large conjugative plasmids that have been completely sequenced, e.g., pTOL plasmids (e.g., pWW0 and pWW53) for toluene/xylenes degradation; pDTG1, NAH7, and pND6-1 for naphthalene (also phenanthrene and anthracene) degradation; pADP-1 for atrazine degradation; pCAR plasmids for carbazole degradation; pTTS12 and pSYT for styrene degradation; pPhe+ and pVI150 for phenol, cresols, and dimethylphenol degradation (Miyakoshi et al. 2007; Köhler et al. 2013; Shintani et al. 2013; Segura et al. 2014). Although these catabolic plasmids play a major role in the distribution of the ability to degrade and utilize aromatic pollutants

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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. In some cases, self-transmissible plasmids are not endowed with catabolic genes but with genes involved in the tolerance/resistance to the toxic effect of aromatic compounds, and they are also of great biotechnological interest to generate solvent tolerant strains. For instance, the tolerance plasmid pGRT1 from P. putida DOT-T1E encodes a solvent efflux pump that is essential for the survival of the strain in the presence of high concentrations of toluene (Ramos et al. 2015; Molina-Santiago et al. 2017). Given the high frequency exchange of catabolic and tolerance plasmids among bacteria, a new approach called “genetic bioaugmentation” has been accomplished to treat polluted environments. This approach involves the introduction of bacteria harboring a relevant self-transmissible catabolic/tolerance plasmid to stimulate the horizontal gene transfer of the target genes into indigenous microorganisms with better fitness for survival in the corresponding ecosystem (Segura et al. 2014). Some considerations about the central and peripheral pathways encoded in the chromosome of different completely sequenced Pseudomonas strains will be presented below, however, a deeper biochemical and genetic analysis can be found in a series of review articles (Harayama and Timmis 1992; Spain 1995; Harwood and Parales 1996; Reineke 1998; Jiménez et al. 2004; Vilchez-Vargas et al. 2010; Díaz et al. 2013), in the Biocatalysis/Biodegradation UM-BBD (https:// www.msi.umn.edu/content/university-minnesota-biocatalysis-and-biodegradationdatabase), and AromaDeg (http://aromadeg.siona.helmholtz-hzi.de) databases (Duarte et al. 2014), as well as in other chapters of this book.

2.1

The b-Ketoadipate Pathway

The β-ketoadipate pathway is considered one of the most widely distributed set of genes for degradation of aromatic compounds in microbes. This central pathway is composed by two ortho-cleavage branches, one for degradation of protocatechuate (pca genes) and the other for degradation of catechol (cat genes). In Pseudomonas, these two branches converge at the intermediate β-ketoadipate enol-lactone, that is finally degraded to TCA cycle intermediates by the activities encoded by the genes pcaD, codes for the lactonase that generates β-ketoadipate, pcaIJ, code for the CoA-transferase that produces β-ketoadipyl-CoA, and pcaF, encodes the thiolase the generates succinate and acetyl-CoA (Harwood and Parales 1996). The genomic analyses reported here showed 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 Pseudomonas, the cat branch is missing in the three available genomes of P. syringae, which do not contain a typical catechol meta-cleavage route (Fig. 1). See Sects. 2.7 and 2.9 for protocatechuate and catechol meta-cleavage pathways in some Pseudomonas strains.

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2.1.1 The Protocatechuate Branch Many abundant aromatic compounds such as p-hydroxybenzoate, and phenylpropanoids ( p-coumarate, caffeate, cinnamate, ferulate, etc.) and some hydroaromatics, e.g. quinate, are degraded through the central intermediate protocatechuate. The protocatechuate pathway is widely distributed in bacteria (Jimenez et al. 2004), and we have identified the pca genes in most Pseudomonas species sequenced except in P. mendocina (Fig. 1). The ortho-cleavage of protocatechuate by the two-component protocatechuate 3,4-dioxygenase (PcaGH) generates a carboxy-cis,cis-muconate that becomes converted to β-ketoadipate enol-lactone by the action of the PcaC and PcaD enzymes. Whereas the pca genes are arranged in a single cluster in P. fluorescens, they are organized in different clusters in other Pseudomonas strains (Jiménez 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). Phenylpropanoids, many of which can be methoxylated (e.g., ferulate), are common carbon sources derived from highly abundant biological polymers like lignin and suberin (Young et al. 2005; Bugg et al. 2011). Ferulate (3-methoxy-4hydroxy-cinnamate) is initially converted through a CoA-dependent nonβ-oxidative pathway, which involves firstly a ferulate-CoA ligase (Fcs) and then a feruloyl-CoA hydratase/lyase (Ech) that generates acetyl-CoA and vanillin (3-methoxy-4-hydroxy-benzaldehyde). Vanillin is further converted into protocatechuate by the action of a Vdh dehydrogenase (encoded by at least four different paralogous genes in P. putida KT2440) and an O-demethylase (encoded by the vanAB gene cluster) (Priefert et al. 2001; Jiménez et al. 2004; Simon et al. 2014) (Fig. 1). In P. putida CSV86, a second gene cluster (verAB) encodes the O-demethylase that converts veratric acid (3,4-dimethoxy-benzaldehyde) and isovanillic acid (3-hydroxy-4-methoxy-benzaldehyde) into vanillic acid and protocatechuate, respectively (Fig. 1) (Mohan and Phale 2017). Nonmethoxylated phenylpropanoid acids, e.g., p-coumarate and caffeate, are also degraded through the CoA-dependent non-β-oxidative pathway (fcs-ech-vdh) to 4-hydroxybenzoate and protocatechuate, respectively (Fig. 1). Phenylpropanoid alcohols 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) (Fig. 1) (Priefert et al. 2001). Interestingly, in some strains two additional genes, aat (encoding a putative β-ketothiolase) and acd (encoding a putative acyl-CoA dehydrogenase) cluster with the fcs-ech genes, and they could be responsible of an alternative CoA-dependent β-oxidative pathway for phenylpropanoids degradation to acetyl-CoA and vanillyl-CoA (Priefert et al. 2001). In some strains, such as P. azelaica HBP1 and P. azelaica Aramco J, the phenylpropanoid degradation pathway is encoded within an ICE element (ICEhbp) (García et al. 2014; El-Said Mohamed et al. 2015).

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p-Hydroxybenzyl alcohol is channeled to p-hydroxybenzoate by the combined action of an alcohol dehydrogenase (Badh) and an aldehyde dehydrogenase (Bzdh). In P. putida CSV86, these enzymes were found to catalyze the conversion of other aromatic compounds such as 1- and 2-hydroxymethylnaphthalene to respective naphthoic acids (dead-end products) (Paliwal et al. 2014). 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; Pillai and Swarup 2002; Chen et al. 2014), but the corresponding genes have not been identified yet in the chromosome of 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, benzyl alcohol, tryptophan, aniline, salicylate, anthranilate, mandelate, including some 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; Jiménez et al. 2004). Some of these peripheral pathways were shown to be present in the Pseudomonas genomes under study (Fig. 1). Catechol can then be subject of ortho- or meta-cleavage depending on the particular pathway and strain under study. In this section, we review the catechol orthocleavage (β-ketoadipate pathway); see Sect. 2.7 for catechol meta-cleavage pathways. Benzoate is a key intermediate in the catabolism of several aromatic compounds, e.g., mandelate and benzyl alcohol, 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 β-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. Moreover, some Pseudomonas, such as P. fluorescens Pf-01 and P. azelaica, contain two unlinked ben-cat clusters, which might represent paralogs that function under different environmental conditions (ecoparalogs). 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. In P. putida CSV86 and GB1, the benzyl alcohol dehydrogenase (Badh) and benzaldehyde dehydrogenase (Bzdh) convert this aromatic alcohol to benzoate (Fig. 1) (Paliwal et al. 2014). Some other common peripheral pathways that funnel into catechol are those for the degradation of tryptophan, salicylate (2-hydroxybenzoate), and phenol (Fig. 1). Tryptophan is converted into anthranilate (2-aminobenzoate) through the kynurenine pathway encoded by the kynA (encodes a tryptophan 2,3-dioxygenase), kynB (encodes a kynurenine formamidase) and kynU (kynureninase) genes (Kurnasov

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et al. 2003; Knoten et al. 2011), which are present in the genomes of P. protegens strain Pf-5 and in 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 oxoquinoline (Miyakoshi et al. 2007), and it is a precursor for the Pseudomonas quinolone-dependent quorum sensing signal (PQS) that controls numerous cellular functions and virulence in P. aeruginosa (Chugani and Greenberg 2010; Drees and Fetzner 2015). 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. protegens Pf-5, the kyn and ant genes are also clustered in the chromosome. The ant and cat genes are linked in the genome of P. knackmussi B13 (Miyazaki et al. 2015). Phenol, a major environmental pollutant, becomes hydroxylated to catechol by the action of a phenol monooxygenase. Although a phc/dmp cluster that codes for a multicomponent diiron phenol hydroxylase (Santos and Sá-Correia 2007) is usually present in plasmids, e.g., pVI150 (Powlowski and Shingler 1994), dmp orthologs have been found in the genome of P. putida GB1 and P. knackmussi B13 (Fig. 1) (Miyazaki et al. 2015). Salicylate suffers oxidative decarboxylation to catechol by the decarboxylating flavoprotein monooxygenase NahG (SalA). A nahG ortholog is found in the genomes of some P. putida strains, e.g., DOT-T1E and GB1 (Fig. 1). Degradation of chloroaromatic pollutants, e.g., chlorobenzene, polychlorobiphenyls (PCBs), etc., leads to chlorocatechols as central intermediates. Chlorocatechols are usually channeled via a modified ortho cleavage pathway (clc genes) that shares the pcaIJ and pcaF genes with the standard β-ketoadipate pathway (Fig. 1) (Reineke 1998). The modified ortho-cleavage pathway involves a chlorocatechol 1,2-dioxygenase (ClcA), a chloromuconate cycloisomerase (ClcB), and a dienelactone hydrolase (ClcD) that release the chloride ion, and a maleylacetate reductase (ClcE) that generates β-ketoadipate. The clc genes have been identified in the ICEclc element present in the chromosome of P. knackmussii B13 (Miyazaki et al. 2015). The B13 strain generates chlorocatechols as a result of the degradation of 3- and 4-chlorobenzoate through the action of a multicomponent dioxygenase that shows similarity to the benzoate (BenABC) and toluate (XylXYZ) 1,2-dioxygenase (Miyazaki et al. 2015). The clc pathway has been extensively studied in P. reinekei MT1 (Cámara et al. 2009). Alternative pathways for degradation of chloroaromatics have been reported in different Pseudomonas strains. Thus, whereas 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); 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 currently available Pseudomonas genomes. Although most bacteria mineralize methylaromatics via meta-cleavage (see Sect. 2.7) and cloroaromatics via ortho-cleavage, there are some exceptions. P. reinekei MT1 is one of the few bacteria described so far that degrade

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4-methylsalicylate via ortho cleavage of 4-methylcatechol with the formation of 4-methylmuconolactone. Further degradation of 4-methylmuconolactone requires a devoted mnl gene cluster that funnels this intermediate into the TCA cycle (Marín et al. 2010). Some P. putida strains, e.g., GJ31, encode novel chlorocatechol 2,3-dioxygenases (CbzE) that are able to productively convert 3-chlorocatechol by proximal meta-cleavage with the release of chlorine and the production of 2-hydroxymuconate that is further degraded through the classical meta-cleavage pathway (see below) (Schmidt et al. 2013).

2.2

Aerobic Hybrid Pathways: The Phenylacetyl-CoA Pathway

Aerobic degradation of phenylacetate differs significantly from the classical strategies that involve the activation of aromatic compounds to dihydroxylated intermediates followed by dioxygenolytic ring-cleavage. The paa phenylacetate pathway is a hybrid pathway that combines properties from the aerobic and anaerobic catabolism of aromatic compounds. Thus, the molecule is firstly activated through the addition of CoA by a phenylacetate-CoA ligase (Fernández et al. 2014). Phenylacetyl-CoA is transformed by a multicomponent oxygenase into its ring-1,2-epoxide that evolves to the formation of an oxepin-CoA intermediate, which is further cleaved to form an aliphatic compound (dehydrosuberyl-CoA) that is channeled through a β-oxidation like mechanism to β-ketoadipyl-CoA, thus converging with the β-ketoadipate central pathway (Teufel et al. 2010; Teufel et al. 2012). β-Ketoadipyl-CoA is then thiolytically splitted into succinyl-CoA and acetyl-CoA by a dedicated Paa thiolase (Nogales et al. 2007). The paa genes have been identified in many Pseudomonas strains (Fig. 1). 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 (Arias et al. 2008). The fad genes encode the β-oxidation of n-phenylalkanoates to phenylacetyl-CoA (Olivera et al. 2001) (Fig. 1). Although 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 been yet reported in the Pseudomonas genus (Ismail and Gescher 2012).

2.3

The Homogentisate Pathway

The central intermediate homogentisate (2,5-dihydroxyphenylacetate) is formed in the degradation of the aromatic amino acids phenylalanine and tyrosine. The homogentisate central pathway is encoded by the hmg genes responsible for the extradiol-ring cleavage of homogentisate (hmgA), isomerization of

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maleylacetoacetate (hmgC), and hydrolysis of fumarylacetoacetate into fumarate and acetoacetate (hmgB) (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: (i) hydroxylation of phenylalanine to tyrosine by the pterin-dependent phenylalanine hydroxylase (phhAB), (ii) deamination of tyrosine to 4-hydroxyphenylpyruvate by the tyrosine aminotransferase (tyrB or phhC), and (iii) oxygenation and decarboxylation of 4-hydroxyphenylpyruvate into homogentisate by a 4-hydroxyphenylpyruvate dioxygenase (hpd) (Arias-Barrau et al. 2004; Jiménez 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, whereas the hpd gene is clustered with the hmg genes in P. syringae, P. stutzeri, and P. mendocina, 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 reported in the published genomes of the sequenced Pseudomonas strains. However, some mha orthologs can be found by sequence comparison analysis in the genomes of P. citronellolis and P. putida NBRC14164T strains.

2.4

The Gentisate Pathway

Gentisate (2,5-dihydroxybenzoate) 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 encoded by the gtd genes responsible for the extradiol-ring cleavage of gentisate (gtdA), isomerization of maleylpyruvate (gtdC) and hydrolysis of fumarylpyruvate to fumarate and pyruvate (gtdB) (Jiménez et al. 2004). The gtdABC orthologs have been found in the genomes of P. aeruginosa and P. entomophila (Fig. 1). As an alternative to the isomerization route, maleylpyruvate can also be degraded via a direct hydrolytic route catalyzed by a maleylpyruvate hydrolase (HbzF) to yield pyruvate and maleate, the latter converted to D-maleate by a maleate hydratase (HbzIJ) in P. alcaligenes NCIMB 9867 (Liu et al. 2015c). 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 (3,4-dihydroxyphenylacetate) is formed during the catabolism of a number of aromatic amines and hydroxylated

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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 conversion of 4-hydroxyphenylacetate into homoprotocatechuate, and hpaG1G2EDFXHI for degradation of homoprotocatechuate to TCA cycle intermediates (Jiménez et al. 2004). Similar hpa genes were identified in P. protegens Pf-5, P. entomophila, P. knackmussi, and P. putida GB1 (Fig. 1), although their arrangement is slightly different to that found in P. aeruginosa. Biogenic amines can be formed by hydrolysis of arylsulfate esters through the action of the AtsA arylsulfatase. Tyramine and dopamine are catabolized via a tyramine oxidase (TynAB) and a hydroxyphenylacetaldehyde dehydrogenase (TynC) that produce 4-hydroxyphenylacetate (from tyramine) and homoprotocatechuate (from dopamine). The tyn cluster has been studied in P. putida U (Arcos et al. 2010) and is conserved in P. putida GB-1 (Fig. 1) closely linked to the hpa cluster. 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 and Nicotine Pathways

Nicotinic acid is a N-heterocyclic aromatic compound that is widely distributed in nature as part of pyridine cofactors (e.g., NAD and NADP), alkaloids (e.g., nicotine and anabasine), and it is essential (vitamin B3) for many organisms. The complete nicotinate degradation pathway (nic genes) has been elucidated in P. putida KT2440, and it involves a nicotinate molibdohydroxylase (nicAB) and 6-hydroxynicotinate monooxygenase (nicC), a peculiar 2,5-dihydroxypyridine (DHP) extradiol dioxygenase (nicX) and N-formylmaleamate deformylase (nicD), and a set of reactions that convert maleamate into ammonia and fumarate (nicF and nicE) (Jiménez et al. 2008). In Pseudomonas, the nic genes are arranged in highly conserved clusters that are only present in P. putida strains (Fig. 1), what is in agreement with the fact that only P. putida strains were reported to use nicotinic acid as sole carbon source (Jiménez et al. 2008, 2011). Some Pseudomonas strains, e.g., Pseudomonas sp. strain S16, are able to degrade nicotine to DHP via the pyrrolidine pathway. This pathway begins with the oxidation of nicotine to N-methylmyosmine by the NicA1/NicA2 nicotine oxidoreductase, followed by spontaneous hydrolysis to form pseudooxynicotine that is then oxidized to 3-succinoyl-pyridine by the combined action of a Pnao amidase and a Sapd dehydrogenase. Succinoyl-pyridine is further hydroxylated to 6-hydroxy-3succinoylpyridine by a SpmABC molibdenum-dependent monooxygenase, and then converted to succinic semialdehyde and DHP by the action of a HspA/HspB monooxygenase (Tang et al. 2013; Hu et al. 2015). DHP is further converted to TCA cycle intermediates by a reaction scheme similar to that of nicotinic acid degradation

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in strain KT2440 (Tang et al. 2012). Most of the genes responsible of nicotine degradation are located in the largest genome island of strain S16, suggesting their acquistion by horizontal gene transfer (Tang et al. 2012). Similar genes responsible for nicotine degradation have been identified in the genome of P. plecoglossicida TND35 (Raman et al. 2015).

2.7

The meta-Cleavage Pathways that Generate 2-Hydroxypentadienoate

Although catechol is usually degraded by the β-ketoadipate pathway (see above), alternative meta-cleavage routes are also present in Pseudomonas strains that degrade aromatic hydrocarbons, e.g., biphenyl, naphthalene, and phenols. 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 monoaromatic hydrocarbon-degraders that use different biochemical strategies for the activation of the hydrocarbons to the corresponding catecholic intermediates. Thus, P. putida mt-2 contains a pWW0encoded 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 by the XylXYZ dioxygenase and XylL dihydrodiol dehydrogenase 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). A similar tod cluster is also present in the chromosome of the solvent resistant P. putida DOT-T1E strain (Udaondo et al. 2013). Both the xyl and tod pathways convert catechol and its alkylderivatives into pyruvate and acetyl-CoA via a meta-cleavage route that generates 2-hydroxypentadienoate (HPD) as 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 crossregulation between the specific XylS and BenR transcriptional regulators (Domínguez-Cuevas et al. 2006). Nevertheless, successive growth of strain mt-2 in benzoate favors the loss of plasmid pWW0 and, therefore, the selection of the ortho-cleavage over the meta-cleavage pathway. Interestingly, any cross-activation of the chromosomal ben genes by the plasmid-encoded XylS activator will lead to a metabolic conflict during the degradation of m-xylene by P. putida mt-2 because 3-methylbenzoate generated as an intermediate can be channeled through the ortho pathway and produce toxic dead-end metabolites (2-methyl-2-enelactone).

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However, it was shown that the natural expression ranges of XylS are insufficient to cause as significant induction of the ben-cat genes when cells face endogenous or exogenous 3-methylbenzoate (Pérez-Pantoja et al. 2015). Besides benzene, toluene, and ethylbenzene, P. putida F1 is able to degrade p-cymene ( p-isopropyltoluene). The oxidation of the methyl group of p-cymene to produce p-cumate and the ring dioxygenation of the latter to produce 2,3-dihydroxyp-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-pcumate 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 and a sep cluster (encodes a solvent efflux pump) thus generating an aromatics catabolic island for aromatic hydrocarbon degradation in P. putida F1 (Kasahara et al. 2012). In P. reinekei MT1, the dhb cluster for 2,3-dihydroxybenzoate degradation encodes a chimeric meta-cleavage pathway where the first two enzymes, DhbA and DhbB, are related to the 2,3-dihydroxy-p-cumate dioxygenase and the decarboxylase acting on the ringcleavage product, and the subsequent steps are catalyzed by enzymes related to those involved in catechol meta-cleavage degradation (Marín et al. 2012). It is worth noting that the Tod enzymes are able to attack a much higher range of aromatic hydrocarbons than those that can be used as carbon sources by P. putida F1, revealing that gene expression may be a limiting factor of the ability of strain F1 to degrade contaminants. Moreover, this observation explains why P. putida F1 can be evolved to grow in additional hydrocarbons (e.g., biphenyl, cumene, propylbenzene and butylbenzene) when the expression of the tod and cym genes is guaranteed due to spontaneous mutations in the cognate regulatory genes, todS and cymR, that now recognize these additional aromatic hydrocarbons as inducers, as in the case of TodS, or inactivate the repressor, as in the case of CymR, and thus allow the expression of the catabolic tod and cym genes in the mutant strains (Choi et al. 2003). A different adaptation strategy was shown to take place when P. putida F1 was evolved to use styrene as sole carbon source (George et al. 2011). In this case, a mutation in the TodA subunit of the toluene dioxygenase leads to a less active enzyme that, hence, generates less amount of 3-vinylcatechol, which is a toxic substrate that inhibits the catechol 2,3-dioxygenase activity, thus allowing productive meta-cleavage of this intermediate and finally styrene mineralization (George and Hay 2012). Degradation of polycyclic aromatic hydrocarbons, such as biphenyl and naphthalene, requires an initial dioxygenation and meta-cleavage step for the generation of a monoaromatic acid, e.g., benzoate or salicylate, and then a second step of dioxygenation (benzoate) or monooxygenation (salicylate) followed by a catechol meta-cleavage reaction that leads to the formation of HPD. The bph genes responsible of the degradation of biphenyl via benzoate and HPD (Fig. 1) are located in a transmissible ICEbph-sal element (encoding also the sal genes for salicylate degradation via meta-cleavage) in P. pseudoalcaligenes KF707

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and P. stutzeri KF716 (Fujihara et al. 2006; Triscari-Barberi et al. 2012; Hirose et al. 2015). Although some P. putida strains contain the bph genes in conjugative plasmids, e.g., P. putida KF715, other strains appear to harbor the bph genes into the chromosome, e.g., P. putida B6-2 (Tang et al. 2011; Suenaga et al. 2017). In P. azelaica HBP1 and Aramco J strains, 2-hydroxybiphenyl is oxidized to 2,3-dihydroxybiphenyl by the HpbA mononoxygenase, and then converted into benzoate and HPD by the sequential action of a HbpC extradiol dioxygenase and a HbpD hydrolase (Fig. 1) (Jaspers et al. 2001). The hbp genes are located in an ICEhbp element that also harbors a pob-pca cluster for 4-hydroxybenzoate degradation (two other pob-pca clusters are located in other regions of the chromosome), a salicylate degradation cluster (see below), and a phenylpropanoid degradation cluster (see above) (García et al. 2014; El-Said Mohamed et al. 2015). Naphthalene becomes oxidized to salicylate via an upper pathway (nah genes) that comprises, (i) hydroxylation to 1,2-dihydroxynaphthalene by naphthalene dioxygenase (NahAaAbAcAd) and the corresponding dihydrodiol dehydrogenase (NahB) enzymes; (ii) meta-cleavage by the 1,2-dihydroxynaphthalene dioxygenase (NahC) and subsequent formation of salicylaldehyde and pyruvate (NahDE); (iii) oxidation of salicylaldehyde to salicylate (NahF). A lower pathway (sal genes) is involved in the oxidative decarboxylation of salicylate to catechol by the decarboxylating flavoprotein monooxygenase NahG, and the further meta-cleavage of catechol to pyruvate and acetyl-CoA by the NahHIJKNLMO enzymes (Fig. 1). Although nah genes are usually present in catabolic plasmids (NAH plasmids), a chromosomal location of nah genes has been shown in some Pseudomonas strains such as P. putida CSV86 and P. stutzeri CCUG29243. In strain CSV86, the nah genes are present in a mobilizable ICEnah element that confers the ability to degrade naphthalene to heterologous hosts (Paliwal et al. 2014). Salicylate is also formed from the degradation of 2,20 -dihydroxybipenyl through the hbp pathway in P. azelaica. A salicylate hydroxylase and a catechol meta-cleavage pathway are also encoded by the salA and dmp genes, respectively, that are located next to the hbp genes within the ICEhbp element (García et al. 2014; El-Said Mohamed et al. 2015), and by the sal gene cluster present in the ICEbph-sal element of P. pseudoalcaligenes KF707 and P. stutzeri KF716 (Fujihara et al. 2006; Triscari-Barberi et al. 2012; Hirose et al. 2015). Although extradiol dioxygenases usually require the presence of two ortho- or para-positioned 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 AmnAB 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 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

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gene cluster has been identified in the genome of P. putida W619 and in the ICEclc element of P. knackmussi B13 (Fig. 1) (Gaillard et al. 2006; Miyazaki et al. 2015).

2.8

The Hydroquinone and Hydroxyhydroquinol Pathways

Hydroquinone (1,4-dihydroxybenzene) is a central intermediate formed during the catabolism of acetophenones, e.g., 4-hydroxyacetophenone, 4-ethylphenol, and 4-nitrophenol in different Pseudomonas strains (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 β-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 β-ketoadipate. 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, b). In P. putida SF1 and DLL-E4 strains, p-nitrophenol (PNP) is degraded through the pnp pathway that generates hydroquinone as central intermediate. PnpA is a single component PNP 4-monooxygenase that oxidizes PNP to p-benzoquinone, which is then reduced to hydroquinone by the PnpB reductase. The pnpCDEF cluster shares significant similarities to the hapCDEF cluster (see above) and is involved in the conversion of hydroquinone to β-ketoadipate (Shen et al. 2010; Tikariha et al. 2016). The PnpCD (HapCD) hydroquinone 1,2-dioxygenase constitutes the prototype of a new structural class of extradiol dioxygenases of the cupin family that exhibits a four-residue coordination environment (Liu et al. 2015b). The pnp pathway (PnpA and PnpB) is also involved in the conversion of 4-nitrocatechol to the central intermediate hydroxyhydroquinol (1,3,4trihydroxybenzene). Then, an intradiol hydroxyhydroquinol 1,2-dioxygenase (PnpG) cleaves the aromatic ring generating maleylacetate. The pnpG gene is also located within the pnp cluster in P. putida SF1 and DLL-E4 strains (Shen et al. 2010; Tikariha et al. 2016). Thus, the hydroxyhydroquinol pathway in Pseudomonas is an ortho-cleavage pathway. Hydroxyhydroquinol is an intermediate formed during the degradation of other aromatic compounds, such as 2,4,5-trichlorophenoxyacetic acid or resorcinylic compounds (Daubaras et al. 1996). In the genome of P. knackmussi B13, there are two adjacent genes encoding putative hydroxyhydroquinol 1,2-dioxygenase and maleylacetate reductase of a predicted hydroxyhydroquinol pathway (Fig. 1) (Miyazaki et al. 2015).

2.9

The Protocatechuate and Gallate meta-Cleavage Pathways

Although the catabolism of protocatechuate via the β-ketoadipate pathway is a widespread feature in Pseudomonas, protocatechuate can be also subject of

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meta-cleavage (oxalomesaconate pathway) in some strains such as P. straminea NGJ1 (Maruyama et al. 2004). This protocatechuate pathway has been well studied in Sphingomonas and Comamonas strains and it involves a 4,5-cleavage of protocatechuate by the two-component protocatechuate 4,5-dioxygenase (LigAB). The resulting product, 4-carboxy-2-hydroxymuconate-6-semialdehyde, is converted to 2-pyrone-4,6-dicarboxylate (PDC) by a LigC dehydrogenase, and the latter hydrolyzed to oxalomesaconate (OMA) by a PDC lactonase (Lig I). OMA is then hydrated by the LigJ hydratase and finally cleaved to pyruvate and oxaloacetate by the LigK aldolase (Masai et al. 2007). The lig genes are organized in a cluster. Although an ortholog gene cluster (pro genes) was described in P. straminea NGJ1 (Maruyama et al. 2004), no ortholog clusters have been identified in the Psedomonas genomes sequenced so far. In contrast, most P. putida genomes contain a gal cluster involved in the degradation of gallate (3,4,5-trihydroxybenzoate) and that encodes some enzymes similar to the Lig enzymes involved in meta-cleavage of protocatechuate (Nogales et al. 2011). Gallate is first uptaken by P. putida by an essential GalT transporter, and subject to meta-cleavage by a gallate dioxygenase (GalA) that generates OMA. GalA is a monocomponent extradiol dioxygenase specific for gallate (it does not recognize protocatechuate) whose two-domain architecture resembles the fusion of the large and small subunits of the LigAB protocatechuate 4,5-dioxygenase (Nogales et al. 2005). OMA is then hydrated to 4-carboxy-4hydroxy-2-oxoadipate by a GalB hydratase (that belongs to an enzyme family different to that of LigJ hydratase), and the latter is cleaved to pyruvate and oxaloacetate by the GalC aldolase (LigK ortholog) (Mazurkewich et al. 2016). It has been suggested that the gal cluster constitutes an evolutionary acquisition, via horizontal gene transfer, by P. putida to increase its metabolic proficiency as a saprophytic omnivore (Nogales et al. 2011).

3

System-Level Response to Aromatic Compounds in Pseudomonas

Microorganisms living in environments enriched and/or polluted by aromatic compounds face an interesting paradox. While many of these compounds, mostly phenol compounds and aromatic hydrocarbons, can be used as carbon and energy source, allowing biodegraders to colonize niches refractory to other microbes, they are also stressors for the bacteria above a certain threshold concentration since they are membrane-damaging and macromolecule-disrupting agents that eventually could lead to cell death (Sikkema et al. 1995; Ramos et al. 2015). Therefore, when bacteria are exposed to these compounds, they exhibit a multifactorial response by adjusting their gene expression programs to adapt to the new environmental scenario. This global response involves: (i) metabolic programs that involve not only the compound-specific pathways (see above) but also their integration within the global metabolism of the host cell; (ii) stress-response programs, e.g., changes in lipid metabolism, efflux pumps, or molecular chaperones, for adaptation to suboptimal growth conditions; and (iii) the social program, including cell motility and

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chemotaxis, the organization of the cell envelope, biofilm formation, and cell-to-cell interactions. All the three major programs are intimately connected and influence each other (Domínguez-Cuevas et al. 2006; Díaz et al. 2013). The recent “omic” approaches allow a system biology view more realistic about 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. Cells exposed to toxic aromatics primarily respond to them as a molecular insult rather than as a carbon source (Domínguez-Cuevas et al. 2006). A pioneer transcriptomic approach showed the short-term response of P. putida KT2440 (pWW0) cells grown in citrate 15 min after they were challenged with a sublethal concentration of an aromatic hydrocarbon such as toluene (Domínguez-Cuevas et al. 2006). Only 5% of the reprogramming response corresponds with genes involved directly in the catabolism of toluene. The main cellular response involves proteins that are directly or indirectly related to stress-response and morphological programs assisting the cell toward the toxic effects of toluene. Similar gene expression profiles were found in other Pseudomonas strains growing in the presence of toluene such as P. putida S12 (Volkers et al. 2015) and P. monteilii CCTCC M2013683 (Yang et al. 2016), highlighting the large set of cellular processes required for the optimal resistance and/or degradation of such compounds. From a spatial-temporal point of view, 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 (Ramos et al. 2015). Time-series transcriptomic analyses upon toluene exposition in P. putida S12 show that differentially expressed genes increased immediately after toluene addition, with a maximum at 5 min. As expected, genes immediately upregulated were involved in lipid and membrane metabolism, but also genes related with energy production and synthesis of chaperones to refold denatured proteins were upregulated (Volkers et al. 2015). The accumulation of aromatic compounds in the cytoplasmic membrane uncouples the proton gradient at some extent, while releasing lipids and proteins. Thus, differential expression of energy-management systems as well as the activation of the heat-shock response (mainly protein folding machinery) driven by RpoH regulon seems to be essential for initial cell survival after exposition to aromatic compounds (Domínguez-Cuevas et al. 2006, Volkers et al. 2009, Li et al. 2015). The initial damage to the cell membrane also leads to a reduction in electron transport chain activity and to an increase in the production of hydrogen peroxide and other reactive oxygen species (ROS), which in turn produce an oxidative stress response. 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 (Domínguez-Cuevas et al. 2006). P. putida exposure to a phenol shock is also triggering 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

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confer increased cell protection and enhanced efficiency of cell repair mechanisms against phenol-induced damage (Santos et al. 2004, 2007). Comparative genomics analysis have shown that many of the enzymes involved in the oxidative stress response are a common feature of the P. putida core genome, strongly linking the ability to counteract oxidative stress with the catabolic potential of this species toward aromatic compounds (Udaondo et al. 2016). A widely observed trait in P. putida is that proteins putatively involved in solute transport, including the outer membrane OprF porin, are downregulated upon cell exposure to aromatics, hence minimizing the access of the toxic compound to the cytoplasm (Santos et al. 2004, 2007). Interestingly, and in contrast to the observed general repression of cell barrier functions that decrease membrane permeability, toluene shock in P. putida leads to the induction of genes encoding RND (resistancenodulation-cell division) efflux pumps involved in the extrusion of toxic chemicals as a defense mechanism (Nikaido and Takatsuka 2009; Wijte et al. 2011; Ramos et al. 2015). An 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). While highly toluene tolerant strains, e.g., DOT-T1E and S12, encode and express three pumps (TtgABC, TtgDEF, TtgGHI), sensitive toluene strains, e.g., KT2440, only encode the TtgABC extrusion pump. Interestingly, the sensitive KT2440 strain activates a larger number of genes in a higher magnitude than the resistant DOT-T1E strain upon toluene exposure (Molina-Santiago et al. 2017). Similarly to the responses observed with highly toxic aromatic compounds, e.g., phenols and aromatic hydrocarbons, a number of proteins involved in detoxification, oxidative stress response, and protein folding mechanisms, such as AhpC, GrpE, or DnaK, are upregulated when P. putida KT2440 is cultivated in aromatic acids, benzoate, or 4-hydroxybenzoate, and the GroEL protein is upregulated 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). Upregulation of efflux pump systems, chaperones, and proteins for cyclopropane fatty acids and threhalose synthesis was identified in proteomic analysis of P. putida KT2440 growing on vanillin as only carbon source (Simon et al. 2014). The involvement of the heat-shock response, oxidative stress response, and efflux pumps in response to aromatic compounds appear to be a general strategy which is also found in other aerobic (Agulló et al. 2007) and anaerobic (Trautwein et al. 2008) biodegraders. After membrane injury due to the presence of toxic aromatic compounds, there is a transient slow-down in motility, chemotaxis, and cell division, all of which are processes that consume large amounts of ATP but do not provide essential traits for survival under these conditions, as a general strategy to save energy for more useful stress endurance programs (see above) (Domínguez-Cuevas et al. 2006, MolinaSantiago et al. 2017). Accordingly, a significant number of proteins that contain the conserved GGDEF (diguanylate cyclase) and/or EAL (c-di-GMP phosphodiesterase) motives become induced by the toluene-shock (Domínguez-Cuevas et al. 2006).

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These proteins are known to control the cellular levels of c-di-GMP, a cyclic dinucleotide alarmone that regulates, among other cellular functions, cell transition to a sessile state (Jenal et al. 2017), and whose participation in motility toward aromatic (and non-aromatic) compounds in Pseudomonas has been shown (Sarand et al. 2008). An interesting finding derived from the transcriptomic experiments with P. putida KT2440 (pWW0) confronted to 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 (Domínguez-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. Upon the first contact with aromatic compounds, the cell requires the activation of an intensive metabolic program. The observed upregulation of genes involved in amino acid and nucleotide biosynthesis could explain the synthesis of this new proteome found in cells exposed to toluene (Domínguez-Cuevas et al. 2006) or phenol (Santos et al. 2007). This hypothesis is supported by the induction of the Tuf-1 elongation factor for protein synthesis and chaperone activity, and an inorganic pyrophosphatase (Ppa) as a major provider of energy for the polypeptidemaking reactions (Domínguez-Cuevas et al. 2006). Similarly, enzymes involved in the synthesis of 6-phosphogluconate, e.g., glucose-6-phosphate-1-dehydrogenase (Zwf-1), necessary to replenish some of the pentose phosphate intermediates needed for de novo biosynthesis of nucleotides, are also upregulated. Interestingly, Zwf-1 plays a key role as source of NADPH and belongs to the SoxR regulon that controls the oxidative stress response. NADPH is, in fact, not only required for anabolic functions but also for the functioning of many antioxidant enzymes. In this way, it has been shown the upregulation of several enzymes that produce NADPH (e.g., glucose-6-phosphate-1-dehydrogenase (Zwf-1), 6-phosphogluconate dehydratase (Edd), glutamate dehydrogenase (GdhA)), when cells are cultivated using an aromatic compound as carbon source or when they are under oxidative environmental stress (Zhao et al. 2004, Domínguez-Cuevas et al. 2006, Singh et al. 2007). In addition to the network-wide rerouting of carbon skeletons toward NADP-dependent dehydrogenases, Pseudomonas strains resort to pyridine nucleotide transhydrogenases, e.g., soluble SthA and membrane-bound proton-pumping PntAB, to fine-tune the redox homeostasis under aromatic compounds biodegradation conditions. The thereby achieved robustness in the redox balance may explain why Pseudomonas species are frequent hosts of oxidative routes for aromatic pollutants (Nikel et al. 2016b).

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A higher energy demand in phenol-stressed cells to cope with the energetically expensive adaptation mechanisms is consistent with the upregulation of TCA cycle dehydrogenases that generate NADH (Icd, SucA) or reduced quinones (SdhA) (Santos et al. 2007). In addition, the upregulation of phosphoenolpyruvate carboxykinase (PckA) in response to phenol may indicate that the TCA cycle also acts rerouting carbon flux to anabolic pathways, e.g., via phosphoenolpyruvate, that eventually leads to an increase in peptidoglycan synthesis (Santos et al. 2007). 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 be also 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 reprogramming observed when the cells face only toluene (see above) 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 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 (downregulated) and by increasing membrane stabilization through higher levels of the OprH porin (Volkers et al. 2006). As expected, the studies of Pseudomonas strains cultured in the presence of different aromatic compounds as sole carbon source reveal the induction of the catabolic pathway involved in the mineralization of the corresponding aromatic compound (enzymes and transport proteins). Metabolic modules responsible of modulating 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, are also upregulated. As an example of this reprogramming, the levels of pyruvate dehydrogenase decreased significantly when P. putida KT2440 is cultured in benzoate as only carbon source. Since acetyl-CoA and succinate are the final products of benzoate degradation via the β-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. Overproduction of

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acetyl-CoA during phenylethylamine degradation upregulates 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, Yun et al. 2011). Further evidence of this intensive metabolic reprogramming comes from substrate shift experiments in chemostat under carbon limitation. Shifts in carbon sources (e.g., from benzoate to glucose and from fructose to benzoate) resulted in a remarkable rearrangement of the gene expression levels and metabolic fluxes involved in the initial uptake and metabolism of these carbon sources in P. putida (Sudarsan et al. 2014). For instance, during the shift from fructose to benzoate, the upregulation of benzoate-activated regulators (BenR, CatR, and PcaR) led to the subsequent expression of the β-ketoadipate pathway for benzoate degradation. Similarly, it was observed the induction of the glcB gene, which encodes the malate synthase, driving the activation of the glyoxylate shunt as an anaplerotic route fixing the acetyl moiety formed during benzoate degradation (Sudarsan et al. 2014). Transcriptomic analyses were only partially consistent with carbon flux estimations. Thus, while the upregulation of glyoxylate shunt was in agreement with the observation that nearly 85% of the flux through the branch point of isocitrate was diverted through the glyoxylate cycle in the presence of benzoate, the large decrease in the flux through the isocitrate dehydrogenase is not reflected in the transcriptomic data. Overall, it was found a strong flux rerouting through central carbon metabolism (TCA cycle) in the presence of benzoate with no concomitant differential gene expression of the cognate enzymes, which argue in favor of additional means for flux regulation, such as allosteric (metabolic) and posttranscriptional mechanisms (Sudarsan et al. 2014). These observations highlight the complex metabolism of aromatic compounds in Pseudomonas and the imperative need of system level approaches in order to fully understand this catabolism. An emerging and previously unrecognized biological function of aromatic degradation pathways is the production of secondary metabolites (exchange factors) some of which play an important role in cell-to-cell communication and microbial interactions. Thus, the catabolism of tryptophan through the kynurenine pathway in P. aeruginosa generates anthranilate, which is a pivotal branch-point metabolite that can be channeled to the synthesis of quorum sensing signals such as the Pseudomonas quinolone signal (PQS). Some quorum sensing signals can, in turn, control the expression of aromatic degradation pathways, such as in the case of the ant-cat genes in P. aeruginosa (Chugani and Greenberg 2010). Some aromatic ring-cleavage dioxygenases, such as the Hod enzyme involved in the quinaldine degradation pathway, have the ability to cleave PQS and, thus, they behave as quorum quenching enzymes (Fetzner 2015). Another quorum quenching mechanism is carried out by protoanemonin, an antibiotic generated by misrouting of some chloroaromatic compounds via the classical β-ketoadipate pathway in some Pseudomonas strains, opening an interesting new function for this molecule present in active concentrations in certain pseudomonads-dominated consortia (Bobadilla Fazzini et al. 2013). In addition to quorum-sensing signals, there is a large number of aromatic exchange factors, e.g., siderophores, indole, antibiotics, that act in cell-to-cell communication.

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Thus, during the catabolism of phenylacetate different bioactive compounds and communication signals, e.g., tropodithietic acid and related tropone-derived compounds with antibacterial activity, can be generated (Teufel et al. 2011; Díaz et al. 2013). An attractive and promising approach comes from the possibility to integrate physiological and omics data into accurate metabolic networks constructed from biochemical and genetic data, e.g., genome-scale metabolic reconstructions (GEMREs). Such reconstructions contain detailed information on the target organism including the reaction stoichiometry and the biochemical and physiological data available. A GEMRE can be further transformed into a mathematical model (genome-scale model, GEM) which not only shows the metabolic capabilities of a particular organism but it allows to predict phenotype from genotype, contextualize omics data, etc., hence constituting an ideal tool for studying complex metabolic processes (O’Brien et al. 2015; Nogales and Agudo 2016). In this sense, the Pseudomonas genus has been subject of an intensive metabolic modeling effort. Today, GEMs exist for many Pseudomonas species including: (i) P. putida KT2440, e.g., iJN746 (Nogales et al. 2008), iJP850 (Puchalka et al. 2008), PpuMBEL1071 (Sohn et al. 2010), iJP962 (Oberhardt et al. 2011), Belda’s model (Belda et al. 2016), PpuQY1140 (Yuan et al. 2017), and iJN1411 (Nogales et al. 2017); (ii) P. aeruginosa PAO1,e.g., iMO1056 (Oberhardt et al. 2008) and iPae1146 (Bartell et al. 2017); (iii) P. fluorescens SBW25 (Borgos et al. 2013); and (iv) P. stutzeri A1501 (Babaei et al. 2014, 2015). However, only models from P. putida have been used to study the metabolism of aromatic compounds at system level. For instance, by using PpuMBEL1071 the capabilities to degrade aromatic compounds were studied by estimating biomass formation on different aromatics as sole carbon and energy sources (Sohn et al. 2010). By including the reactions from TOL plasmid, iJN746 was used to study the metabolism of toluene in P. putida contextualizing experimental data with in silico analysis (Nogales et al. 2008). Thus, when the model was constrained with experimental toluene and oxygen uptake rates, important discrepancies, such as a much lower in silico growth rate than in vivo growth rate, were found. Further in silico analyses revealed that the three oxidative reactions involved in the conversion of toluene to 2-hydroxymuconate semialdehyde, i.e., toluene monooxygenase, benzoate 1,2-dioxygenase, and catechol 2,3-dioxygenase, consumed up to 40% of the available oxygen. These observations supported the hypothesis that oxygen was one of the main limiting factors for toluene catabolism (Nogales et al. 2008), and that an incomplete toluene catabolism can be due to the secretion of pathway intermediates. Interestingly, there is an increasing number of evidences supporting the secretion of intermediate metabolites as a general strategy in Pseudomonas strains growing both on aromatic and nonaromatic carbon sources. For instance, P. putida secretes gluconate and 2-ketogluconate during glucose metabolism (del Castillo and Ramos 2007; Blank et al. 2008) and vanillic acid is often accumulated in the media during ferulic acid or vanillin catabolism (Graf and Altenbuchner 2014; Ravi et al. 2017). This behavior has been suggested to be responsible of redox homeostasis and/or as a consequence of potential metabolic overflows. Thus, global understanding of

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carbon flux during aromatics catabolism becomes critical in order to optimize the biodegradative capabilities of Pseudomonas. In this sense, a recent study addressing the dynamics of benzoate metabolism in P. putida by constructing a kinetic model based on deterministic rate equations provided a better understanding of the β-ketoadipate pathway in Pseudomonas (Sudarsan et al. 2016). The model predicted the stationary and dynamic changes in the concentration of intracellular and extracellular metabolites. Interestingly, a large secretion of intermediates, including β-ketoadipate, benzoate cis-diol, cis,cis-muconate, and catechol, was found driven by the limited capacity of some of the cognate enzymatic steps in P. putida KT2440, which argue in favor of an important metabolic overflow during benzoate metabolism. Consistently, in silico analysis identified negative Gibbs energies for several reactions of the β-ketoadipate pathway, including benzoate cis-diol dehydrogenase, catechol 1,2-dioxygenase, muconate cycloisomerase, β-ketoadipyl-CoA thiolase, and β-ketoadipate succinyl-CoA transferase. Overall, catechol formation and cleavage, and specially benzoate uptake were identified as the major flux bottlenecks in the β-ketoadipate pathway, thus being key targets for optimizing the metabolism of benzoate and other aromatics catabolized through this widely distributed pathway in Pseudomonas (Sudarsan et al. 2016). Similar data-driven dynamic modeling approaches will improve our current knowledge about aromatics metabolism in Pseudomonas. Nevertheless, considering a certain bacterial strain in isolation is an oversimplified view of the natural conditions where microorganisms thrive in complex communities in which the fitness of a single cell depends on the interaction with other cells of the same population and/or with cells of another population(s) generating consortia. Thus, a population-based approach accounting for the interaction between the cells should be considered (West et al. 2006). The division of labour within a bacterial population could be a relevant strategy when cells face difficult-to-degrade substrates, e.g., certain aromatic compounds, that might be toxic for individual cells and this metabolic specialization may also explain why many aromatic degradation pathways have evolved as upper and lower segments with a different expression pattern between different members of the population (Nikel et al. 2014b). In this respect, the TOL pathway of P. putida devoted to toluene/xylene degradation is structured in an upper pathway that converts toluene into benzoate, and a lower pathway responsible for mineralization of benzoate (see above). Interestingly, it was shown that when a clonal population of P. putida mt-2 was exposed to m-xylene many of the cells displayed a near bimodal distribution expressing either the upper or the lower operon. It is tempting to speculate that by engaging part of the population in an energy-demanding process (conversion of m-xylene into 3-methylbenzoate) while the remaining members take full advantage from the metabolic intermediates produced (3-methylbenzoate acting as a common good that provides energy and carbon to support growth), the entire community would balance the cost-to-benefit ratio of using a difficult-to-degrade aromatic hydrocarbon (Nikel et al. 2014b).

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Biotechnological Applications of the Metabolism of Aromatic Compounds in Pseudomonas

Due to the fact that Pseudomonas strains have a demonstrated capacity to tolerate and metabolize a large number of aromatic compounds, together with the fact that they are genetically amenable and reach a high biomass yield with a high growth rate and a low maintenance demand, they have been extensively proposed and used as biocatalysts for different biotechnological processes (Wackett 2003; Poblete-Castro et al. 2012; Nikel et al. 2014a; Lieder et al. 2015; Loeschcke and Thies 2015; Nikel et al. 2016a; Belda et al. 2016; Kim et al. 2016). In silico global genome analyses of Pseudomonas strains (Choi et al. 2007; Winsor et al. 2009, 2011, 2016; Silby et al. 2011; Jeukens et al. 2014; Winsor and Brinkman 2014; Sharma et al. 2015; Koehorst et al. 2016) and genome-scale reconstructions (see above) have been performed during the last decade. These efforts, when combined with the emergent synthetic biology approaches, can be used to explore the potential of these bacteria in different applications (Sohn et al. 2010; van Duuren et al. 2013). Table 1 presents some representative examples of the use of the Pseudomonas strains for different biotechnological applications related to the metabolism of aromatic compounds. The broad degradative potential of Pseudomonas toward aromatic compounds that are environmental pollutants makes them highly suitable for bioremediation purposes. For instance, native and/or recombinant Pseudomonas strains have been used in biodegradation of phenols, monoaromatic and polyaromatic hydrocarbons, chloro- and nitro-aromatic compounds, biogenic amines, or oil biodesulfurization. A nonexhaustive list of compounds that can be degraded by Pseudomonas strains is shown in Table 1. 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, P. knackmussi 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 could 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). Degradation of certain aromatic xenobiotics, such as polychlorobiphenyls, is facilitated by the use of consortia made of different bacterial strains some of which are specialized in the initial attack to the chlorinated hydrocarbons whereas other members are specialized in the dehalogenation and mineralization of the intermediate chloroaromatic acids (Hernández-Sánchez et al. 2013). Many aromatic compounds are high value-added products because they are used as platform chemicals for the production of a wide variety of industrial compounds. These aromatic compounds are currently produced in energy intensive chemical

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Table 1 Some biotechnological applications derived from the metabolism of aromatics in Pseudomonas Strain P. putida JLR11 P. putida PCL P. putida UWC3 P. knackmussii B13 P. pseudoalcaligenes KF707 P. putida F1 P. aeruginosa RW41 P. putida 1274 P. putida and P. fluorescens P. aeruginosa P. putida PD1 Pseudomonas sp. JP1 Pseudomonas sp. Ph6-gfp P. aeruginosa RS1 Pseudomonas GSa and GSb P. aeruginosa LY11 P. stutzeri CECT 930

Biotechnological process/approach Compound Bioremediation 2,4,6-trinitrotoluene (TNT) Naphthalene 2,4Dichlorophenoxyacetate 3- and 4-Chlorocatechols, 2-aminophenol Polychlorobiphenyls (PCBs) Benzene, toluene, phenol 4-Chlorobenzenesulfonate p-Nitrophenol Aromatic naphthenic acids Naphthalene Phenanthrene Polycyclic aromatic hydrocarbons Phenanthrene Pyrene 2,40 -Dichlorobiphenyl Decabromodiphenyl ether (BDE-209) Phenanthrene

Pseudomonas sp. OS2

Pyrene Nitrobenzene Naphthalene 4-Chloro-3-nitrophenol Polycyclic aromatic hydrocarbons and a diazo dye Polycyclic aromatic hydrocarbons Halogenated phenols

P. putida CSV86

Benzyl alcohol

Pseudomonas sp. JPN2 Pseudomonas sp. a3 P. gessardii LZ-E Pseudomonas sp. JHN P. stutzeri

Pseudomonas sp. WJ6

Reference Van Dillewijn et al. 2007 Kuiper et al. 2001 Dejonghe et al. 2000 Gaillard et al. 2006 Taira et al. 1992 Reardon et al. 2000 Blasco et al. 2008 Samuel et al. 2014 Zhang et al. 2015 Karimi et al. 2015 Khan et al. 2014 Liang et al. 2014 Sun et al. 2014 Ghosh et al. 2014 Gayathri and Shobha 2015 Liu et al. 2015a Moscoso et al. 2012 Jin et al. 2016 Wu et al. 2012 Huang et al. 2016 Arora et al. 2014 Álvarez et al. 2015

Xia et al. 2014 Kaczorek et al. 2016 Nigam et al. 2012 (continued)

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Table 1 (continued) Strain P. aeruginosa N6P6 P. aeruginosa NY3 P. monteilii SB3078 P. putida U Recombinant strains (pathway expansion) P. putida F1 (pVAD, pWW0) P. putida KTH2 (pESOX3) P. putida KT2440 (pIZdszB1A1C1-D1) P. fluorescens F113 L::1180 P. putida KT2442::44ES P. putida KT2440 (pCAR1) P. knackmussii B13 SN45P-1 P. putida DOT-T1 (pWW0-km) P. putida F1 (B1)

P. knackmussi B13 (pTOL) P. putida KT2440 (KTH2) P. putida KT2440 (KTU-PGC) P. putida KT2440 (KTU-9) P. putida X3 P. putida KT2440 (KT-ΔUPP-MP)

P. putida KT2440 (pNAH7)

Biotechnological process/approach Compound Polycyclic aromatic hydrocarbons Polycyclic aromatic hydrocarbons Benzene, toluene, ethylbenzene Tyramine, dopamine

Benzene, toluene, ethylbenzene, xylenes, styrene Dibenzothiophene (desulfurization) Dibenzothiophene (desulfurization) Polychlorobiphenyls (PCBs) 3-Chlorobenzoate Carbazole Chloro- and methylphenols (bioprotection) m-xylene, p-xylene

Reference Mangwani et al. 2015 Nie et al. 2010 Dueholm et al. 2015 Arcos et al. 2010

Lorenzo et al. 2004 Galán et al. 2000 Martínez et al. 2016 Villacieros et al. 2005 Klemba et al. 2000 Miyakoshi et al. 2007 Erb et al. 1997

Ramos et al. 1995

n-Propylbenzene, n-butylbenzene, cumene, biphenyl m-Chlorotoluene

Choi et al. 2003

Abril et al. 1989

Phenol

Prieto et al. 1996

Carbofuran, chlorpyrifos

Gong et al. 2016b

Methyl-parathion

Gong et al. 2016a

Methyl-parathion Methyl-parathion, fenitrothion, chlorpyrifos, permethrin, fenpropathrin, cypermethrin Naphthalene

Zhang et al. 2016 Zuo et al. 2015

Fernández et al. 2012 (continued)

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Table 1 (continued) Strain P. aeruginosa PaJC P. putida KT2440pK18mobtynhpa

Biotechnological process/approach Compound Benzene, toluene, p-xylene Tyramine, dopamine

Reference Kahraman and Geckil 2005 Arcos et al. 2010

Bioproduction of aromatics 3-Methylcatechol (from m-xylene) p-Hydroxybenzoate (from toluene) 6-Hydroxynicotinate (from nicotinate) 6-Hydroxy-3-succinoylpyridine (from nicotine) 3-succinoyl-pyridine (from nicotine) Indigo (from indole) Benzene, toluene, chlorobenzene cis-diols (from the aromatic hydrocarbons) (S)-styrene oxide (from styrene)

P. putida DOT-T1E derivative P. putida DOT-T1E derivative P. putida KT2440dnicC P. putida P-HSP P. putida S16dspm P. putida B4-01 P. putida KTOY02

P. taiwanensis VLB120ΔC

Rojas et al. 2004 Ramos-González et al. 2001 Jiménez et al. 2008 Yu et al. 2014 Wang et al. 2015 Cheng et al. 2016 Ouyang et al. 2007

Schmutzler et al. 2016

Other bioconversions PHAs production P. putida U P. fluorescens BM07

P. putida CA-3

From n-phenylalkanoic acids From 11-phenoxyundecanoic acid From styrene, phenylacetate and other aromatic compounds

Pseudomonads

From phenylacetate

P. putida mt2

From benzene, toluene, ethylbenzene, p-xylene From terephthalate

P. putida GO16

Luengo et al. 2003 Choi et al. 2009, 2010 O’Leary et al. 2005; Ward and O’Connor 2005; Dunn et al. 2005; Ward et al. 2005, 2006; Goff et al. 2007 Tobin and O’Connor 2005 Nikodinovic et al. 2008 Kenny et al. 2012 (continued)

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Table 1 (continued) Biotechnological process/approach Compound From APL (alkaline pretreated lignin) Bioproduction of other building blocks P. putida KT2440 β-Ketoadipate and derivative muconolactone (from protocatechuate) P. putida KT2440 Cis,cis-muconate (from derivatives lignin derived monoaromatic compounds) P. putida KT2440 Cis,cis-muconate (from derivative benzoate) P. putida KT2440 Pyruvate and lactate (from derivative p-coumarate and benzoate) Bioproduction of aromatics from renewable sources P. putida S12 Phenol, cinnamate (from glucose) P. putida S12 p-Hydroxybenzoate (from glycerol) Pseudomonas sp. HR199 Vanillin (from eugenol) derivative P. putida KT2440 Vanillin (from ferulic derivative acid) Strain P. putida KT2440

P. fluorescens BF13 derivative P. putida KT2440 derivative P. putida S12 derivative

Vanillin (from ferulic acid) Anthranilate (from glucose) p-Hydroxybenzoate (from xylose) Phloroglucinol (from glucose)

P. protegens Pf-5

Reference Linger et al. 2014

Okamura-Abe et al. 2016 Vardon et al. 2015; Johnson et al. 2016 Van Duuren et al. 2011 Johnson and Beckham 2015

Wierckx et al. 2008 Verhoef et al. 2014 Priefert et al. 2001 Graf and Altenbuchner 2014 Di Gioia et al. 2011 Kuepper et al. 2015 Meijnen et al. 2011 Achkar et al. 2005

Biosensors P. putida KT2440:: DmpR (pVI360) Pseudomonas sp.Y2 PAL1

Phenol, cresols Styrene

Shingler and Moore 1994 Alonso et al. 2003 (continued)

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Table 1 (continued) Biotechnological process/approach Compound Toluene, benzene, phenol, p-xylene, m-xylene P. putida pPG7-JAMA21 Naphthalene, 2-methylnaphthalene, salicylate P. putida F1G4 Benzene, toluene, ethylbenzene, xylenes, naphthalene P. putida BX5Pu-LUX14 2,4-Dinitrotoluene (2,4-DNT) P. putida F1G4 (PpF1G4) Aromatic compounds and surfactans P. putida KT2440 Benzene, toluene, derivatives ethylbenzene, xylenes Strain P. putida TVA8

Toluene, flavonoids

P. putida DOT-T1E derivative P. putida KT2440 and DOT-T1E (pKST-1)

Benzene, toluene, xylenes

p-Nitrophenol

P. monteilii LZU-3

Reference Applegate et al. 1998 Werlen et al. 2004

Phoenix et al. 2003 de las Heras et al. 2008 Keane et al. 2008 de las Heras and de Lorenzo 2011, 2012 Espinosa-Urgel et al. 2015 HernándezSánchez et al. 2016 Chen et al. 2016

Biocontainment P. putida KT2440ICS

Styrene

P. putida KT2440ICB

Biphenyl, 4-chlorobiphenyl 3-Methylbenzoate

P. putida MCR8

Lorenzo et al. 2004 Munthali et al. 1996 Ronchel and Ramos 2001

processes that generate toxic byproducts. Thus, their synthesis by bacterial biocatalysts in a cost-effective, environmentally friendly, and regioselective manner has attracted during the last decades a great industrial interest. The biocatalytic potential of Pseudomonas makes these bacteria as flexible cell factories for the bioproduction of value-added aromatic compounds (Gosset 2009; Vargas-Tah and Gosset 2015; Molina-Santiago et al. 2016; Lee and Wendisch 2016). Some examples are the production of bulk chemicals, such as phenol, p-hydroxybenzoate, styreneoxide, 6-hydroxynicotinic acid, 3-succinoyl-pyridine, and anthranilate; flavors, such as vanillin; or dyes, such as indigo (Table 1). The use of bacterial biofilms has attracted increasing attention as self-immobilized biocatalysts since they feature increased resistance to toxic chemicals and physical robustness compared with their planktonic counterparts (Schmutzler et al. 2016; Benedetti et al. 2016). The development of bacterial synthetic consortia is also a promising strategy to enhance

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the efficiency of bioconversion of certain aromatic compounds such as dibenzothiophene in biodesulfurization processes (Martínez et al. 2016). P. putida when cultured in chemically defined media containing different aromatic carbon sources (e.g., phenylalkanoates, styrene, or other toxic aromatic hydrocarbons) is able to produce valuable bioplastics such as the polyhydroxyalkanoates (PHA) (García et al. 1999; O’Leary et al. 2005; Ward and O’Connor 2005; Dunn et al. 2005; Tobin and O’Connor 2005; Nikodinovic et al. 2008; Ward et al. 2005, 2006; Goff et al. 2007; Choi et al. 2009, 2010; Kenny et al. 2012). The disruption or deletion in these bacteria of the fadB and/or fadA genes involved in the β-oxidation multienzymatic complex generates bioplastic overproducers. The monomer composition and length of these polymers can be modified by changing the relative proportion of the phenylalkanoate precursor added to the culture broth (Luengo et al. 2003). Since the monomeric units of PHA are enantiomerically pure R-3hydroxyalkanoic 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). Recently, there is a strong motivation to find alternative routes to produce aromatic compounds used as platform chemicals in green production processes from renewable resources that can reduce the dependence on fossil fuels and allow a sustainable bioeconomy. Solvent-tolerant Pseudomonas strains (e.g., P. putida DOT-T1E and S12 strains) are especially useful as whole-cell biocatalysts in double-phase systems for the production of toxic compounds that partition in the organic phase of a solvent/water mixture. Thus, engineered P. putida strains have been used for the bioconversion of glucose to produce anthranilate (Kuepper et al. 2015), phenol (Wierckx et al. 2008), or cinnamate (Nijkamp et al. 2005), for the bioconversion of glycerol to produce p-hydroxybenzoate (Verhoef et al. 2014), or for the bioconversion of xylose to produce p-hydroxybenzoate (Meijnen et al. 2011). In all these examples, the aromatic compounds have been synthesized by utilizing endogenous precursors derived from the ubiquitous shikimate pathway. An exception to this general strategy is the production of phloroglucinol derived from malonyl-CoA by the Phl biosynthetic pathway in P. protegens Pf-5 (Achkar et al. 2005). Bioconversion of plastic waste (e.g., polystyrene, polyethylene terephthalate (PET)) into higher value biodegradable polymers (e.g., PHAs) is also an emerging strategy for plastic upcycling (Wierckx et al. 2015). Lignocellulosic biomass offers a vast, renewable resource for the sustainable production of fuels, chemicals, and materials. Polysaccharides have been the primary fraction of interest in selective conversion processes, leaving the aromatic polymer lignin as a major target for valorizing the underutilized lignocellulosic fraction. Lignin depolymerization, either by biological or chemical approaches, provides a heterogeneous slate of aromatic species that can be bioconverted into several classes of molecules including those that maintain the aromatic ring, e.g., vanillin, ringopened species, e.g., dicarboxylic acids, and products derived after the carbon enters the central metabolism, e.g., PHAs (Beckham et al. 2016). Different P. putida strains have been engineered to valorize lignin-derived aromatic compounds (Table 1). The most obvious intermediate to produce is cis,cis-muconate, the ortho-cleavage

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product of catechol. Muconic acid can be converted into myriad downstream products, including adipic acid, which is the most commercially important dicarboxylic acid, and terephthalic acid (Vardon et al. 2015; Johnson et al. 2016; Beckham et al. 2016). Other building blocks that are produced from lignin-derived aromatic compounds in Pseudomonas are muconolactone and β-ketoadipate (Okamura-Abe et al. 2016). The production of pyruvate and L-lactate from lignin-derived aromatic compounds revealed that the lower pathways (ortho- or meta-cleavage pathways) of catechol and protocatechuate metabolism are interchangeable, and because they differ in carbon yield, cofactor regeneration, and the final products that enter the TCA cycle, the pathways employed can have a profound effect on product yield (Johnson and Beckham 2015). The conversion of ligninderived streams into medium chain-length PHAs (mcl-PHAs) in P. putida has been also demonstrated. Interestingly, these mcl-PHAs can be depolymerized by several routes and the resulting monomeric acids can be used for products beyond bioplastics, e.g., chemical precursors or fuel-range hydrocarbons (Linger et al. 2014). The knowledge on the regulation of the aromatic catabolic pathways has been utilized to design different Pseudomonas whole-cell biosensors for a wide range of target compounds (e.g., styrene, naphthalene, benzene, toluene, ethylbenzene, PCBs, phenol, etc.), and using different types of reporter genes (e.g., lacZ, lux, luc, gfp, etc.) (Carmona et al. 2008; van der Meer and Belkin 2010; Xue et al. 2014; Hernández-Sánchez et al. 2016) (Table 1). The regulatory systems of some efflux pumps, e.g., the TtgR and SepR regulators and the cognate promoters that control expression of solvent extrusion pumps of P. putida, can be also engineered to develop whole-cell biosensors able to detect a wide range of structurally diverse antibiotics, aromatic hydrocarbons, and flavonoids (Phoenix et al. 2003; EspinosaUrgel et al. 2015). A step forward on the design of biosensors 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. A variant of the toluene-recognizing transcriptional regulator XylR that responds to 2,4dinitrotoluene has been obtained and 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; de las Heras and de Lorenzo 2011) have developed a comprehensive system for the long-term preservation of P. putida cells genetically designed for biosensing benzene, toluene, ethylbenzene, and xylenes (BTEX) in soil, along with a procedure to formulate, spread, and vigorously activate such bacteria at the desired site and occasion. These authors have also developed a new platform that utilizes mini-transposon vectors tailored for engineering an artificial expression cascade that operates as an amplifier of the signal/response ratio of the biosensor (de las Heras and de Lorenzo 2012). An alternative biosensing technology based on microbial fuel cells (MFCs) was developed for in situ real-time environmental monitoring. This type of biosensor can directly generate a measurable electrical signal from substrates. P. monteilii LZU-3, which is able to degrade p-nitrophenol aerobically and generate voltage in a two-chamber MFC with an aerobic anode chamber, was tested as a biosensor for p-nitrophenol monitoring (Chen et al. 2016).

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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 able to degrade biphenyl/chlorobiphenyl or styrene and endowed with an active gene containment system have been described (Table 1) (García and Díaz 2014).

5

Research Needs

The increased use of the “omic” techniques, e.g. genomics, proteomics, metabolomics and fluxomics, as well as the systems biology approaches for addressing biological complexity from a holistic perspective, have contributed significantly to accelerate and complete our understanding on different aspects of the physiology, ecology, biochemistry, and regulatory mechanisms underlying the catabolism of aromatic compounds in Pseudomonas. Pseudomonas becomes, thus, a paradigmatic bacterial genus both for increasing basic knowledge and for applied research in the field of aromatic compounds bioconversions. Although the role of lateral gene transfer as a major mechanism for adaptation and evolution of novel catabolic abilities toward aromatic compounds has been clearly shown in members of the Pseudomonas genus, a more detailed reconstruction of the evolutionary history of the catabolic clusters will benefit from the continued effort to sequence environmentally relevant strains. However, our current knowledge about the degradative potential of Pseudomonas is still far from complete. There are still many issues regarding the catabolism of aromatic compounds that remain unknown or poorly studied and that, therefore, require future attention. Genomic analyses point to the existence in bacterial chromosomes of many parologous genes likely involved in the degradation of aromatic compounds, which, in turn, raises the question on the physiological role of this genetic redundancy and points to the existence of additional catabolic pathways whose physiological relevance needs to be studied. The role of genes of unknown function present in aromatic gene clusters, and the exploration of the degradative capabilities of non-cultivable bacteria by metagenomic approaches are also topics that deserve further research. The bacterial attack and degradation of complex aromatic molecules, e.g., the lignin polymer, is an emerging and exciting issue that is still in its infancy and should be further studied. Another important aspect that should be stressed in the near future is the transmembrane trafficking of aromatic substrates and metabolites, especially in those

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situations that involve metabolic overflows and/or syntrophic interactions between two or more different partners of the community for the complete mineralization of the aromatic substrates. As individual cells rarely metabolize a wide range of substrates, metabolic specialization within the bacterial population could be a relevant trait in the assembly of efficient microbial biodegrader communities. Phenotypic diversification may constitute part of an evolutionary long-term strategy toward the success of the whole population and it may be a relevant strategy for the evolution of biodegradation pathways. Genetic and biochemical analysis of single cells will provide further clues to unravel this puzzling issue. The interplay between regulatory proteins that recognize aromatic compounds, e.g., sensor histidine-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 ecophysiological meaning of the diversity found in the regulation of the hierarchical utilization of aromatic compounds among closely related strains that share ecological niches, and the understanding of the molecular mechanisms underlaying carbon catabolite repression phenomena should be further explored. The role and mechanism of action of some nucleotides, e.g., c-di-GMP, as second messengers that control motility, and perhaps other adaptive responses to toxic carbon sources, should be also studied in the near future. Further research should be carried out to better define the timing, content, and cross-talk of the different cellular programs, i.e., metabolic, stress, and social programs, that are induced when bacteria face aromatic compounds. The interplay between central carbon catabolism and peripheral redox reactions during oxidative biodegradation of aromatic compounds should be a major focus of future research since robustness in the redox balance is a main issue for an efficient and versatile metabolism of these carbon sources. Aromatic degradation pathways are also an important source of metabolic exchange factors and they might modulate cell-to-cell communication. The role of quorum sensing in controlling aromatic degradation pathways and, in turn, the latter as a source of metabolic signals and/or quorum quenching mechanisms that modulate bacterial communication are exciting topics that need to be addressed and that point to the aromatic degradation pathways as possible targets for drug development. Although some genome-scale metabolic reconstructions and models have been recently developed for several Pseudomonas strains, this is still an emerging field and much more effort should be devoted to refine these models and to generate new ones for other relevant strains. The reconstruction of new and improved Pseudomonas models, including dynamic modeling approaches, not only will allow the better understanding of the Pseudomonas potential toward aromatics, but also will drive systems metabolic engineering approaches toward the revalorization of such compounds and the identification of still unknown metabolic bottlenecks. Moreover, integrating the effector-specific regulatory and sensing 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 understanding and redesign the

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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. The extension of the metabolic modeling approach at population level should yield a much more complete view of the biodegradation of aromatic compounds in nature while facilitating the construction of bacterial consortia-based biotechnological applications. The knowledge of the full range of metabolic capabilities of the microorganisms, as well as the monitoring of such capacities and their distribution at contaminated sites, will allow to predict how these organisms are likely to respond to changes in environmental conditions during natural attenuation and/or bioaugmentation approaches for bioremediation of polluted sites. Finally, the valorization of aromatic compounds present in biowaste (e.g., lignin, plastics) to produce biofuels, biopolymers, commodity chemicals, and other value-added products of biotechnological interest, will enable a sustainable knowledge-based bioeconomy with social and environmental rewards. Acknowledgments Work in our laboratory was supported by the Ministry of Economy and Competitiveness of Spain Grant BIO2012-39501, BIO2016-79736-R, BIO2014-59528-JIN, and PCIN2014-113, European Union FP7 Grant 311815, and Fundación Ramón-Areces XVII CN.

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Contents 1 2 3 4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aerobic Aromatic Catabolic Routes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sequenced Bacterial Genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Spread of Members of Gene Families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Intradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 EXDO I Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Lig B Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Cupin Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Other Extradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Diiron Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Flavoprotein Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Rieske Non-heme Iron Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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D. Pérez-Pantoja Departamento de Bioquímica y Biología Molecular, Facultad de Ciencias Biológicas, Universidad de Concepción, Concepción, Chile e-mail: [email protected] R. Donoso · B. González Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile e-mail: [email protected]; [email protected] H. Junca Research Group Microbial Ecology: Metabolism, Genomics and Evolution, Microbiomas Foundation, Chia, Colombia e-mail: [email protected] D. H. Pieper (*) Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany e-mail: [email protected] # Springer Nature Switzerland AG 2019 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils, and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, https://doi.org/10.1007/978-3-319-50418-6_33

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5 Metabolism Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Metabolism by Bacteria Outside the Actinobacterial and Proteobacterial Phyla . . . . 5.2 Actinobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Proteobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Pathway Redundancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5 Gene Redundancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6 Superbugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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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 available in 2008 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; Pérez-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

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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 (for an appropriate novel database see Duarte et al. 2014). 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 (Duarte et al. 2014). Various pathways for degradation of these compounds by bacteria have been reported. The activation of the aromatic ring commonly proceeds by members of one of three superfamilies: the Rieske non-heme 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. 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

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1-carboxylate structural motif. The 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 Fe 2+-dependent dioxygenases (Moonen et al. 2008).

3

Sequenced Bacterial Genomes

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 was 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 δ- (23) and ε- classes (28); and the highly represented phylum Firmicutes (182) and the α- (112) β- (71) and γ- (223) classes of Proteobacteria (besides two 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 cross-reactivity 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

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

Fig. 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

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1,2-dioxygenases, cluster 3 actinobacterial catechol 1,2-dioxygenases, and clusters 5 and 7 the α- and β-subunits of protocatechuate 3,4-dioxygenases, respectively. Enzymes of cluster 6 are obviously related to the β-subunits of protocatechuate dioxygenases, however, in no case genes encoding these enzymes are clustered with genes encoding putative α- subunits, and the function of these enzymes remains to be elucidated. Similarly, the function of enzymes of cluster 4 wait for clarification.

Fig. 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 α- and β-subunits of protocatechuate 3,4-dioxygenases, respectively. The functions of enzymes of clusters 4 and 6 remain to be elucidated

ä Fig. 1 (continued) 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

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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 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 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 α-proteobacterial, 2 out of 10 β-proteobacterial, and 4 out of 29 γ-proteobacterial families and are absent in δ- or ε- 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 α-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). The EXDO I family comprises enzymes that catalyze the dioxygenolytic ring fission of the catecholic derivatives in several bacterial monoand polyaromatics biodegradation pathways (Eltis and Bolin 1996; Duarte et al. 2014) (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),

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Table 1 Intradiol dioxygenases observed in genomes of Proteobacteria

Class α α α α α α α α α α α β β γ γ γ γ

Family Caulobacteraceae (2) Aurantimonadaceae (2) Bradyrhizobiaceae (11) Brucellaceae (6) Methylobacteriaceae (3) Phyllobactriaceae (3) Rhizobiaceae (6) Rhodobacteraceae (24) Xanthobacteriaceae (2) Acetobacteraceae (3) Sphingomonadaceae (5) Burkholderiaceae (43) Comamonadaceae (8) Oceanosprillaceae (3) Moraxellaceae (5) Pseudomonadaceae (19) Xanthomonadaceae (11)

Protocatechuate 3,4-dioxygenase (Pca34) ++ + ( ) ++ + ++ ++ ++ ++ + ( ) ++ ( ) + ++

Catechol 1,2-dioxygenase (Cat12)

Hydroxybenzoquinol dioxygenase (Hqu)

++ +

+ + ++ + + + ++

++ + ++ ( ) + + + ( )

+

++; 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

1,2-dihydroxynaphthalene (NahC), homoprotocatechuate (homoprotocatechuate 2,3-dioxygenases, HpaD), 2,3-dihydroxy-p-cumate (2,3-dihydroxy-p-cumate-3,4dioxygenases 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

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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,

Fig. 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

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13 strains belong to the β-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 β- and γ-proteobacterial genomes, but surprisingly most abundant in α-proteobacterial genomes (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. 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 1,2-dihydroxynaphthalene dioxygenase of Bacillus sp. JF8 (Miyazawa et al. 2004) (subcluster 1D) are not observed in any Bacillus species, but encoded in four α-proteobacterial genomes. Also the three subcluster 1E proteins, where no closely related proteins have been characterized so far, are encoded in two α-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 α- and β-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

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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 α-, eight β-, and five γ-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 γ-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 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; Duarte et al. 2014). LigB type extradiol dioxygenases are well

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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 β-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 β-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 β-subunits of protocatechuate dioxygenases. Analysis of the primary structure revealed that the N-terminal regions showed a significant amino acid sequence identity with the β-subunit of protocatechuate 4,5-dioxygenases, whereas the C-terminal region has similarity to the corresponding small α-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 β-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 α-, β-, and γ-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 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 α-Proteobacteria and are thus not a dominant trait in this group.

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Fig. 4 Evolutionary relationships among LigB-type dioxygenases. Subcluster 1A comprises protocatechuate 4,5-dioxygenase β-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 β- and α-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

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 α- and β-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 α- and β-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 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.

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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 β- and α-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 LigBtype 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; Duarte et al. 2014) 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 α-, 5 out of 10 β-, 16 out of 29 γ-, and 4 out of 11 δ-proteobacterial families, although they were absent in ε-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

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in sequenced representatives of the families Hyphomonadaceae, Neisseriaceae, Aeromonadaceae, Idiomarinaceae, Moritellaceae, Chromatiaceae, Legionellaceae, Hahellaceae, Bdellovibrionaceae, 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 α-, three β-, and three γ-proteobacterial families and being absent from δ- and ε-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 Fe 2+-dependent dioxygenase, benzoquinol 1,2-dioxygenase, which is a α 2β 2 heterotetramer where the α- and β-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.

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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 α-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 a reductase (and, in some cases, a ferredoxin), a catalytic effector and a terminal heteromultimeric oxygenase composed by α, β, and γ subunits whose α-subunits are assumed to be the site of substrate hydroxylation (Leahy et al. 2003). According to the presence of genes putatively coding for α subunit, benzene/toluene multicomponent monooxygenase are found almost exclusively in β-Proteobacteria, including Burkholderia, Cupriavidus, Ralstonia, Methylibium, and Dechloromonas strains with the only exceptions of Bradyrhizobium sp. BTAi1 and Frankia sp. CcI3. In the β-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 γ-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 α-, 5 out of 10 β-, and 8 out of 29 γ-proteobacterial families. They are, however, absent from δand ε-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 α- and β-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

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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 para-hydroxylate 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 (Pérez-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 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 α-Proteobacteria, specifically in Bradyrhizobiaceae and Rhodobacteraceae. Also among β-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 γ-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,

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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 γ-Proteobacteria, but it is observed in Plesiocystis pacifica SIR-1 (a δ-proteobacterium). The pathway is abundant in β-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 α-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.

Fig. 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

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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 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 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 two-component 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 γ-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-3monooxygenase from P. azelaica HBP1, which is capable of oxidizing various 2-substituted phenols, but not phenol (Suske et al. 1997), OhpB

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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 β-proteobacteria such genes are only observed in C. testosteroni and B. phymatum, however, also three γ-Proteobacteria harbor such gene, and 3-hydroxybenzoate-4-hydroxylases seem to be frequently encoded in the genome of α-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. 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

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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 γ-proteobacteria,

Fig. 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

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predominantly in Enterobacteriaceae (19 out of 61 genomes) and Pseudomonas (5 out of 18 genomes), and outside of this group only in two α-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 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 α- and γ-proteobacteria: S. stellata, R. sphaeroides, Marinomonas sp., V. shilonii, V. vulnificus, A.vinelandii, P. entomophila, and one P. putida strain. Additional 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 β-proteobacterium Polaromonas sp. JS666.

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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; Duarte et al. 2014). 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 ferredoxinNADH-reductase). The catalytic ISPs are usually heteromultimers composed of a large α-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 β-subunit, however, some enzymes, such as phthalate 4,5-dioxygenases contain an oxygenase composed only of α-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 β-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 β-proteobacterial genomes (overall in five), but also in one γ-proteobacterium and in two α-proteobacteria, among them strain HTCC2654. A similar spread is observed for enzymes related to phenoxybenzoate dioxygenase (observed in seven β-, four α-, and one γ-Proteobacterium). 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 α-, β-, and γ-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 (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

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Fig. 7 Evolutionary relationships among the α-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

two-component dioxygenases are obviously restricted to γ-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 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.

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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 β-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), three-component anthranilate dioxygenases (cluster E5, see above), and terephthalate dioxygenases (Sasoh et al. 2006) (cluster E3). Respective sequences are nearly exclusively observed in β-proteobacteria and in Sphingomonads out of the α-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 C. testosteroni

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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

3-Hydroxybenzoate 4-hydroxylase

Enzyme group Protocatechuate 3,4-dioxygenase Catechol 1,2-dioxygenase Hydroxybenzoquinol 1,2-dioxygenase Chlorocatechol 1,2-dioxygenase Catechol 2,3-dioxygenase 2,3-Dihydroxybiphenyl 1,2-dioxygenase Homoprotocatechuate 2,3-dioxygenase Protocatechuate 4,5-dioxygenase Gallate 4,5-dioxygenase Homoprotocatechuate 2,3-dioxygenase 2,3-Dihydroxyphenylpropionate 1,2-dioxygenase 2-Aminophenol 1,6-dioxygenase Gentisate 1,2-dioxygenase Homogentisate 1,2-dioxygenase 3-Hydroxyanthranilate 3,4-dioxygenase Benzoquinol 1,2-dioxygenase Benzoyl-CoA oxygenase Phenylacetyl-CoA oxygenase 2-Aminobenzoyl-CoA monooxygenase/ reductase 4-Hydroxybenzoate 3-hydroxylase LigB-type dioxygenase Cupin superfamily dioxygenase Cupin superfamily dioxygenase Cupin superfamily dioxygenase Type IV extradiol dioxygenase Soluble diiron oxygenase Soluble diiron oxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase

Family Intradiol dioxygenase Intradiol dioxygenase Intradiol dioxygenase Intradiol dioxygenase Type I extradiol dioxygenase Type I extradiol dioxygenase Type I extradiol dioxygenase LigB-type dioxygenase LigB-type dioxygenase LigB-type dioxygenase LigB-type dioxygenase + ++ ++ ++ + ++ ++ ++

Pathway marker ++ ++ ++ + ++ # ++ ++ ++ ++ ++

Table 2 Key groups of catabolic enzymes discussed in the metabolic diversity section

Phylogenomics of Aerobic Bacterial Degradation of Aromatics (continued)

Mhb4H

Phb3H

Forming protocatechuate Forming protocatechuate

Amn Gen Hge Han Bqu Box Paa Abc

Abbreviation Pca34 Cat12 Hqu Cca Cat23 Dhb HpcEXDOI Pca45 Gal HpcLigB Dhp

Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Dearomatization Dearomatization Dearomatization

Enzyme function Intradiol cleavage Intradiol cleavage Intradiol cleavage Intradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage

26 715

Terephthalate 1,2-dioxygenase Phthalate 3,4-dioxygenase Anthranilate 1,2-dioxygenase (2 component) Anthranilate 1,2-dioxygenase (3 component) Benzoate 1,2-dioxygenase

Resorcinol 4-hydroxylase

4-Hydroxyphenylacetate 3-hydroxylases

Chlorophenol 4-hydroxylase

Phenol 2-hydroxylase

4-Hydroxyphenylacetate 3-hydroxylases

Phenol/benzoquinol hydroxylase

3-Hydroxyphenylpropionate 2-hydroxylase 3-Hydroxyphenylacetate 6-hydroxylase

3-Hydroxybenzoate 6-hydroxylase

Enzyme group Salicylate 1-hydroxylase

Table 2 (continued)

Channeling to catechol Channeling to catechol

Rieske nonheme iron oxygenase

Forming hydroxybenzoquinol

Forming homoprotocatechuate

Forming chlorobenzoquinol

Forming catechol

Forming homoprotocatechuate

Forming catechol/hydroxybenzoquinol

Forming 2,3-dihydroxyphenylpropionate Forming homogentisate

Forming gentisate

Rieske nonheme iron oxygenase

Enzyme function Forming catechol

Channeling to protocatechuate Channeling to protocatechuate Channeling to catechol

Pathway marker

Family Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class D flavoprotein monooxygenase Class D flavoprotein monooxygenase Class D flavoprotein monooxygenase Class D* flavoprotein monooxygenase Class D* flavoprotein monooxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase

BenDO

AntDO

TphDO Pht34DO AntDO

Res4H

Pha3H

Ph4H

Ph2H

Pha3H

Pbq2H

Mha6H

Mhp2H

Mhb6H

Abbreviation Ohb1H

716 D. Pérez-Pantoja et al.

Channeling to protocatechuate Channeling to protocatechuate Forming phenol Forming catechol

Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Soluble diiron oxygenase Soluble diiron oxygenase

Biphenyl 2,3-dioxygenase type Naphthalene inducible dioxygenase (NidA) type Phthalate dioxygenase 4,5-dioxygenase Isophthalate dioxygenase Toluene/benzene monooxygenase Phenol monooxygenase

PhtDO IphDO Tmo Pmo

BphDO NidDO

Sal5H PhpDO

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

Channeling to gentisate Channeling to 2,3-dihydroxyphenylpropionate Activation of hydrophobic aromatics Polycyclic aromatic degradation

Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase

Salicylate 5-hydroxylase Phenylpropionate 2,3-dioxygenase

26 Phylogenomics of Aerobic Bacterial Degradation of Aromatics 717

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D. Pérez-Pantoja et al.

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 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 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

26

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

719

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. 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

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D. Pérez-Pantoja et al.

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 (α, β, and γ) 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 δ proteobacteria and none in the ε proteobacterial class. The α 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 ring-cleavage 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 α-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.

+

+

+

++

+

+

+ +

++

+

+ ++

++

++

+

+

+

+

+

+

+

(+)

+

+

(+)

+

+

++

++

++

+

++

++

+

++

+

++; 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)

++

+

+

++

++

+

Streptomycetaceae (2) ++

+

+

++

Nocardioidaceae (3)

+

++

+

+

+

++

+

++

+

Mycobacteriaceae (17)

++

++

++

++

++

Pca34 Phb3H Mhb4H Pht34DO TphDO Cat12 BenDO Gen Mhb6H Hge HppDO Mha6H Dhp Mhp2H Hpcexdoi Pha3H NidDO Dhb Paa Abc

Micromonosporaceae (2)

Micrococcaceae (2)

Microbacteriaceae (2)

Frankiaceae (3)

Corynebacteriaceae (5)

Cellulomonadaceae (2)

Actinobacterial families

Table 3 Catabolic gene markers of Proteobacteria

26 Phylogenomics of Aerobic Bacterial Degradation of Aromatics 721

(+)

+

Pht34DO

(+)

TphDO

(+)

Cat23 (+)

Ohb1H (+)

AntDO ++

(+)

PMO (+)

(+)

Ph2H (+)

(+)

+

(+)

(+)

(+)

+

(+)

+

++

++

+

++

(+)

(+)

(+)

+

(+)

+

Neisseriaceae (5)

++

++

(+)

+

(+)

(+)

Comamonadaceae (8)

Oxalobacteraceae (2)

+

(+)

++

Burkholderiaceae (43)

(+)

++

Alcaligenaceae (3)

β Proteobacterial families

(+)

+

++ +

(+) (+)

++ (+)

(+)

+

+

++

+

+

+

(+)

(+)

++

++

++

++

++

++ +

+

++

++

++

+

++

(+)

+

++

+

+

+

++

Sphingomonadaceae (5)

(+)

++

(+)

+

(+)

++

++

HppDO

Erythrobacteraceae (3)

SAR11 (2)

Rickettsiaceae (16)

Anaplasmataceae (10)

Rhodospirillaceae (3)

(+)

+

BenDO +

(+)

Pbq2H

+

++

++

+

TMO (+)

Hqu

++

++

Xanthobacteraceae (2)

++

+

IphDO

(+)

Res4H

+

++

Rhizobiaceae (5)

++

+

+

Bqu

Acetobacteraceae (3)

++

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

Cat12

Table 4 Catabolic gene markers of Actinobacteria Mha6H +

(+)

(+)

(+)

Han (+)

(+)

(+)

(+)

(+)

Dhp (+)

+

Mhp2H (+)

Gal (+)

Hpc LigB ++

+

(+)

++

(+)

+

Hpc EXDOI +

+

+

+

Pha3H (+)

+

Paa +

++

++

(+)

+

+

+

+

++

Box ++

(+)

++

(+)

(+)

Abc ++

+

+

(+)

+

+

+

+

+

+

722 D. Pérez-Pantoja et al.

PhpDO

+

(+)

(+)

+

(+)

(+)

(+)

+

+

+

+

(+)

+ (+)

+

+

+ +

(+)

(+) +

++

++

++

++

++

++

in the analysis

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

++; More than 60% of the sequenced genomes of these Actinobacterial families comprise a gene encoding the mentioned activity (number of sequenced representatives is

++

+

++

++

Xanthomonadaceae (11)

+

++

Vibrionaceae (30)

Thiotrichaceae (2)

+

Pseudomonadaceae (19)

Francisellaceae (7)

+

++

Moraxellaceae (5)

Pasteurellaceae (21)

++ +

++

+

Oceanospirillaceae (3)

+

Legionellaceae (4)

Coxiellaceae (5)

Enterobacteriaceae (61)

(+)

+

+

+

(+)

+

+

+

++

++

(+)

(+)

+

+

(+)

+

(+)

+

(+)

+

(+)

+

+

(+)

++

(+)

+

(+)

(+)

++

+

+

++

+

+

Ectothiorhodospiraceae (3)

(+)

(+)

+

+

(+)

(+)

+

++

(+)

(+)

(+)

+

Shewanellaceae (18)

(+)

(+)

+

Psychromonadaceae (2)

(+)

+

+

++

+

(+)

+

Pseudoalteromonadaceae (3)

+

+

++

(+)

+

Idiomarinaceae (2)

Aeromonadaceae (7)

γ Proteobacterial families

Rhodocyclaceae (3)

Nitrosomonadaceae (3) ++

++

26 Phylogenomics of Aerobic Bacterial Degradation of Aromatics 723

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D. Pérez-Pantoja et al.

The most broadly distributed pathways in the α 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 has the Dhp pathway. The Cca, Amn, and Bqu pathways are not found in any α proteobacterial genome. Regarding peripheral pathways, α 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 α 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 β class of proteobacteria harbors all major central aromatic catabolic pathways listed in Table 2. The distribution of these catabolic pathways among β-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 (Pérez-Pantoja 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 β 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