Methods in Enzymology 495 - Methods in Methane Metabolism, Part B: Methanotrophy

METHODS IN ENZYMOLOGY Editors-in-Chief

JOHN N. ABELSON AND MELVIN I. SIMON Division of Biology California Institute of Technology Pasadena, California Founding Editors

SIDNEY P. COLOWICK AND NATHAN O. KAPLAN

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32 Jamestown Road, London NW1 7BY, UK First edition 2011 Copyright # 2011, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: permissions@ elsevier.com. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made For information on all Academic Press publications visit our website at elsevierdirect.com ISBN: 978-0-12-386905-0 ISSN: 0076-6879 Printed and bound in United States of America 11 12 13 14 10 9 8 7 6 5 4 3 2 1

CONTRIBUTORS

June I. Bagstevold Department of Molecular Biology, University of Bergen, Bergen, Norway Ramakrishnan Balasubramanian Department of Molecular Biosciences, and Department of Chemistry, Northwestern University, Evanston, Illinois, USA Nathan L. Bandow Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA David A. C. Beck Department of Chemical Engineering, and eScience Institute, University of Washington, Seattle, Washington, USA Lee Behling Department of Chemistry, University of Wisconsin-Eau Claire, Eau Claire, Wisconsin, USA Frode S. Berven Department of Molecular Biology, and Proteomic Unit (PROBE), Department of Biomedicine, University of Bergen, Bergen, Norway John P. Bowman Tasmanian Institute of Agricultural Research, University of Tasmania, Hobart, Tasmania, Australia Klaus Butterbach-Bahl Institute for Meteorology and Climate Research, Atmospheric Environmental Research (IMK-IFU), Karlsruhe Institute of Technology (KIT), GarmischPartenkirchen, Germany Sunney I. Chan Institute of Chemistry, Academia Sinica, Taipei, Taiwan, and Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena, California, USA Kelvin H.-C. Chen Department of Chemical Biology, National Pingtung University of Education, Pingtung, Taiwan

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Ludmila Chistoserdova Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Dong W. Choi Department of Biological and Environmental Science, Texas A&M UniversityCommerce, Commerce, Texas, USA Andrew Crombie School of Life Sciences, University of Warwick, Coventry, United Kingdom Svetlana N. Dedysh Winogradsky Institute of Microbiology, Russian Academy of Sciences, Moscow, Russia Alan A. DiSpirito Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA Peter F. Dunfield Department of Biological Sciences, University of Calgary, Calgary, Alberta, Canada Anne Fjellbirkeland Department of Molecular Biology, and Department of Biology, Centre for Geobiology, University of Bergen, Bergen, Norway Warren H. Gallagher Department of Chemistry, University of Wisconsin-Eau Claire, Eau Claire, Wisconsin, USA Valerie S. Gilles Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA David W. Graham School of Civil Engineering and Geosciences, Newcastle University, Newcastle Upon Tyne, United Kingdom Scott C. Hartsel Department of Chemistry, University of Wisconsin-Eau Claire, Eau Claire, Wisconsin, USA Harald B. Jensen Department of Molecular Biology, University of Bergen, Bergen, Norway Marina G. Kalyuzhnaya Department of Microbiology, University of Washington, Seattle, Washington, USA

Contributors

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Odd A. Karlsen Department of Molecular Biology, University of Bergen, Bergen, Norway Valentina N. Khmelenina Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Ralf Kiese Institute for Meteorology and Climate Research, Atmospheric Environmental Research (IMK-IFU), Karlsruhe Institute of Technology (KIT), GarmischPartenkirchen, Germany Hyung J. Kim Departments of Medicine and Biochemistry, University of Utah Health Sciences Center, Salt Lake City, Utah, USA Michael C. Konopka Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Stephan M. Kraemer Department of Environmental Geosciences, University of Vienna, Althanstrasse, Vienna, Austria Øivind Larsen Department of Molecular Biology, and Uni Environment, University of Bergen, Bergen, Norway Mary E. Lidstrom Department of Chemical Engineering, and Department of Microbiology, University of Washington, Seattle, Washington, USA Chunyan Liu Institute of Atmospheric Physics, Chinese Academy of Sciences (IAP-CAS), Beijing, China Sarah McQuaide Department of Electrical Engineering, University of Washington, Seattle, Washington, USA Akimitsu Miyaji Department of Environmental Chemistry and Engineering, Tokyo Institute of Technology, Nagatsuta-cho, Midori-ku, Yokohama, Japan J. Colin Murrell School of Life Sciences, University of Warwick, Coventry, United Kingdom Ildar I. Mustakhimov Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia

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H.-Hoa T. Nguyen Transmembrane BioSciences, Pasadena, California, USA David S. Ojala Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Alexander S. Reshetnikov Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Amy C. Rosenzweig Department of Molecular Biosciences, and Department of Chemistry, Northwestern University, Evanston, Illinois, USA Olga N. Rozova Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Jeremy D. Semrau Department of Civil and Environmental Engineering, The University of Michigan, Ann Arbor, Michigan, USA Stephen M. Smith Department of Molecular Biosciences, and Department of Chemistry, Northwestern University, Evanston, Illinois, USA Thomas J. Smith Biomedical Research Centre, Sheffield Hallam University, Sheffield, United Kingdom Yuri A. Trotsenko Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Sukhwan Yoon Department of Civil and Environmental Engineering, The University of Michigan, Ann Arbor, Michigan, USA, and Department of Biogeochemistry, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany Steve S.-F. Yu Institute of Chemistry, Academia Sinica, Taipei, Taiwan

PREFACE

The production and consumption of methane by microorganisms is central to the global carbon cycle. It has been 2 decades since a Methods in Enzymology volume focused explicitly on this field (Volume 188, Hydrocarbons and Methylotrophy). During that time, interest in methane metabolism has steadily increased in the context of dwindling petroleum reserves, increased greenhouse gas emissions, and environmental hydrocarbon pollution. A field previously dominated by microbiology and protein biochemistry has exploded in multiple directions. In particular, the advent of genomic and proteomic techniques has transformed the way methane metabolic pathways are studied. In these volumes, we cover both the generation (Part A) and utilization (Part B) of methane. Part A describes recent developments that enable a wide variety of experiments with methanogenic Archaea, which seemed intractable two decades ago to all but those initiates who “grew up” studying anaerobes. Methods are presented to readily culture and to perform genetic experiments on these oxygen-sensitive microbes, as well as to characterize the remarkable enzymes and respiratory proteins that allow methanogens to generate energy for growth and produce as a byproduct a very important fuel that may be adopted as the “fuel of the future.” Included is the state of the art in “biomethanation,” the biotechnological use of methanogens to produce this important energy source. Finally, approaches are described for the deployment of “omics” technologies to understand how methanogens regulate metabolism. The first methanotroph genome sequence was completed in 2004, and research has become progressively more “omic” in the past few years. Part B combines traditional approaches to methanotroph isolation and enzyme chemistry with state-of-the-art genomic and proteomic techniques. Novel methods have been developed and used to address challenging problems such as linking metagenomic data with environmental function or generating mutant forms of the methane monooxygenase enzymes. Moreover, whole new areas of research, such as the study of the copper chelator methanobactin, have been discovered. Taken together, we hope that the two volumes capture the excitement that pervades this rapidly developing field. We thank an outstanding group of colleagues with diverse points of view for providing ideas on content and ultimately contributing a series of excellent chapters. The high level of enthusiasm in the community resulted in the xv

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unexpected final production of two, rather than one, volumes. The methods and novel approaches described here should inspire and guide future research in the field as well as provide a central resource for researchers interested in methane metabolism. AMY C. ROSENZWEIG AND STEPHEN W. RAGSDALE

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VOLUME I. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME II. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME III. Preparation and Assay of Substrates Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME IV. Special Techniques for the Enzymologist Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME V. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VI. Preparation and Assay of Enzymes (Continued) Preparation and Assay of Substrates Special Techniques Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VII. Cumulative Subject Index Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VIII. Complex Carbohydrates Edited by ELIZABETH F. NEUFELD AND VICTOR GINSBURG VOLUME IX. Carbohydrate Metabolism Edited by WILLIS A. WOOD VOLUME X. Oxidation and Phosphorylation Edited by RONALD W. ESTABROOK AND MAYNARD E. PULLMAN VOLUME XI. Enzyme Structure Edited by C. H. W. HIRS VOLUME XII. Nucleic Acids (Parts A and B) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XIII. Citric Acid Cycle Edited by J. M. LOWENSTEIN VOLUME XIV. Lipids Edited by J. M. LOWENSTEIN VOLUME XV. Steroids and Terpenoids Edited by RAYMOND B. CLAYTON xvii

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VOLUME XVI. Fast Reactions Edited by KENNETH KUSTIN VOLUME XVII. Metabolism of Amino Acids and Amines (Parts A and B) Edited by HERBERT TABOR AND CELIA WHITE TABOR VOLUME XVIII. Vitamins and Coenzymes (Parts A, B, and C) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME XIX. Proteolytic Enzymes Edited by GERTRUDE E. PERLMANN AND LASZLO LORAND VOLUME XX. Nucleic Acids and Protein Synthesis (Part C) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME XXI. Nucleic Acids (Part D) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XXII. Enzyme Purification and Related Techniques Edited by WILLIAM B. JAKOBY VOLUME XXIII. Photosynthesis (Part A) Edited by ANTHONY SAN PIETRO VOLUME XXIV. Photosynthesis and Nitrogen Fixation (Part B) Edited by ANTHONY SAN PIETRO VOLUME XXV. Enzyme Structure (Part B) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVI. Enzyme Structure (Part C) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVII. Enzyme Structure (Part D) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVIII. Complex Carbohydrates (Part B) Edited by VICTOR GINSBURG VOLUME XXIX. Nucleic Acids and Protein Synthesis (Part E) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XXX. Nucleic Acids and Protein Synthesis (Part F) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME XXXI. Biomembranes (Part A) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME XXXII. Biomembranes (Part B) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME XXXIII. Cumulative Subject Index Volumes I-XXX Edited by MARTHA G. DENNIS AND EDWARD A. DENNIS VOLUME XXXIV. Affinity Techniques (Enzyme Purification: Part B) Edited by WILLIAM B. JAKOBY AND MEIR WILCHEK

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VOLUME LIII. Biomembranes (Part D: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LIV. Biomembranes (Part E: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LV. Biomembranes (Part F: Bioenergetics) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LVI. Biomembranes (Part G: Bioenergetics) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LVII. Bioluminescence and Chemiluminescence Edited by MARLENE A. DELUCA VOLUME LVIII. Cell Culture Edited by WILLIAM B. JAKOBY AND IRA PASTAN VOLUME LIX. Nucleic Acids and Protein Synthesis (Part G) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME LX. Nucleic Acids and Protein Synthesis (Part H) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME 61. Enzyme Structure (Part H) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 62. Vitamins and Coenzymes (Part D) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 63. Enzyme Kinetics and Mechanism (Part A: Initial Rate and Inhibitor Methods) Edited by DANIEL L. PURICH VOLUME 64. Enzyme Kinetics and Mechanism (Part B: Isotopic Probes and Complex Enzyme Systems) Edited by DANIEL L. PURICH VOLUME 65. Nucleic Acids (Part I) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME 66. Vitamins and Coenzymes (Part E) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 67. Vitamins and Coenzymes (Part F) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 68. Recombinant DNA Edited by RAY WU VOLUME 69. Photosynthesis and Nitrogen Fixation (Part C) Edited by ANTHONY SAN PIETRO VOLUME 70. Immunochemical Techniques (Part A) Edited by HELEN VAN VUNAKIS AND JOHN J. LANGONE

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VOLUME 71. Lipids (Part C) Edited by JOHN M. LOWENSTEIN VOLUME 72. Lipids (Part D) Edited by JOHN M. LOWENSTEIN VOLUME 73. Immunochemical Techniques (Part B) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 74. Immunochemical Techniques (Part C) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 75. Cumulative Subject Index Volumes XXXI, XXXII, XXXIV–LX Edited by EDWARD A. DENNIS AND MARTHA G. DENNIS VOLUME 76. Hemoglobins Edited by ERALDO ANTONINI, LUIGI ROSSI-BERNARDI, AND EMILIA CHIANCONE VOLUME 77. Detoxication and Drug Metabolism Edited by WILLIAM B. JAKOBY VOLUME 78. Interferons (Part A) Edited by SIDNEY PESTKA VOLUME 79. Interferons (Part B) Edited by SIDNEY PESTKA VOLUME 80. Proteolytic Enzymes (Part C) Edited by LASZLO LORAND VOLUME 81. Biomembranes (Part H: Visual Pigments and Purple Membranes, I) Edited by LESTER PACKER VOLUME 82. Structural and Contractile Proteins (Part A: Extracellular Matrix) Edited by LEON W. CUNNINGHAM AND DIXIE W. FREDERIKSEN VOLUME 83. Complex Carbohydrates (Part D) Edited by VICTOR GINSBURG VOLUME 84. Immunochemical Techniques (Part D: Selected Immunoassays) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 85. Structural and Contractile Proteins (Part B: The Contractile Apparatus and the Cytoskeleton) Edited by DIXIE W. FREDERIKSEN AND LEON W. CUNNINGHAM VOLUME 86. Prostaglandins and Arachidonate Metabolites Edited by WILLIAM E. M. LANDS AND WILLIAM L. SMITH VOLUME 87. Enzyme Kinetics and Mechanism (Part C: Intermediates, Stereo-chemistry, and Rate Studies) Edited by DANIEL L. PURICH VOLUME 88. Biomembranes (Part I: Visual Pigments and Purple Membranes, II) Edited by LESTER PACKER

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VOLUME 89. Carbohydrate Metabolism (Part D) Edited by WILLIS A. WOOD VOLUME 90. Carbohydrate Metabolism (Part E) Edited by WILLIS A. WOOD VOLUME 91. Enzyme Structure (Part I) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 92. Immunochemical Techniques (Part E: Monoclonal Antibodies and General Immunoassay Methods) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 93. Immunochemical Techniques (Part F: Conventional Antibodies, Fc Receptors, and Cytotoxicity) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 94. Polyamines Edited by HERBERT TABOR AND CELIA WHITE TABOR VOLUME 95. Cumulative Subject Index Volumes 61–74, 76–80 Edited by EDWARD A. DENNIS AND MARTHA G. DENNIS VOLUME 96. Biomembranes [Part J: Membrane Biogenesis: Assembly and Targeting (General Methods; Eukaryotes)] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 97. Biomembranes [Part K: Membrane Biogenesis: Assembly and Targeting (Prokaryotes, Mitochondria, and Chloroplasts)] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 98. Biomembranes (Part L: Membrane Biogenesis: Processing and Recycling) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 99. Hormone Action (Part F: Protein Kinases) Edited by JACKIE D. CORBIN AND JOEL G. HARDMAN VOLUME 100. Recombinant DNA (Part B) Edited by RAY WU, LAWRENCE GROSSMAN, AND KIVIE MOLDAVE VOLUME 101. Recombinant DNA (Part C) Edited by RAY WU, LAWRENCE GROSSMAN, AND KIVIE MOLDAVE VOLUME 102. Hormone Action (Part G: Calmodulin and Calcium-Binding Proteins) Edited by ANTHONY R. MEANS AND BERT W. O’MALLEY VOLUME 103. Hormone Action (Part H: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 104. Enzyme Purification and Related Techniques (Part C) Edited by WILLIAM B. JAKOBY

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VOLUME 105. Oxygen Radicals in Biological Systems Edited by LESTER PACKER VOLUME 106. Posttranslational Modifications (Part A) Edited by FINN WOLD AND KIVIE MOLDAVE VOLUME 107. Posttranslational Modifications (Part B) Edited by FINN WOLD AND KIVIE MOLDAVE VOLUME 108. Immunochemical Techniques (Part G: Separation and Characterization of Lymphoid Cells) Edited by GIOVANNI DI SABATO, JOHN J. LANGONE, AND HELEN VAN VUNAKIS VOLUME 109. Hormone Action (Part I: Peptide Hormones) Edited by LUTZ BIRNBAUMER AND BERT W. O’MALLEY VOLUME 110. Steroids and Isoprenoids (Part A) Edited by JOHN H. LAW AND HANS C. RILLING VOLUME 111. Steroids and Isoprenoids (Part B) Edited by JOHN H. LAW AND HANS C. RILLING VOLUME 112. Drug and Enzyme Targeting (Part A) Edited by KENNETH J. WIDDER AND RALPH GREEN VOLUME 113. Glutamate, Glutamine, Glutathione, and Related Compounds Edited by ALTON MEISTER VOLUME 114. Diffraction Methods for Biological Macromolecules (Part A) Edited by HAROLD W. WYCKOFF, C. H. W. HIRS, AND SERGE N. TIMASHEFF VOLUME 115. Diffraction Methods for Biological Macromolecules (Part B) Edited by HAROLD W. WYCKOFF, C. H. W. HIRS, AND SERGE N. TIMASHEFF VOLUME 116. Immunochemical Techniques (Part H: Effectors and Mediators of Lymphoid Cell Functions) Edited by GIOVANNI DI SABATO, JOHN J. LANGONE, AND HELEN VAN VUNAKIS VOLUME 117. Enzyme Structure (Part J) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 118. Plant Molecular Biology Edited by ARTHUR WEISSBACH AND HERBERT WEISSBACH VOLUME 119. Interferons (Part C) Edited by SIDNEY PESTKA VOLUME 120. Cumulative Subject Index Volumes 81–94, 96–101 VOLUME 121. Immunochemical Techniques (Part I: Hybridoma Technology and Monoclonal Antibodies) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 122. Vitamins and Coenzymes (Part G) Edited by FRANK CHYTIL AND DONALD B. MCCORMICK

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VOLUME 123. Vitamins and Coenzymes (Part H) Edited by FRANK CHYTIL AND DONALD B. MCCORMICK VOLUME 124. Hormone Action (Part J: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 125. Biomembranes (Part M: Transport in Bacteria, Mitochondria, and Chloroplasts: General Approaches and Transport Systems) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 126. Biomembranes (Part N: Transport in Bacteria, Mitochondria, and Chloroplasts: Protonmotive Force) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 127. Biomembranes (Part O: Protons and Water: Structure and Translocation) Edited by LESTER PACKER VOLUME 128. Plasma Lipoproteins (Part A: Preparation, Structure, and Molecular Biology) Edited by JERE P. SEGREST AND JOHN J. ALBERS VOLUME 129. Plasma Lipoproteins (Part B: Characterization, Cell Biology, and Metabolism) Edited by JOHN J. ALBERS AND JERE P. SEGREST VOLUME 130. Enzyme Structure (Part K) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 131. Enzyme Structure (Part L) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 132. Immunochemical Techniques (Part J: Phagocytosis and Cell-Mediated Cytotoxicity) Edited by GIOVANNI DI SABATO AND JOHANNES EVERSE VOLUME 133. Bioluminescence and Chemiluminescence (Part B) Edited by MARLENE DELUCA AND WILLIAM D. MCELROY VOLUME 134. Structural and Contractile Proteins (Part C: The Contractile Apparatus and the Cytoskeleton) Edited by RICHARD B. VALLEE VOLUME 135. Immobilized Enzymes and Cells (Part B) Edited by KLAUS MOSBACH VOLUME 136. Immobilized Enzymes and Cells (Part C) Edited by KLAUS MOSBACH VOLUME 137. Immobilized Enzymes and Cells (Part D) Edited by KLAUS MOSBACH VOLUME 138. Complex Carbohydrates (Part E) Edited by VICTOR GINSBURG

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VOLUME 139. Cellular Regulators (Part A: Calcium- and Calmodulin-Binding Proteins) Edited by ANTHONY R. MEANS AND P. MICHAEL CONN VOLUME 140. Cumulative Subject Index Volumes 102–119, 121–134 VOLUME 141. Cellular Regulators (Part B: Calcium and Lipids) Edited by P. MICHAEL CONN AND ANTHONY R. MEANS VOLUME 142. Metabolism of Aromatic Amino Acids and Amines Edited by SEYMOUR KAUFMAN VOLUME 143. Sulfur and Sulfur Amino Acids Edited by WILLIAM B. JAKOBY AND OWEN GRIFFITH VOLUME 144. Structural and Contractile Proteins (Part D: Extracellular Matrix) Edited by LEON W. CUNNINGHAM VOLUME 145. Structural and Contractile Proteins (Part E: Extracellular Matrix) Edited by LEON W. CUNNINGHAM VOLUME 146. Peptide Growth Factors (Part A) Edited by DAVID BARNES AND DAVID A. SIRBASKU VOLUME 147. Peptide Growth Factors (Part B) Edited by DAVID BARNES AND DAVID A. SIRBASKU VOLUME 148. Plant Cell Membranes Edited by LESTER PACKER AND ROLAND DOUCE VOLUME 149. Drug and Enzyme Targeting (Part B) Edited by RALPH GREEN AND KENNETH J. WIDDER VOLUME 150. Immunochemical Techniques (Part K: In Vitro Models of B and T Cell Functions and Lymphoid Cell Receptors) Edited by GIOVANNI DI SABATO VOLUME 151. Molecular Genetics of Mammalian Cells Edited by MICHAEL M. GOTTESMAN VOLUME 152. Guide to Molecular Cloning Techniques Edited by SHELBY L. BERGER AND ALAN R. KIMMEL VOLUME 153. Recombinant DNA (Part D) Edited by RAY WU AND LAWRENCE GROSSMAN VOLUME 154. Recombinant DNA (Part E) Edited by RAY WU AND LAWRENCE GROSSMAN VOLUME 155. Recombinant DNA (Part F) Edited by RAY WU VOLUME 156. Biomembranes (Part P: ATP-Driven Pumps and Related Transport: The Na, K-Pump) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER

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VOLUME 157. Biomembranes (Part Q: ATP-Driven Pumps and Related Transport: Calcium, Proton, and Potassium Pumps) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 158. Metalloproteins (Part A) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 159. Initiation and Termination of Cyclic Nucleotide Action Edited by JACKIE D. CORBIN AND ROGER A. JOHNSON VOLUME 160. Biomass (Part A: Cellulose and Hemicellulose) Edited by WILLIS A. WOOD AND SCOTT T. KELLOGG VOLUME 161. Biomass (Part B: Lignin, Pectin, and Chitin) Edited by WILLIS A. WOOD AND SCOTT T. KELLOGG VOLUME 162. Immunochemical Techniques (Part L: Chemotaxis and Inflammation) Edited by GIOVANNI DI SABATO VOLUME 163. Immunochemical Techniques (Part M: Chemotaxis and Inflammation) Edited by GIOVANNI DI SABATO VOLUME 164. Ribosomes Edited by HARRY F. NOLLER, JR., AND KIVIE MOLDAVE VOLUME 165. Microbial Toxins: Tools for Enzymology Edited by SIDNEY HARSHMAN VOLUME 166. Branched-Chain Amino Acids Edited by ROBERT HARRIS AND JOHN R. SOKATCH VOLUME 167. Cyanobacteria Edited by LESTER PACKER AND ALEXANDER N. GLAZER VOLUME 168. Hormone Action (Part K: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 169. Platelets: Receptors, Adhesion, Secretion (Part A) Edited by JACEK HAWIGER VOLUME 170. Nucleosomes Edited by PAUL M. WASSARMAN AND ROGER D. KORNBERG VOLUME 171. Biomembranes (Part R: Transport Theory: Cells and Model Membranes) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 172. Biomembranes (Part S: Transport: Membrane Isolation and Characterization) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER

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VOLUME 173. Biomembranes [Part T: Cellular and Subcellular Transport: Eukaryotic (Nonepithelial) Cells] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 174. Biomembranes [Part U: Cellular and Subcellular Transport: Eukaryotic (Nonepithelial) Cells] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 175. Cumulative Subject Index Volumes 135–139, 141–167 VOLUME 176. Nuclear Magnetic Resonance (Part A: Spectral Techniques and Dynamics) Edited by NORMAN J. OPPENHEIMER AND THOMAS L. JAMES VOLUME 177. Nuclear Magnetic Resonance (Part B: Structure and Mechanism) Edited by NORMAN J. OPPENHEIMER AND THOMAS L. JAMES VOLUME 178. Antibodies, Antigens, and Molecular Mimicry Edited by JOHN J. LANGONE VOLUME 179. Complex Carbohydrates (Part F) Edited by VICTOR GINSBURG VOLUME 180. RNA Processing (Part A: General Methods) Edited by JAMES E. DAHLBERG AND JOHN N. ABELSON VOLUME 181. RNA Processing (Part B: Specific Methods) Edited by JAMES E. DAHLBERG AND JOHN N. ABELSON VOLUME 182. Guide to Protein Purification Edited by MURRAY P. DEUTSCHER VOLUME 183. Molecular Evolution: Computer Analysis of Protein and Nucleic Acid Sequences Edited by RUSSELL F. DOOLITTLE VOLUME 184. Avidin-Biotin Technology Edited by MEIR WILCHEK AND EDWARD A. BAYER VOLUME 185. Gene Expression Technology Edited by DAVID V. GOEDDEL VOLUME 186. Oxygen Radicals in Biological Systems (Part B: Oxygen Radicals and Antioxidants) Edited by LESTER PACKER AND ALEXANDER N. GLAZER VOLUME 187. Arachidonate Related Lipid Mediators Edited by ROBERT C. MURPHY AND FRANK A. FITZPATRICK VOLUME 188. Hydrocarbons and Methylotrophy Edited by MARY E. LIDSTROM VOLUME 189. Retinoids (Part A: Molecular and Metabolic Aspects) Edited by LESTER PACKER

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VOLUME 190. Retinoids (Part B: Cell Differentiation and Clinical Applications) Edited by LESTER PACKER VOLUME 191. Biomembranes (Part V: Cellular and Subcellular Transport: Epithelial Cells) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 192. Biomembranes (Part W: Cellular and Subcellular Transport: Epithelial Cells) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 193. Mass Spectrometry Edited by JAMES A. MCCLOSKEY VOLUME 194. Guide to Yeast Genetics and Molecular Biology Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 195. Adenylyl Cyclase, G Proteins, and Guanylyl Cyclase Edited by ROGER A. JOHNSON AND JACKIE D. CORBIN VOLUME 196. Molecular Motors and the Cytoskeleton Edited by RICHARD B. VALLEE VOLUME 197. Phospholipases Edited by EDWARD A. DENNIS VOLUME 198. Peptide Growth Factors (Part C) Edited by DAVID BARNES, J. P. MATHER, AND GORDON H. SATO VOLUME 199. Cumulative Subject Index Volumes 168–174, 176–194 VOLUME 200. Protein Phosphorylation (Part A: Protein Kinases: Assays, Purification, Antibodies, Functional Analysis, Cloning, and Expression) Edited by TONY HUNTER AND BARTHOLOMEW M. SEFTON VOLUME 201. Protein Phosphorylation (Part B: Analysis of Protein Phosphorylation, Protein Kinase Inhibitors, and Protein Phosphatases) Edited by TONY HUNTER AND BARTHOLOMEW M. SEFTON VOLUME 202. Molecular Design and Modeling: Concepts and Applications (Part A: Proteins, Peptides, and Enzymes) Edited by JOHN J. LANGONE VOLUME 203. Molecular Design and Modeling: Concepts and Applications (Part B: Antibodies and Antigens, Nucleic Acids, Polysaccharides, and Drugs) Edited by JOHN J. LANGONE VOLUME 204. Bacterial Genetic Systems Edited by JEFFREY H. MILLER VOLUME 205. Metallobiochemistry (Part B: Metallothionein and Related Molecules) Edited by JAMES F. RIORDAN AND BERT L. VALLEE

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VOLUME 206. Cytochrome P450 Edited by MICHAEL R. WATERMAN AND ERIC F. JOHNSON VOLUME 207. Ion Channels Edited by BERNARDO RUDY AND LINDA E. IVERSON VOLUME 208. Protein–DNA Interactions Edited by ROBERT T. SAUER VOLUME 209. Phospholipid Biosynthesis Edited by EDWARD A. DENNIS AND DENNIS E. VANCE VOLUME 210. Numerical Computer Methods Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 211. DNA Structures (Part A: Synthesis and Physical Analysis of DNA) Edited by DAVID M. J. LILLEY AND JAMES E. DAHLBERG VOLUME 212. DNA Structures (Part B: Chemical and Electrophoretic Analysis of DNA) Edited by DAVID M. J. LILLEY AND JAMES E. DAHLBERG VOLUME 213. Carotenoids (Part A: Chemistry, Separation, Quantitation, and Antioxidation) Edited by LESTER PACKER VOLUME 214. Carotenoids (Part B: Metabolism, Genetics, and Biosynthesis) Edited by LESTER PACKER VOLUME 215. Platelets: Receptors, Adhesion, Secretion (Part B) Edited by JACEK J. HAWIGER VOLUME 216. Recombinant DNA (Part G) Edited by RAY WU VOLUME 217. Recombinant DNA (Part H) Edited by RAY WU VOLUME 218. Recombinant DNA (Part I) Edited by RAY WU VOLUME 219. Reconstitution of Intracellular Transport Edited by JAMES E. ROTHMAN VOLUME 220. Membrane Fusion Techniques (Part A) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 221. Membrane Fusion Techniques (Part B) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 222. Proteolytic Enzymes in Coagulation, Fibrinolysis, and Complement Activation (Part A: Mammalian Blood Coagulation Factors and Inhibitors) Edited by LASZLO LORAND AND KENNETH G. MANN

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VOLUME 223. Proteolytic Enzymes in Coagulation, Fibrinolysis, and Complement Activation (Part B: Complement Activation, Fibrinolysis, and Nonmammalian Blood Coagulation Factors) Edited by LASZLO LORAND AND KENNETH G. MANN VOLUME 224. Molecular Evolution: Producing the Biochemical Data Edited by ELIZABETH ANNE ZIMMER, THOMAS J. WHITE, REBECCA L. CANN, AND ALLAN C. WILSON VOLUME 225. Guide to Techniques in Mouse Development Edited by PAUL M. WASSARMAN AND MELVIN L. DEPAMPHILIS VOLUME 226. Metallobiochemistry (Part C: Spectroscopic and Physical Methods for Probing Metal Ion Environments in Metalloenzymes and Metalloproteins) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 227. Metallobiochemistry (Part D: Physical and Spectroscopic Methods for Probing Metal Ion Environments in Metalloproteins) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 228. Aqueous Two-Phase Systems Edited by HARRY WALTER AND GO¨TE JOHANSSON VOLUME 229. Cumulative Subject Index Volumes 195–198, 200–227 VOLUME 230. Guide to Techniques in Glycobiology Edited by WILLIAM J. LENNARZ AND GERALD W. HART VOLUME 231. Hemoglobins (Part B: Biochemical and Analytical Methods) Edited by JOHANNES EVERSE, KIM D. VANDEGRIFF, AND ROBERT M. WINSLOW VOLUME 232. Hemoglobins (Part C: Biophysical Methods) Edited by JOHANNES EVERSE, KIM D. VANDEGRIFF, AND ROBERT M. WINSLOW VOLUME 233. Oxygen Radicals in Biological Systems (Part C) Edited by LESTER PACKER VOLUME 234. Oxygen Radicals in Biological Systems (Part D) Edited by LESTER PACKER VOLUME 235. Bacterial Pathogenesis (Part A: Identification and Regulation of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 236. Bacterial Pathogenesis (Part B: Integration of Pathogenic Bacteria with Host Cells) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 237. Heterotrimeric G Proteins Edited by RAVI IYENGAR VOLUME 238. Heterotrimeric G-Protein Effectors Edited by RAVI IYENGAR

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VOLUME 239. Nuclear Magnetic Resonance (Part C) Edited by THOMAS L. JAMES AND NORMAN J. OPPENHEIMER VOLUME 240. Numerical Computer Methods (Part B) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 241. Retroviral Proteases Edited by LAWRENCE C. KUO AND JULES A. SHAFER VOLUME 242. Neoglycoconjugates (Part A) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 243. Inorganic Microbial Sulfur Metabolism Edited by HARRY D. PECK, JR., AND JEAN LEGALL VOLUME 244. Proteolytic Enzymes: Serine and Cysteine Peptidases Edited by ALAN J. BARRETT VOLUME 245. Extracellular Matrix Components Edited by E. RUOSLAHTI AND E. ENGVALL VOLUME 246. Biochemical Spectroscopy Edited by KENNETH SAUER VOLUME 247. Neoglycoconjugates (Part B: Biomedical Applications) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 248. Proteolytic Enzymes: Aspartic and Metallo Peptidases Edited by ALAN J. BARRETT VOLUME 249. Enzyme Kinetics and Mechanism (Part D: Developments in Enzyme Dynamics) Edited by DANIEL L. PURICH VOLUME 250. Lipid Modifications of Proteins Edited by PATRICK J. CASEY AND JANICE E. BUSS VOLUME 251. Biothiols (Part A: Monothiols and Dithiols, Protein Thiols, and Thiyl Radicals) Edited by LESTER PACKER VOLUME 252. Biothiols (Part B: Glutathione and Thioredoxin; Thiols in Signal Transduction and Gene Regulation) Edited by LESTER PACKER VOLUME 253. Adhesion of Microbial Pathogens Edited by RON J. DOYLE AND ITZHAK OFEK VOLUME 254. Oncogene Techniques Edited by PETER K. VOGT AND INDER M. VERMA VOLUME 255. Small GTPases and Their Regulators (Part A: Ras Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL

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VOLUME 256. Small GTPases and Their Regulators (Part B: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 257. Small GTPases and Their Regulators (Part C: Proteins Involved in Transport) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 258. Redox-Active Amino Acids in Biology Edited by JUDITH P. KLINMAN VOLUME 259. Energetics of Biological Macromolecules Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 260. Mitochondrial Biogenesis and Genetics (Part A) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 261. Nuclear Magnetic Resonance and Nucleic Acids Edited by THOMAS L. JAMES VOLUME 262. DNA Replication Edited by JUDITH L. CAMPBELL VOLUME 263. Plasma Lipoproteins (Part C: Quantitation) Edited by WILLIAM A. BRADLEY, SANDRA H. GIANTURCO, AND JERE P. SEGREST VOLUME 264. Mitochondrial Biogenesis and Genetics (Part B) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 265. Cumulative Subject Index Volumes 228, 230–262 VOLUME 266. Computer Methods for Macromolecular Sequence Analysis Edited by RUSSELL F. DOOLITTLE VOLUME 267. Combinatorial Chemistry Edited by JOHN N. ABELSON VOLUME 268. Nitric Oxide (Part A: Sources and Detection of NO; NO Synthase) Edited by LESTER PACKER VOLUME 269. Nitric Oxide (Part B: Physiological and Pathological Processes) Edited by LESTER PACKER VOLUME 270. High Resolution Separation and Analysis of Biological Macromolecules (Part A: Fundamentals) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 271. High Resolution Separation and Analysis of Biological Macromolecules (Part B: Applications) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 272. Cytochrome P450 (Part B) Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 273. RNA Polymerase and Associated Factors (Part A) Edited by SANKAR ADHYA

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VOLUME 274. RNA Polymerase and Associated Factors (Part B) Edited by SANKAR ADHYA VOLUME 275. Viral Polymerases and Related Proteins Edited by LAWRENCE C. KUO, DAVID B. OLSEN, AND STEVEN S. CARROLL VOLUME 276. Macromolecular Crystallography (Part A) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 277. Macromolecular Crystallography (Part B) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 278. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 279. Vitamins and Coenzymes (Part I) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 280. Vitamins and Coenzymes (Part J) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 281. Vitamins and Coenzymes (Part K) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 282. Vitamins and Coenzymes (Part L) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 283. Cell Cycle Control Edited by WILLIAM G. DUNPHY VOLUME 284. Lipases (Part A: Biotechnology) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 285. Cumulative Subject Index Volumes 263, 264, 266–284, 286–289 VOLUME 286. Lipases (Part B: Enzyme Characterization and Utilization) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 287. Chemokines Edited by RICHARD HORUK VOLUME 288. Chemokine Receptors Edited by RICHARD HORUK VOLUME 289. Solid Phase Peptide Synthesis Edited by GREGG B. FIELDS VOLUME 290. Molecular Chaperones Edited by GEORGE H. LORIMER AND THOMAS BALDWIN VOLUME 291. Caged Compounds Edited by GERARD MARRIOTT VOLUME 292. ABC Transporters: Biochemical, Cellular, and Molecular Aspects Edited by SURESH V. AMBUDKAR AND MICHAEL M. GOTTESMAN

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VOLUME 293. Ion Channels (Part B) Edited by P. MICHAEL CONN VOLUME 294. Ion Channels (Part C) Edited by P. MICHAEL CONN VOLUME 295. Energetics of Biological Macromolecules (Part B) Edited by GARY K. ACKERS AND MICHAEL L. JOHNSON VOLUME 296. Neurotransmitter Transporters Edited by SUSAN G. AMARA VOLUME 297. Photosynthesis: Molecular Biology of Energy Capture Edited by LEE MCINTOSH VOLUME 298. Molecular Motors and the Cytoskeleton (Part B) Edited by RICHARD B. VALLEE VOLUME 299. Oxidants and Antioxidants (Part A) Edited by LESTER PACKER VOLUME 300. Oxidants and Antioxidants (Part B) Edited by LESTER PACKER VOLUME 301. Nitric Oxide: Biological and Antioxidant Activities (Part C) Edited by LESTER PACKER VOLUME 302. Green Fluorescent Protein Edited by P. MICHAEL CONN VOLUME 303. cDNA Preparation and Display Edited by SHERMAN M. WEISSMAN VOLUME 304. Chromatin Edited by PAUL M. WASSARMAN AND ALAN P. WOLFFE VOLUME 305. Bioluminescence and Chemiluminescence (Part C) Edited by THOMAS O. BALDWIN AND MIRIAM M. ZIEGLER VOLUME 306. Expression of Recombinant Genes in Eukaryotic Systems Edited by JOSEPH C. GLORIOSO AND MARTIN C. SCHMIDT VOLUME 307. Confocal Microscopy Edited by P. MICHAEL CONN VOLUME 308. Enzyme Kinetics and Mechanism (Part E: Energetics of Enzyme Catalysis) Edited by DANIEL L. PURICH AND VERN L. SCHRAMM VOLUME 309. Amyloid, Prions, and Other Protein Aggregates Edited by RONALD WETZEL VOLUME 310. Biofilms Edited by RON J. DOYLE

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VOLUME 311. Sphingolipid Metabolism and Cell Signaling (Part A) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN VOLUME 312. Sphingolipid Metabolism and Cell Signaling (Part B) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN VOLUME 313. Antisense Technology (Part A: General Methods, Methods of Delivery, and RNA Studies) Edited by M. IAN PHILLIPS VOLUME 314. Antisense Technology (Part B: Applications) Edited by M. IAN PHILLIPS VOLUME 315. Vertebrate Phototransduction and the Visual Cycle (Part A) Edited by KRZYSZTOF PALCZEWSKI VOLUME 316. Vertebrate Phototransduction and the Visual Cycle (Part B) Edited by KRZYSZTOF PALCZEWSKI VOLUME 317. RNA–Ligand Interactions (Part A: Structural Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 318. RNA–Ligand Interactions (Part B: Molecular Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 319. Singlet Oxygen, UV-A, and Ozone Edited by LESTER PACKER AND HELMUT SIES VOLUME 320. Cumulative Subject Index Volumes 290–319 VOLUME 321. Numerical Computer Methods (Part C) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 322. Apoptosis Edited by JOHN C. REED VOLUME 323. Energetics of Biological Macromolecules (Part C) Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 324. Branched-Chain Amino Acids (Part B) Edited by ROBERT A. HARRIS AND JOHN R. SOKATCH VOLUME 325. Regulators and Effectors of Small GTPases (Part D: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 326. Applications of Chimeric Genes and Hybrid Proteins (Part A: Gene Expression and Protein Purification) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 327. Applications of Chimeric Genes and Hybrid Proteins (Part B: Cell Biology and Physiology) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON

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VOLUME 328. Applications of Chimeric Genes and Hybrid Proteins (Part C: Protein–Protein Interactions and Genomics) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 329. Regulators and Effectors of Small GTPases (Part E: GTPases Involved in Vesicular Traffic) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 330. Hyperthermophilic Enzymes (Part A) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 331. Hyperthermophilic Enzymes (Part B) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 332. Regulators and Effectors of Small GTPases (Part F: Ras Family I) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 333. Regulators and Effectors of Small GTPases (Part G: Ras Family II) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 334. Hyperthermophilic Enzymes (Part C) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 335. Flavonoids and Other Polyphenols Edited by LESTER PACKER VOLUME 336. Microbial Growth in Biofilms (Part A: Developmental and Molecular Biological Aspects) Edited by RON J. DOYLE VOLUME 337. Microbial Growth in Biofilms (Part B: Special Environments and Physicochemical Aspects) Edited by RON J. DOYLE VOLUME 338. Nuclear Magnetic Resonance of Biological Macromolecules (Part A) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 339. Nuclear Magnetic Resonance of Biological Macromolecules (Part B) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 340. Drug–Nucleic Acid Interactions Edited by JONATHAN B. CHAIRES AND MICHAEL J. WARING VOLUME 341. Ribonucleases (Part A) Edited by ALLEN W. NICHOLSON VOLUME 342. Ribonucleases (Part B) Edited by ALLEN W. NICHOLSON VOLUME 343. G Protein Pathways (Part A: Receptors) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 344. G Protein Pathways (Part B: G Proteins and Their Regulators) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT

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VOLUME 345. G Protein Pathways (Part C: Effector Mechanisms) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 346. Gene Therapy Methods Edited by M. IAN PHILLIPS VOLUME 347. Protein Sensors and Reactive Oxygen Species (Part A: Selenoproteins and Thioredoxin) Edited by HELMUT SIES AND LESTER PACKER VOLUME 348. Protein Sensors and Reactive Oxygen Species (Part B: Thiol Enzymes and Proteins) Edited by HELMUT SIES AND LESTER PACKER VOLUME 349. Superoxide Dismutase Edited by LESTER PACKER VOLUME 350. Guide to Yeast Genetics and Molecular and Cell Biology (Part B) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 351. Guide to Yeast Genetics and Molecular and Cell Biology (Part C) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 352. Redox Cell Biology and Genetics (Part A) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 353. Redox Cell Biology and Genetics (Part B) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 354. Enzyme Kinetics and Mechanisms (Part F: Detection and Characterization of Enzyme Reaction Intermediates) Edited by DANIEL L. PURICH VOLUME 355. Cumulative Subject Index Volumes 321–354 VOLUME 356. Laser Capture Microscopy and Microdissection Edited by P. MICHAEL CONN VOLUME 357. Cytochrome P450, Part C Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 358. Bacterial Pathogenesis (Part C: Identification, Regulation, and Function of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 359. Nitric Oxide (Part D) Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 360. Biophotonics (Part A) Edited by GERARD MARRIOTT AND IAN PARKER VOLUME 361. Biophotonics (Part B) Edited by GERARD MARRIOTT AND IAN PARKER

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VOLUME 362. Recognition of Carbohydrates in Biological Systems (Part A) Edited by YUAN C. LEE AND REIKO T. LEE VOLUME 363. Recognition of Carbohydrates in Biological Systems (Part B) Edited by YUAN C. LEE AND REIKO T. LEE VOLUME 364. Nuclear Receptors Edited by DAVID W. RUSSELL AND DAVID J. MANGELSDORF VOLUME 365. Differentiation of Embryonic Stem Cells Edited by PAUL M. WASSAUMAN AND GORDON M. KELLER VOLUME 366. Protein Phosphatases Edited by SUSANNE KLUMPP AND JOSEF KRIEGLSTEIN VOLUME 367. Liposomes (Part A) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 368. Macromolecular Crystallography (Part C) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 369. Combinational Chemistry (Part B) Edited by GUILLERMO A. MORALES AND BARRY A. BUNIN VOLUME 370. RNA Polymerases and Associated Factors (Part C) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 371. RNA Polymerases and Associated Factors (Part D) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 372. Liposomes (Part B) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 373. Liposomes (Part C) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 374. Macromolecular Crystallography (Part D) Edited by CHARLES W. CARTER, JR., AND ROBERT W. SWEET VOLUME 375. Chromatin and Chromatin Remodeling Enzymes (Part A) Edited by C. DAVID ALLIS AND CARL WU VOLUME 376. Chromatin and Chromatin Remodeling Enzymes (Part B) Edited by C. DAVID ALLIS AND CARL WU VOLUME 377. Chromatin and Chromatin Remodeling Enzymes (Part C) Edited by C. DAVID ALLIS AND CARL WU VOLUME 378. Quinones and Quinone Enzymes (Part A) Edited by HELMUT SIES AND LESTER PACKER VOLUME 379. Energetics of Biological Macromolecules (Part D) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS VOLUME 380. Energetics of Biological Macromolecules (Part E) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS

Methods in Enzymology

VOLUME 381. Oxygen Sensing Edited by CHANDAN K. SEN AND GREGG L. SEMENZA VOLUME 382. Quinones and Quinone Enzymes (Part B) Edited by HELMUT SIES AND LESTER PACKER VOLUME 383. Numerical Computer Methods (Part D) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 384. Numerical Computer Methods (Part E) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 385. Imaging in Biological Research (Part A) Edited by P. MICHAEL CONN VOLUME 386. Imaging in Biological Research (Part B) Edited by P. MICHAEL CONN VOLUME 387. Liposomes (Part D) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 388. Protein Engineering Edited by DAN E. ROBERTSON AND JOSEPH P. NOEL VOLUME 389. Regulators of G-Protein Signaling (Part A) Edited by DAVID P. SIDEROVSKI VOLUME 390. Regulators of G-Protein Signaling (Part B) Edited by DAVID P. SIDEROVSKI VOLUME 391. Liposomes (Part E) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 392. RNA Interference Edited by ENGELKE ROSSI VOLUME 393. Circadian Rhythms Edited by MICHAEL W. YOUNG VOLUME 394. Nuclear Magnetic Resonance of Biological Macromolecules (Part C) Edited by THOMAS L. JAMES VOLUME 395. Producing the Biochemical Data (Part B) Edited by ELIZABETH A. ZIMMER AND ERIC H. ROALSON VOLUME 396. Nitric Oxide (Part E) Edited by LESTER PACKER AND ENRIQUE CADENAS VOLUME 397. Environmental Microbiology Edited by JARED R. LEADBETTER VOLUME 398. Ubiquitin and Protein Degradation (Part A) Edited by RAYMOND J. DESHAIES

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VOLUME 399. Ubiquitin and Protein Degradation (Part B) Edited by RAYMOND J. DESHAIES VOLUME 400. Phase II Conjugation Enzymes and Transport Systems Edited by HELMUT SIES AND LESTER PACKER VOLUME 401. Glutathione Transferases and Gamma Glutamyl Transpeptidases Edited by HELMUT SIES AND LESTER PACKER VOLUME 402. Biological Mass Spectrometry Edited by A. L. BURLINGAME VOLUME 403. GTPases Regulating Membrane Targeting and Fusion Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 404. GTPases Regulating Membrane Dynamics Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 405. Mass Spectrometry: Modified Proteins and Glycoconjugates Edited by A. L. BURLINGAME VOLUME 406. Regulators and Effectors of Small GTPases: Rho Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 407. Regulators and Effectors of Small GTPases: Ras Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 408. DNA Repair (Part A) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 409. DNA Repair (Part B) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 410. DNA Microarrays (Part A: Array Platforms and Web-Bench Protocols) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 411. DNA Microarrays (Part B: Databases and Statistics) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 412. Amyloid, Prions, and Other Protein Aggregates (Part B) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 413. Amyloid, Prions, and Other Protein Aggregates (Part C) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 414. Measuring Biological Responses with Automated Microscopy Edited by JAMES INGLESE VOLUME 415. Glycobiology Edited by MINORU FUKUDA VOLUME 416. Glycomics Edited by MINORU FUKUDA

Methods in Enzymology

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Characterization of the Recombinant PyrophosphateDependent 6-Phosphofructokinases from Methylomicrobium alcaliphilum 20Z and Methylococcus capsulatus Bath Valentina N. Khmelenina, Olga N. Rozova, and Yuri A. Trotsenko Contents 1. Introduction 2. Methods 2.1. Culture conditions 2.2. Routine genetic manipulations 2.3. PPi-PFKs cloning, expression, and purification 2.4. Assay of pyrophosphate-dependent 6-phosphofructokinase activity 2.5. Assay of fructosebisphosphate aldolase activity by using the recombinant PPi-PFK 3. Brief Overview of Current Knowledge of PPi-PFK in Microorganisms 4. Properties of PPi-PFK from Methanotrophs 4.1. PPi-PFK from Mc. capsulatus Bath 4.2. PPi-PFK from Mm. alcaliphilum 20Z 5. Conclusion Acknowledgments References

2 4 4 4 4 4 6 7 8 8 10 10 11 11

Abstract The Embden–Meyerhof–Parnas (EMP) glycolysis is the starting point of the core carbon metabolism. Aerobic methanotrophs possessing activity of the pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK) instead of the classical glycolytic enzyme ATP-dependent 6-phosphofructokinase (ATP-PFK) are Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00001-2

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2011 Elsevier Inc. All rights reserved.

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promising model bacteria for elucidation of the role of inorganic pyrophosphate (PPi) and PPi-dependent glycolysis in microorganisms. Characterization of the His6-tagged PPi-PFKs from two methanotrophs, halotolerant alkaliphilic Methylomicrobium alcaliphilum 20Z and thermotolerant Methylococcus capsulatus Bath, showed differential capabilities of PPi-PFKs to phosphorylate sedoheptulose-7phosphate and this property correlated well with the metabolic patterns of these bacteria assimilating C1 substrate either via the ribulosemonophosphate (RuMP) pathway (Mm. alcaliphilum 20Z) or simultaneously via the RuMP and serine pathways and the Calvin cycle (Mc. capsulatus Bath). Analysis of the genomic draft of Mm. alcaliphilum 20Z (https://www.genoscope.cns.fr/agc/mage) has provided in silico evidence for the existence of a PPi-dependent pyruvate-phosphate dikinase (PPDK). Expression of the ppdk gene at oxygen limitation along with the presence of PPi-PFK in Mm. alcaliphilum 20Z implied functioning of PPi-dependent glycolysis and PPi recycling under conditions when oxidative phosphorylation is hampered.

1. Introduction Energy metabolism of many organisms is based on glycolysis. In aerobic microorganisms, glycolysis is a minor source of energy, since ca. 95% of ATP production comes from subsequent tricarboxylic acid cycle reactions and oxidative phosphorylation. In contrast, anaerobic organisms rely almost exclusively on glycolysis and fermentation for ATP production (Mertens, 1991; Mertens et al., 1993). One significant variation of the standard glycolytic pathway is pyrophosphate-dependent glycolysis, which uses PPi instead of ATP as a phosphoryl donor (Mertens, 1991; Saavedra et al., 2005). In this version of glycolysis, two key glycolytic enzymes, ATP-PFK and ADP-pyruvate kinase (PK), are replaced by pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK) and pyruvate phosphate dikinase (PPDK) the latter catalyzing the reversible reaction of AMP- and PPi-dependent conversion of phosphoenolpyruvate (PEP) into pyruvate accompanied by ATP synthesis (Mertens, 1991; Mertens et al., 1993). PPi-dependent glycolysis can in theory yield five ATP molecules instead of the two yielded by standard glycolysis. This increase of 2.5-fold can be crucial under severe energy-limiting conditions. This modified glycolysis has been well studied in anaerobic bacteria and protists (Saavedra et al., 2005). Apart from the widely distributed ATP-PFK catalyzing an irreversible catabolic reaction, the phosphorylation of fructose-6-phosphate (Fru 6-P) to fructose-1,6-bisphosphate (Fru 1,6-P2), PPi-PFK (EC 2.7.1.90) catalyzes the same reaction in reversible way and can thus function both in glycolysis and gluconeogenesis (Reeves et al., 1974, 1976).

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Mg2þ

Fructose-6-phosphate þ PPi > Fructose-1; 6-bisphosphate þ Pi

PPi-PFK activity was detected in several protozoans, in higher plants and in prokaryotes (Bruchhaus et al., 1996; Carnal and Black, 1983; Mertens et al., 1993; O’Brien et al., 1975; Petzel et al., 1989). In many organisms, PPi-PFK replaces allosteric ATP-PFK which catalyses an essentially irreversible catabolic reaction of classical glycolysis (Slamovits and Keeling, 2006; Wood et al., 1977). In some cases, PPi-PFK is present alongside ATP-PFK in the same organism, suggesting that PPi-PFK may perform an unknown specific function (Alves et al., 1994, 2001; Van Praag, 1997). Activity of PPi-PFK was found in 10 strains of types I, II, and X aerobic methanotrophic bacteria (Beschastny et al., 1992, 2008; Trotsenko and Shishkina, 1990). Also, intracellular PPi concentrations 20 times higher than in Escherichia coli cells and only low ATP concentration were found in the methanotrophs studied and such metabolic features correlated with extremely low inorganic pyrophosphatase activity (Beschastny et al., 2008; Chetina and Trotsenko, 1987; Trotsenko and Shishkina, 1990). The role of PPi and PPi-dependent enzymes in methanotrophs is not obvious. It is generally accepted that aerobic methanotrophs oxidize methane to carbon dioxide and water via the intermediates of methanol, formaldehyde, and formate, thereby providing energy for biosynthesis of cell components (Trotsenko and Murrell, 2008). ATP production is thought to come from oxidative phosphorylation but with low efficiency in methanotrophs studied (Chetina and Trotsenko, 1987). ATP production relying on substratelevel phosphorylation may play a crucial role in these bacteria. Methanotrophs are different in their constructive metabolic pattern. Type I methanotrophic bacteria assimilate C1 compounds via the ribulosemonophosphate (RuMP) cycle of formaldehyde fixation, in which the first intermediates are C6 phosphosugars. Phosphotrioses are formed by cleavage of phosphohexoses through both the Entner–Doudoroff and Embden– Meyerhof–Parnas (EMP) pathways (Shishkina and Trotsenko, 1982; Stro¨m et al., 1974). In contrast, type II methanotrophs use the serine pathway of C1 assimilation, where C3 compounds are firstly formed after condensation of formaldehyde and glycine (Stro¨m et al., 1974; Trotsenko and Murrell, 2008); therefore, C6 phosphosugar synthesis via gluconeogenesis reactions is needed. Notably, simultaneous functioning of three pathways: the RuMP cycle as the major pathway, the serine pathway and the Calvin cycle was detected for the type X methanotrophs Methylococcus capsulatus Bath and Methylocaldum szegediense (Medvedkova et al., 2009; Trotsenko and Murrell, 2008). Operation of the Calvin cycle may be related to the thermophilic nature of these methanotrophic species (Trotsenko et al., 2009). Thus, in type I and type X methanotrophs, both C3 phosphotrioses and hexosephosphates can be formed bypassing the classical EMP

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pathway. This suggests that glycolysis and the PPi-dependent PFK may perform unknown specific function(s). Availability of the genome sequences of several methanotrophs, including halotolerant Mm. alcaliphilum 20Z (https:// www.genoscope.cns.fr/agc/mage) and thermotolerant Mc. capsulatus Bath (Ward et al., 2004), has generated the required template to investigate in detail their metabolic pattern at both genetic and enzymatic levels. In this study, we describe properties of purified PPi-PFKs from Mm. alcaliphilum 20Z and Mc. capsulatus Bath.

2. Methods 2.1. Culture conditions Cells of Mm. alcaliphilum were grown under methane-air atmosphere (1:1) in liquid mineral medium containing 3% NaCl (Khmelenina et al., 1999). The standard NMS medium (Whittenbury et al., 1970) was used for Mc. capsulatus Bath cultivation at 43  C.

2.2. Routine genetic manipulations DNA from Mm. alcaliphilum 20Z and Mc. capsulatus Bath was isolated by a standard phenol:chloroform method (Sambrook and Russell, 2001). For RNA isolation from 10 ml of exponentially grown culture (OD600 ¼ 0.8), the RNA extraction method of Chomczynski and Sacchi (1987) was used as described in Reshetnikov et al. (2006).

2.3. PPi-PFKs cloning, expression, and purification The standard procedures may be used for cloning of the pfp genes into the bacterial expression vector pET22b designed to express a C-terminal His6tagged fusion protein under the control of the T7 promoter. A single step of immobilized metal ion affinity chromatography on a Ni2þ-NTA column was performed for purification of His6-tagged PPi-PFK fusion protein from E. coli extracts (Reshetnikov et al., 2008).

2.4. Assay of pyrophosphate-dependent 6-phosphofructokinase activity 2.4.1. Assay of PPi-PFK with Fru 6-P as phosphoryl acceptor 1 ml of reaction mixture contained: 50 mM HEPES–HCl, pH 7.0; 20 mM Fru 6-P;

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2 mM PPi; 5 mM MgCl2; 0.25 mM NADH; 0.2 U of aldolase (from rabbit muscle); 0.3 U of a-glycerophosphate dehydrogenase; 0.3 U of triosephosphate isomerase (Sigma). After preincubation at 30  C for 3 min, 1–2 mg of PPi-PFK was added and the NADH oxidation rate was measured by the decrease in absorbance at 340 nm. 2.4.2. Assay of PPi-PFK with sedoheptulose-7-phosphate as phosphoryl acceptor 1 ml of reaction mixture contained: 50 mM HEPES–HCl, pH 7.0 5 mM sedoheptulose-7-phosphate; 2 mM PPi; 5 mM MgCl2; 0.25 mM NADH; 0.2 U of aldolase (from rabbit muscle); 0.3 U of a-glycerophosphate dehydrogenase; After preincubation at 30  C for 3 min, 1–2 mg of PPi-PFK was added and the NADH oxidation rate was measured by the decrease in absorbance at 340 nm. 2.4.3. Assay of PPi-PFK with Fru 1,6-P2 1 ml of reaction mixture contained: 50 mM HEPES–HCl, pH 7.0; 2 mM Fru 1,6-P2; 15 mM NaH2PO4; 5 mM MgCl2; 0.3 mM NADPþ; 0.12 U of glucose-6-phosphate dehydrogenase; 0.24 U of glucose phosphate isomerase. Incubate at 30  C for 3 min. Add 1 mg of PPi-PFK. Monitor NADPþ reduction at 340 nm. 2.4.4. Assay of PPi-PFK with sedoheptulose-1,7-bisphosphate or ribulose-1,5-bisphosphate 0.4 ml reaction mixture contained: 50 mM HEPES–HCl, pH 7.0;

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mM sedoheptulose-1,7-bisphosphate (or 3 mM ribulose-1,5bisphosphate); 15 mM NaH2PO4; 5 mM MgCl2. The mixture was incubated for 3 min at 30  C and reaction was stopped by addition of 0.4 ml of 1 M H2SO4. Estimation of PPi (Heinonen et al., 1981): 0.4 ml of reagent consisting of 4 ml of 40 mM ammonium heptamolybdate, 1 ml of 5 M H2SO4, 50 mM of triethylamine is added to the mixture and let stand for at least 15 min and then centrifuged 5 min at 2000g. Then 0.2 ml of 5 M H2SO4 was added to the supernatant and centrifuged at the same condition. To the second supernatant 0.08 ml of 1 M 2-mercaptoethanol was added by mixing and the color developed for 15 min. The absorbance at a wavelength of 700 nm was measured. PPi concentration was calculated from a calibration line made with Na4P2O7 standard solutions (5–200 nmol in 0.4 ml of H2O).

2.5. Assay of fructosebisphosphate aldolase activity by using the recombinant PPi-PFK Activity of the fructosebisphosphate aldolase (FBA) toward the Fru 1,6-P2 synthesis was rarely studied due to the complexity of Fru 1,6-P2 detection. This problem may be resolved by using the highly active preparation of the recombinant PPi-PFK as the coupling enzyme (Rozova et al., 2010a). 1 ml of reaction mixture contained: 50 mM Tris–HCl, pH 7.5; 2 mM dihydroxyacetone-3-phosphate; 2 mM glyceraldehyde-3-phosphate; 1 mM NADPþ; 1 mM NaH2PO4; 4 mM MgCl2; 0.5 U of glucose phosphate isomerase (Sigma); 0.5 U of glucose-6-phosphate dehydrogenase (from rabbit muscle, Sigma); 0.5 U of recombinant PPi-PFK from Mm. alcaliphilum 20Z or Methylomonas methanica (Reshetnikov et al., 2005). After preincubation at 30  C for 3 min, 15 mU of FBA was added and the NADPþ reduction rate at 340 nm was measured.

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3. Brief Overview of Current Knowledge of PPi-PFK in Microorganisms PPi-PFK was first found in the anaerobic amitochondrial protozoan parasite Entamoeba histolytica H200 (Reeves et al., 1974, 1976). Subsequently, PPi-PFK activity was detected in other protozoans, in higher plants and in prokaryotes (Bruchhaus et al., 1996; Carnal and Black, 1983; Mertens et al., 1993; Petzel et al., 1989). Two types of PPi-PFK are known. Activity of type I PPi-PFK, which is a homopolymer with a subunit molecular mass of 40–50 kDa, is independent on fructose-2,6-bisphosphate (Fru 2,6-P2) and this enzyme has been found in protists and prokaryotes. Type II PPiPFK is a heterodimer, stimulated by Fru 2,6-P2 and found in photosynthetic organisms, higher plants and Euglena gracilis (Bruchhaus et al., 1996; Miyatake et al., 1984). The first report of the presence of PPi-PFK in the prokaryotic organism Propionibacterium freudenreichii appeared in 1975 (O’Brien et al., 1975). The enzyme (2  48 kDa) functions in either glycolysis or gluconeogenesis depending on growth substrate (glucose vs. glycerol or lactate). Synthesis of lipids, carbohydrates, proteins, nucleic acids, or NADH-dependent reduction of fumarate into succinate connected with the electron transport chain were proposed as sources of PPi for this enzyme (O’Brien et al., 1975). PPi-PFK has no allosteric effectors with the exception of the enzyme from Rhodospirillum rubrum, which was inhibited by AMP, ADP, and ATP (Pfleiderer and Klemme, 1980). In many cases, the physiological role of PPi-PFK is not obvious (Siebers et al., 1998). More precisely, the simultaneous presence of PPi-PFK and ATP-PFK (i.e., of a reversible and an irreversible enzyme) in a single organism, even if rarely reported, has suggested that PPi-PFK may perform an alternative unknown function (Alves et al., 2001; Van Praag, 1997). ATP-PFK and PPi-PFK share a common ancestry, but phylogenies show a very complex evolutionary history (Bapteste et al., 2003; Siebers et al., 1998). Several amino acids are essential for ATP-PFK and PPi-PFK functions (Moore et al., 2002). PFK working with ATP (with some exception) harbors Gly at positions 104 and 124 (according to the numbering of E. coli ATP-PFK), while PPi-PFK has an Asp104 and a Lys124. A singlepoint mutation can induce a change of the phospho-donor (Chi and Kemp, 2000). In addition to possible convergent adaptive mutations, horizontal gene transfer (HGT) events, including those of mutated sequences, further complicate the distribution of the enzymes. Hence, based on the PFK phylogenies it is not possible to conclude which phosphoryl donor, ATP or PPi, was ancestrally used by PFK (Bapteste et al., 2003).

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4. Properties of PPi-PFK from Methanotrophs PPi-PFK from Mc. capsulatus Bath is homodimeric (2  45), similar to the most known eubacterial enzymes, and that from Mm. alcaliphilum 20Z is homotetrameric (4  45). PPi-PFKs from both methanotrophs are not subjected to allosteric activation by organic compounds (ATP, ADP, AMP, NADP, NAD, and NADPH) and are thus probably regulated exclusively at the level of substrate and product concentrations (Reshetnikov et al., 2008; Rozova et al., 2010b). ATP, ADP, AMP, CTP, CDP, CMP, UTP, UDP, UMP, GTP, GDP, GMP at 1 mM as well as polyphosphates (n ¼ 3, n ¼ 9, n ¼ 15, and n ¼ 45) at 0.1 mg ml 1 could not substitute for PPi.

4.1. PPi-PFK from Mc. capsulatus Bath The enzyme from Mc. capsulatus Bath has the lowest affinity for both Fru 6-P and Fru 1,6-P2 in comparison to all known bacterial PPi-PFKs (Reshetnikov et al., 2008). It also reversibly phosphorylates Se 7-P with much higher activity and affinity and also displays activity with ribulose-5-P and ribulose-1,5-P2 (Table 1.1). Based on these properties of the enzyme, Table 1.1 Properties of PP-PFK from methanotrophs

Parameters

Vmax (U/mg) F-6-P S-7-P FBP Km (mM) PPi F-6-P S-7-P FBP Pi Mg2þ (forward reaction) Mg2þ (reverse reaction) Mr (kDa) (subunit number) Optimum pH Optimum temperature ( C) Effectors

Methylomicrobium alcaliphilum 20Z (Rozova et al., 2010b)

Methylomonas methanica 12 (Beschastny et al., 1992)

Methylococcus capsulatus Bath (Reshetnikov et al., 2008)

577 0.18 805

840 n.d. 850

7.6 31 9.0

0.118 0.64 1.01 0.095 3.4 0.22 0.33 180 (4  45) 7.5 30 No

0.051 0.39 n.d. 0.1 1.7 0.038 0.35 92 (2  45) 8.0 40 No

0.027 2.27 0.030 0.328 8.69 0.028 2 90 (2  45) 7.0 30 No

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its involvement in the nonoxidative pentose phosphate segment of the Calvin cycle, in which sedoheptulose and ribulose mono- and bisphosphates are the obligatory intermediates, may be proposed. The use of Se 7-P as a substrate was earlier shown for the E. histolytica PPi-PFK (Susskind et al., 1982). However, E. histolytica PPi-PFK has a lower affinity for Se 7-P (Km ¼ 64 mM) than for Fru 6-P (Km ¼ 38 mM). 4.1.1. Transcriptional organization of pfp and hpp genes in Mc. capsulatus Bath and their phylogeny The gene pfp encoding PPi-PFK and the gene hpp (MCA1252) encoding a putative V-type Hþ-pyrophosphatase (Hþ-PPi-ase, EC 3.6.1.1) are colocated in chromosome of Mc. capsulatus Bath (Ward et al., 2004). The closest homologues of the Hþ-PPi-ase is a protein from Geobacter sulfurreducens (TIGR_35554j2947) and R. rubrum (AAC38615) (62% and 58% identities). Cotranscription of the pfp and hpp genes was demonstrated by a standard RT-PCR approach (Reshetnikov et al., 2008). Putative promoter sequences (10 region TAAGTT;  35 region TTGTAA) were revealed 30 bp upstream of the start codon of pfp. According the accepted classification (Bapteste et al., 2003; Mu¨ller et al., 2001), PPi-PFK from Mc. capsulatus is clustered in clade B2 of the type II PFK enzymes and this clade is represented by eubacterial PPi-PFKs only. The most closely related neighbors of the Mc. capsulatus PPi-PFK are the enzymes from lithoautotrophic ammonia oxidizers bacteria Nitrosomonas europaea (Klotz et al., 2006) and Nitrosospira multiformis (Norton et al., 2008) and the facultative b-proteobacterial methylotroph Methylibium petroleiphilum PM1 (Kane et al., 2007). Similarly to Mc. capsulatus, the genomes of these bacteria also contain genes for ribulose bisphosphate carboxylase. Since no genes encoding ATP-PFK or fructose bisphosphatase (FBPase, EC 3.1.3.11) were revealed in the genome of Mc. capsulatus Bath (Ward et al., 2004), the single reversible PPi-PFK may fulfill the functions of these enzymes. Importantly, the bacteria containing PPi-PFK highly homologous to Mc. capsulatus Bath enzyme exhibit common biochemical features. They use monooxygenases requiring NADH or reduced cytochromes for energy substrate oxidation: methane monooxygenase (in methanotrophs), ammonia monooxygenase (in Nitrosospira and Nitrosomonas species), or toluene/ benzene monooxygenase (in M. petroleiphilum PM1) (Kane et al., 2007). Thus, oxidative phosphorylation may be hampered due to limiting by NADH or reduced cytochromes. Therefore, the use of PPi instead of ATP in some metabolic reactions may economize ATP and provide some advantages. Notably, in the lithoautotrophic bacteria, the pfp and hpp genes are also colocated in chromosomes. Coexpression of PPi-PFK and the putative Hþ-PPase suggests the hypothesis that the source of PPi for Fru 6-P phosphorylation may be energy-dependent PPi synthesis on the cell membrane. Conversely, PPi-PFK functioning in the reverse direction

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produces PPi, which may be utilized for generation of a Hþ gradient across the membrane, supporting ATP synthesis (Baltschevsky et al., 1999). The Mc. capsulatus PPi-PFK is clearly separated from a phylogenetic group “P” of PPi-PFKs from P. freudenreichii (O’Brien et al., 1975), Mm. alcaliphilum 20Z and M. methanica 12 (Reshetnikov et al., 2005) (12.6% and 16.5% identity, respectively).

4.2. PPi-PFK from Mm. alcaliphilum 20Z PPi-PFK from Mm. alcaliphilum 20Z showed very high activities and affinities for the forward and reverse reaction substrates, and could be involved in both glycolysis and gluconeogenesis (Rozova et al., 2010b). It appears to transform Fru 1,6-P2 more efficiently than Fru 6-P suggesting that a role of the enzyme in gluconeogenesis may be physiologically more significant. It also phosphorylates Se 7-P but with 103 times less activity than with Fru 6-P. In Mm. alcaliphilum 20Z, C3 compounds for biosynthetic needs (pyruvate and phosphotrioses) can be generated from C6 phosphosugars (the first products of the RuMP cycle) by cleavage via the Entner–Doudoroff pathway (Khmelenina et al., 1997, 2010). Thus, both C3 and C6 compounds can be formed bypassing the EMP pathway. PPi-PFK, together with FBA, may participate in regulating the ratio of C6 and C3 compounds in the cell. Since a candidate gene for the fructose-1,6-bisphosphatase (FBPase) is also present in the genome, some redundancy of the gluconeogenic activity occurs in Mm. alcaliphilum 20Z. Simultaneous functioning of PPi-PFK and FBPase may comprise a futile cycle for removing excess PPi which is the byproduct of many anabolic reactions including synthesis of sucrose accumulated by this halotolerant methanotroph at high levels in high salinity growth conditions (Khmelenina et al., 1999). The differential metabolic properties and functions of PPi-PFKs from Mm. alcaliphilum 20Z and Mc. capsulatus Bath correlate with their large sequence differences (16.5% identity), suggesting the possibility of an independent origin of these enzymes, along the lines of the hypothesis of “rampant” horizontal transfer of genes coding for microbial phosphofructokinases (Bapteste et al., 2003; Mu¨ller et al., 2001).

5. Conclusion Inorganic pyrophosphate created during biosynthetic polymerization reactions in most organisms is hydrolyzed by inorganic pyrophosphatase in order to thermodynamically favor the anabolic processes. Studied methanotrophs are characterized by negligible activity of inorganic

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pyrophosphatase and use PPi as the phosphoryl donor at least in the key glycolytic reaction phosphorylating Fru 6-P. Based on differential properties of PPi-PFKs from two methanotrophs and analyses of the recent genomic data, several important functions may be proposed for the enzyme: involvement in PPi recycling to economize ATP synthesis, regulation of hexosephosphates and triosephosphates interconversion as well as PPi levels in cells (in type I methanotrophic bacteria) and additionally in the Calvin cycle operation (in the type X methanotroph Mc. capsulatus Bath). Moreover, a gene encoding for PPDK catalyzing the PPi- and AMP-dependent conversion of PEP to pyruvate accompanied by ATP formation is present in the genomes of Mm. alcaliphilum 20Z and Mc. capsulatus Bath. Transcriptome analysis of Mm. alcaliphilum 20Z culture indicated that expression of PPDK increased under conditions of oxygen limitation (Kalyuzhnaya, personal communication). Thus, PPi-dependent glycolysis involving the two PPi-dependent enzymes, PPi-PFK and PPDK, may be proposed for this methanotroph. These two enzymes make glycolysis fully reversible. Importantly, PK and PEP synthetase encoding genes can also be expressed in this Mm. alcaliphilum 20Z. High flexibility at the key glycolytic steps may provide some advantages for methanotrophs, allowing metabolism to adapt to different energy states of cell. However, these hypotheses must be verified experimentally, particularly by characterizing the properties of the respective proteins. Further studies for unraveling the evolutionary history of PPi-dependent enzymes of glycolysis and the physiological role of PPi in aerobic methanotrophic bacteria are needed.

ACKNOWLEDGMENTS The work was supported by grants RFBR (08-04-01484-a and 10-04-01224-a), CRDF (Rub1-2946-PU-09), and joint PICS CNRS grant No3380. We thank the team at Ge´noscope for high-throughput sequencing and annotation of the M. alcaliphilum 20Z genome (IbiSA-Ge´noscope 2008 campaign) and M. G. Kalyuzhnaya for kindly providing microarray analysis information.

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C H A P T E R

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Genes and Enzymes of Ectoine Biosynthesis in Halotolerant Methanotrophs Alexander S. Reshetnikov, Valentina N. Khmelenina, Ildar I. Mustakhimov, and Yuri A. Trotsenko Contents 1. Introduction 2. PCR-Based Approach for Identification of the Ectoine Biosynthesis Genes in Methanotrophs 3. Transcriptional Regulation of the Ectoine Biosynthesis Genes 4. Key Enzymes of Ectoine Biosynthesis 4.1. Diaminobutyric acid aminotransferase 4.2. DABA acetyltransferase 4.3. Ectoine synthase 4.4. Aspartate kinase 5. Conclusion Acknowledgments References

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Abstract Ectoine (1,4,5,6-tetrahydro-2-methyl-4-pyrimidine carboxylic acid) is a widely distributed compatible solute accumulated by halophilic and halotolerant microorganisms to prevent osmotic stress in highly saline environments. Ectoine as a highly water keeping compound stabilizing biomolecules and whole cells can be used in scientific work, cosmetics, and medicine. Detailed understanding of the organization/regulation of the ectoine biosynthetic pathway in various producers is an active area of research. Here we review current knowledge on some genetic and enzymatic aspects of ectoine biosynthesis in halophilic and halotolerant methanotrophs. By using PCR methodology, the genes coding for the specific enzymes of ectoine biosynthesis, diaminobutyric acid (DABA) aminotransferase (EctB), DABA acetyltransferase (EctA), and ectoine synthase (EctC), were identified in several methanotrophic species. Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00002-4

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Organization of these genes in either ectABC or ectABC-ask operons, the latter additionally encoding aspartate kinase isozyme (Ask), correlated well with methanotroph halotolerance and intracellular ectoine level. A new gene, ectR1 encoding the MarR-like transcriptional regulatory protein EctR1, negatively controlling transcription of ectoine biosynthetic genes was found upstream of ectABC-ask operon in Methylomicrobium alcaliphilum 20Z. The ectR-like genes were also found in halotolerant methanol utilizers Methylophaga alcalica and Methylophaga thalassica as well as in several genomes of nonmethylotrophic species. The His6-tagged DABA acetyltransferases from Mm. alcaliphilum, M. alcalica, and M. thalassica were purified and the enzyme properties were found to correlate with the ecophysiologies of these bacteria. All these discoveries should be very helpful for better understanding the biosynthetic mechanism of this important natural compound, and for the targeted metabolic engineering of its producers.

1. Introduction Most halophilic and halotolerant microorganisms adapt to high environmental salinity by accumulation of low-molecular weight organic compounds called compatible solutes that equilibrate external osmotic pressure and support intracellular turgor that is higher than in the surroundings. The cyclic imino acid ectoine was originally discovered as an osmoprotectant in anoxygenic phototrophs of the Ectothiorhodospira group (Galinski et al., 1985) and subsequently found in many other Gram-negative and Gram-positive bacteria (Galinski, 1995; Kempf and Bremer, 1998; Severin et al., 1992), including the salt-dependent methanotrophs Methylomicrobium alcaliphilum, Mm. buryatense, Mm. kenyense, Methylobacter marinus (Kalyuzhnaya et al., 2001, 2008; Khmelenina et al., 1999, 2010; Trotsenko and Khmelenina, 2002) as well as methanol- and methylamine-utilizing bacteria of genera Methylophaga and Methylarcula (Doronina et al., 1998, 2000, 2003a,b). Ectoine is a compound compatible with cell metabolism even if accumulated to high intracellular concentrations. It has also been shown to act as a chemical chaperone increasing the stability of proteins. The mechanism of such stabilization can provide insights into protein folding. Moreover, ectoine and its hydroxylated derivative, hydroxyectoine, have attracted commercial interest for use in medicine and cosmetics as stabilizers of biomolecules and whole cells against different damaging factors such as heating, freezing, desiccation, and UV radiation (Buenger and Driller, 2004; Buenger and Driller, 2004; Graf et al., 2008; Jebbar et al., 1992; Sauer and Galinski, 1998). Since ectoines are produced only biotechnologically, understanding their biosynthetic pathways and regulation has been active area of research. The necessity of elucidation of the genes and enzymes responsible for ectoine biosynthesis in halophilic and halotolerant

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methylotrophs is conditioned by practical demands for new technologies of ectoine production from methane and methanol. The pathway of ectoine biosynthesis in the bacteria studied represents a biochemical sequence that is a branch of aspartate family amino acids synthesis (Peters et al., 1990). Three special enzymes are involved in this pathway: diaminobutyric acid (DABA) aminotransferase (EctB, EC 2.6.1.76), catalyzing transamination of aspartate semialdehyde into DABA, DABA acetyltransferase (EctA, EC 2.3.1.178), acetylating DABA into Ngacetyl-DABA, and ectoine synthase (EctC, EC 4.2.1.108), which forms ectoine by cycling of Ng-acetyl-DABA. In most halophilic bacteria studied, the genes encoding these enzymes are combined in chromosomes in the cluster ectABC (Canovas et al., 1999; Go¨ller et al., 1998; Kuhlmann and Bremer, 2002; Louis and Galinski, 1997). Database searches reveal the highly homologous ectoine biosynthetic genes in 200 finished genomes of bacteria belonging to phyla Proteobacteria, Actinobacteria, and one archaeon Nitrosopumilus maritimus. Sometimes, an additional gene ectD coding for ectoine hydroxylase is present and this enzyme converts ectoine to hydroxyectoine (Bursy et al., 2007; Garcı´a-Estepa et al., 2006; Prabhu et al., 2004). High homology of the ectoine biosynthesis genes in various microorganisms of different taxonomic position and physiological properties is a reliable indication of the evolutionary conservation of this biochemical pathway (Kuhlmann and Bremer, 2002). We present below a brief overview of current knowledge on the peculiarities of the ectoine biosynthetic pathway in aerobic halophilic and halotolerant methanotrophs, focusing on the main genetic and biochemical techniques that have been used to unravel mechanisms regulating this important natural product biosynthesis. In the first section, we highlight the strategies that are used for identification of ectoine biosynthetic genes in methanotrophs. In the second section, we describe initial results of transcriptional analysis of ect-genes. The third section briefly describes some properties of the ectoine biosynthesis enzymes. Despite the focus on the ectoine biosynthetic pathway in methanotrophs, many of the techniques could be readily adapted to the study of other ectoine producers.

2. PCR-Based Approach for Identification of the Ectoine Biosynthesis Genes in Methanotrophs Since the genes coding for the ectoine biosynthetic enzymes contain highly conserved regions and are generally organized in an ectABC operon, PCR was very helpful methodology to identify them in Mm. alcaliphilum 20Z (Reshetnikov et al., 2006). Assuming that ect-genes in other

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methanotrophs are also combined in the chromosome as an ectABC operon, we have been exploring the use of better-focused probes more specific for the ectoine biosynthetic gene cluster (Reshetnikov et al., 2006). Based on the alignment of the conserved regions of DABA aminotransferase, DABA acetyltransferase, and ectoine synthase, several degenerate primers Tra1, Tra4, Tra3, CR, and Atf for PCR have been designed and tested (Table 2.1, Fig. 2.1A). Primers Tra1 and Tra4 gave a single specific product about 750 bp in size with Mm. alcaliphilum genomic DNA as a template and this product was shown to match the predicted sequence. Degenerate primer pair Tra3/CR was used to amplify a
1000-bp fragment containing the 30 end of ectB and 50 end of ectC genes. By using another degenerate primer, Atf, corresponding to a segment of ectA and the primer Tra2.1 homologous to 50 region of ectB, a DNA fragment of 700 bp was amplified. All these PCR products were sequenced and analyzed using Vector NTI v.9 (Invitrogen). The assembled sequence of 2100 bp contained three orfs (one complete and two incomplete) oriented in the same direction. These orfs were revealed by database searches using BLAST

Table 2.1 Oligonucleotide primer sequences used for studying of ectoine biosynthesis genes in methanotrophs

Primers

Gene targeted

Sequences (50 –30 )

Tra1 Tra2 Tra2.1 Tra3 Tra4 CR C20 EF AskF AskR AskIR AskIF AtrI Atf ATZ AMOFor AMOA AMOR AMOZ AMOF 7CF 7CR

ectB ectB ectB ectB ectB ectC “vectorette” ectC ask ectB ask ask ectA ectA ectA ectA ectA ectB ectB ectC ectC ectB

CCCTIAA(T/C)TA(T/C)GGICA(T/C)AA(C/T) CC(G/A)TG(A/G)AAI(G/C)C(G/A)TTIGT(G/A)AA ATCTTCAGTGCCGCTTCAACC ACCGG(T/C)ACITT(C/T)TT(C/T)AGITT(C/T)GA CG(G/A)AAIGTGCC(A/G)TT(G/A)TG(T/C)TCNCC GGIGG(A/G)TT(A/G)AANAC(A/G)CA CTCTCCCTTCTCGAATCGTAA TGCTGCTGAAGGATGACGGTATGGGCT AGCATCTTTGTCGGCACGGTCATT CCCGCTTTGCTGACCTCTTTACTGA GCCTCATCCGCCTTGGTCAGTA TGCGTAGAGATTATTGCCGGGTGT ACAAAACCAACCAACTCATCGCCA CTIGA(T/C)(C/T)(T/C)IAA(T/C)TCI(T/G/A)(C/T)ITA TT(C/T)GT(G/T/C)TGGCA(A/G)GTIGCNGT CGAAATGCTGGTTAAGATTGGTCCGT ATTGGGCGGATTGATCGTAGTTTC ATTTGATCGGCGAAACGGTTAT CTTGAGGCTTGCCTCGGCTATC CGACGATTTATCAAGGTGCGGA CGATCAAGGCGGGCACGGTCTAT TCGACAATGGCCTGCAGACTT

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Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs

O. B. B. M. S. V. C.

iheyensis pasteurii halodurans halophilus coelicolor cholerae salexigens

:GAGALNYGHNPSEM: :GAGSLNYGHNNEKM: :GAGALNYGHNDEKM: :GAGALNYGHNDENM: :GAGSLNYGHNNAVL: :GAGALNYGHNNAVL: :GAGTLNYGHNHPKL:

O. B. B. M. S. V. C.

iheyensis pasteurii halodurans halophilus coelicolor cholerae salexigens

Tra1 (forvard) CCCTIAA(T/C)TA(T/C)GGICA(T/C)AA(C/T)

O. B. B. M. S. V. C.

iheyensis pasteurii halodurans halophilus coelicolor cholerae salexigens

:PGEHNGTFRGNN LA: :PGEHNGTFRGNN MA: :PGEHNGTFRGNN HA: :PGEHNGTFRGNN FA: :PGEHNGTFRGNN PA: :PGEHNGTFRGNN HA: :PGQYNGTFRGFNLA:

Tra4 (reversed) CG(A/G)AAIGTGCC(G/T)TT(A/G)TG(C/T)TCNCC

: : : : : : :

GRTGTFFSFEEAGINPD GRTGTFFSFEPAGIQPD GRTGTFFSFEDAGITPD GRTGTFFSFEPAGIKPD GRTGAFFSFEEAGITPD GRTGTFFSFEPSGIEPD GRTGKFFSFEHAGITPD

Tra3 (forvard) ACCGG(T/C)ACITT(C/T)TT(C/T)AGITT(C/T)GA

M. B. B. V. C. S. O.

halophilus : halodurans : pasteurii : cholerae : salexigens : coelicolor : iheyensis :

MVCVFNPPL MVCVFNPPI LICTFNPPL MACVFNPPL VACVFNPAL CICVFNPPV MVCVFNPAL

CR (reversed)

GGIGG(G/A)TT(G/A)AANAC(G/A)CA

Figure 2.1 Conserved regions of DABA aminotransferase (EctB) and ectoine synthase (EctC) homologues and degenerate primers designed for ectB (Tra1, Tra3, and Tra4) and ectC (CR) genes.

to encode proteins with high amino acid sequence identities to the ectoine biosynthesis enzymes: DABA acetyltransferase (EctA), DABA aminotransferase (EctB), and ectoine synthase (EctC). Then, the newly obtained sequences were used to design primers AtrI (homologous to ectA) and EF (homologous to ectC) which were used in inverse PCR (IPCR) to clone the missing 50 end of ectA and 30 end of ectC. In the PCR product of 1.8 kb, a new incomplete orf downstream of ectC was found which showed a significant degree of derived polypeptide (50% amino acid sequence identity) similar to ectoine-associated aspartokinase (Ask) from Vibrio cholerae. To identify the 30 region of the putative ask gene, IPCR was performed with the self-ligated PvuI-fragment and primers AskR homologous to ect and AskF homologous to ask (Fig. 2.2A). For the second IPCR, 500 ng of chromosomal DNA was digested with MunI in a 50 ml reaction volume. DNA fragments were then ligated at 16  C overnight in a 100 ml reaction volume containing 200 ng of digested DNA and 2U of T4 DNA ligase in 1 ligase buffer as recommended by the manufacturer. Circular DNA molecules were used as a template for PCR with primers AskIR and AskIF homologous to the two ask segments. In the sequenced and assembled resultant products, another orf of 948 bp was found 360 bp downstream of ectABC-ask (Fig. 2.2A). Its closest homologue was a gene encoding ectoine hydroxylase (EctD) from Streptomyces avermitilis MA-4680 (GenBank BAC 74106) and S. chrysomallus (GenBank AAS02097) (40% and 42% identities), which is a member of the nonheme

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Alexander S. Reshetnikov et al.

A PvuI

MunI Atf

ectR

Tra2.1 Tra2 Tra3

ectA AtrI

B

ectB Tra1

Tra4

AMOA

phyH

ask

EF

Tra4

Tra1

MunI

ectC CR

AclI

Eco RI Tra3

ectA C20

MunI

AskF

ectB

ectA

C

PvuI

AskIR AskIF

MunI AMOR AMOZ AMOF Tra 3

AMOFor

ATZ

CR

ectC

AclI

AclI

MunI

AskR

CR

ectB

ectC 7CR

7CF

Na-transporter

C20

Figure 2.2 Sequencing strategies used for identification of ectoine biosynthetic genes in Methylomicrobium alcaliphilum 20Z (A), Methylomicrobium kenyense AMO1 (B), and Methylobacter marinus 7C (C). The nucleotide sequences of oligonucleotide primers are given in Table 2.1.

iron(II)- and 2-oxoglutarate-dependent dioxygenase family (Reuter et al., 2010). Although no hydroxyectoine accumulation was detected by 1H NMR in Mm. alcaliphilum 20Z cells grown at salinities of 3% or 9% NaCl, it is still possible that the organism can make this compound under some growth conditions. From the assembled five-gene cluster, only four genes encoding DAB acetyltransferase (EctA, 18.8 kDa), DAB transaminase (EctB, 47.8 kDa), ectoine synthase (EctC, 15.2 kDa), and L-aspartokinase (Ask, 53.3 kDa) are transcribed as a single polycistronic mRNA as was shown by RT-PCR (Reshetnikov et al., 2006). The ectABC genes could be functional in Escherichia coli XL1-Blue since recombinant cells carrying them in the low-copy-number vector pHSG575 grew in the presence of 4% NaCl and synthesized ectoine (Reshetnikov et al., 2006). The analogous PCR strategy was used for deciphering the nucleotide sequences of ect-genes in another alkaliphilic methanotroph, Mm. kenyense AMO1 (Fig. 2.2B). Only three orfs corresponding to ectA, ectB, and ectC were found in a 3.5 kb genomic fragment of Mm. kenyense AMO1 with intergenic regions of 47 bp between ectA and ectB and 170 bp between ectB and ectC. No additional orf was found within 600 bp downstream of the ectC gene. PCR with degenerate primers Tra3/CR as the first step was also used to identify ect-genes in Mb. marinus 7C. On the basis of the sequence obtained (
1000 bp), the homologous primers 7CF and 7CR were synthesized and

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used in “Vectorette” PCR performed as described by Ko et al. (2003). For this purpose, samples of genomic DNA were digested by EcoRI. After annealing and ligation of double strand “vectorette” unit (vect 57TTAA which complemented vect 53) to DNA digested by EcoRI, a “vectorette” PCR amplification with primer 7CF was performed (Table 2.1, Fig. 2.2C). For the second “vectorette” PCR, genomic DNA was digested by AclI. After annealing and ligation of “vectorette” unit (vect 57CG which also complemented vect 53), primer 7CR was used in the second round of PCR amplification. Generated fragments were sequenced and assembled. The 4.25 kb DNA fragment from M. marinus 7C included tightly linked ectABC genes without intergenic regions. An additional orf4 (1371 bp) was detected in the DNA locus 81 bp downstream of ectC. The product of this Orf4 (49 kDa) was similar to transport proteins belonging to Naþ/solute symporter family (SSF) from Mariprofundus ferrooxydans PV-1 ZP_01451115 (56% identity of amino acid sequences), Nitrosococcus oceani ATCC 19707 YP_343605 (46%), and Hyphomonas neptunium ATCC 15444 YP_760353 (43%). Two main types of ectoine gene organization were found in the methanotrophs which correlated with their salt tolerance and intracellular ectoine contents. Thus, Mb. marinus 7C and Mm. kenyense AMO1 possessing ectABC genes accumulated ectoine up to 70 mg per g of DCW and these methanotrophs were capable of growth at salinity 4–5% NaCl (Khmelenina et al., 2010). Conversely, Mm. alcaliphilum 20Z carrying the ectABC-ask operon was able to grow at higher salinity (up to 10% NaCl) and accumulate more ectoine (>120 mg per g of DCW). This implied an important role of a specific aspartate kinase in ectoine synthesis. Conversion of aspartate into b-aspartyl phosphate by aspartate kinase represents the starting point of the pathway for biosynthesis of both aspartate family amino acids and ectoine. Being osmotically controlled, aspartate kinase could make ectoine accumulation more independent from complex machinery regulating the divergent pathways leading from aspartate semialdehyde. Specialized ectoine-associated Ask enzymes can also be found in other halophilic Proteobacteria as judged from homology analysis of aspartate kinases (Lo et al., 2009). Remarkably, no Gram-positive bacteria contain an ectABC-ask gene cluster. The absence of the ask homologs in ectoine gene clusters of Firmicutes is explainable since carbon flow must also be directed to biosynthesis of the osmoprotectants belonging to the glutamate family amino acids, proline and/or glutamine, contributing to osmotic balance (Kuhlmann and Bremer, 2002; Saum and Muller, 2008). The second Ask encoded by the separate gene may occur in the Mm. alcaliphilum 20Z genome (https://www.genoscope.cns.fr/agc/mage) and it shares 17% identity with the osmotically controlled Ask. Notably, Ect proteins from the gammaproteobacterial alkaliphilic methanotrophs Mm. alcaliphilum 20Z and Mm. kenyense AMO1 are only

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distantly related (37–51% identities) to the enzymes from neutrophilic Mb. marinus 7C (also belonging to the Gammaproteobacteria) being most closely related to Ect proteins from zetaproteobacterial M. ferrooxydans (55– 80% identities). So far, Mb. marinus is the single species of the Methylobacter genus that is salt-resistant. It could be proposed that the ect operon ubiquity in ancient prokaryotic world that was largely marine was followed by loss in lineages that became adapted to a terrestrial environment (Lo et al., 2009).

3. Transcriptional Regulation of the Ectoine Biosynthesis Genes Routine genetic manipulations (DNA and RNA isolation from Mm. alcaliphilum 20Z, construction of mutant strains, Northern hybridization, and mapping of the transcriptional start sites) are described previously (Mustakhimov et al., 2010a) and elsewhere in this volume (Ojala et al., 2010). Analysis of the promoter region sequence showed that the ectABC-ask operon in Mm. alcaliphilum 20Z is initiated from two s70-like promoters ectAp1 and ectAp2. ectAp1 has a high level of identity with the consensus sequence of the E. coli s70-recognized promoter while the 10 and 35 sequences of ectAp2 differ from the respective regions of the E. coli s70promoter. Therefore, expression from ectAp2 could be less effective than from ectAp1. Northern hybridization also indicated that the ectC and ask genes might be cotranscribed as an additional transcriptional unit from the promoter region located upstream of the ectC gene. Transcription of ectR1 was carried out from a single s70-like promoter ectR1p located between ectAp1 and ectAp2. This implies that transcription from ectR1p may be controlled by its own product (Mustakhimov et al., 2010a). Upstream of ectA an additional gene transcribed in the reverse orientation was revealed (Mustakhimov et al., 2010a). This gene was named ectR1 (ectoine biosynthesis regulator) and it encoded a protein (20.6 kDa) homologous to the MarR-type transcriptional regulators (Wilkinson and Grove, 2006). A regulatory function of EctR1 was confirmed by studies of the ectR1 knockout mutant. Like most MarR-family regulators, EctR1 was shown to negatively control transcription of the ect operon by binding the promoter region (Mustakhimov et al., 2010a,b). It protects from DNAse I action an asymmetrical DNA locus that includes the 10 sequence of the ectA1p. The EctR1 binding site contains a pseudopalindromic sequence composed of 8-bp half-sites separated by 2 bp (TATTTAGT-GT-ACTATATA) suggesting dimeric association of the EctR1 with the DNA in which each subunit binds an inverted repeat. Gel-filtration data showed that EctR1 is

Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs

23

a dimer both in free solution (44–45 kDa) and in the DNA-binding state (50–55 kDa). The ectR1-like genes were identified in the methanol- and methylamine-utilizing bacteria Methylophaga alcalica, Methylophaga thalassica, and Methylarcula marina (76%, 66%, and 54% identities of translated amino acid sequences) as well as in 22 genomes of heterotrophic halophiles (35.5–55.1% identities) (Mustakhimov et al., 2010b; Reshetnikov et al., 2010). All the EctR-like proteins showed relatively low identity (20%) with the characterized MarR-family transcriptional regulators. MarRfamily proteins include a diverse group of regulators controlling various physiological functions including response to environmental stresses, virulence factors, and aromatic catabolic pathways (Wilkinson and Grove, 2006). Although EctR1 and the MarR-family regulators have only low sequence identities, they likely have analogous protein structures due to the presence of helix-turn-helix DNA-binding motifs and flanking “wing 1” regions (Mustakhimov et al., 2010a).

4. Key Enzymes of Ectoine Biosynthesis As described above, the genes for ectoine biosynthesis in Mm. alcaliphilum 20Z are organized in one transcription unit ectABC-ask and have high similarity to the genes occurring in all groups of bacteria synthesizing ectoine. The encoded proteins have the following enzyme activities: EctB is diaminobutyric acid aminotransferase, EctA is diaminobutyric acid acetyltransferase, EctC is ectoine synthase, and Ask is aspartate kinase.

4.1. Diaminobutyric acid aminotransferase This enzyme was tested in a 1-ml assay volume containing 50 mM Tris– HCl buffer, pH 8.5, 10 mM 2-oxoglutarate, 20 mM diaminobutyric acid, 10 mM pyridoxal-50 -phosphate, 100 mM KCl and an enzyme preparation with activity in the range of 100 mU. The mixture was incubated at 25  C for 30 min. The reaction was stopped by boiling the mixture for 5 min and insoluble material was separated by centrifugation. In all, 20 ml samples were withdrawn and mixed with 5 ml of o-phthalaldehyde reagent (50 mg o-phthalaldehyde was dissolved in 1.25 ml of methanol, 50 ml of mercaptoethanol, and 11.2 ml 0.4 M borate buffer, pH 9.5). After 1 min exposure, 45 ml Na-acetate, pH 7.0, was added and immediately loaded onto a reversed-phase SEPARON SIX C-18 column (150  3.3 mm, 5 mm, Czech Republic). Degassed solutions of 0.1 M Na-acetate, pH 7.2, and methanol (0–40%) were used for gradient elution with a flow of 1 ml/min. Measurements were carried out on a fluorimeter “Gilson 121” (France) in

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9 ml cuvette with filters for excitation 305–395 and emission at 420–650 nm. Glutamic acid was used as inner internal standard. DABA aminotransferase from Mm. alcaliphilum 20Z was shown to be a 300 kDa hexameric enzyme with specific activity of 0.027 U/mg of protein and a calculated pI 5.49. DABA aminotransferase from Halomonas elongata purified by classical biochemical methods (Ono et al., 1999) was also a homohexameric (
250 kDa) pyridoxal-50 -phosphate-dependent enzyme. It required Kþ for activity and stability and was more active in the presence of 0.01–0.5 M KCl than at the same concentrations of NaCl, specific to L-glutamate as aminodonor (Km for L-glutamate 9.1 mM) and to D,L-aspartyl semialdehyde (Km 4.5 mM). The reaction catalyzed by DABA aminotransferase is thought to be a limiting step of the ectoine biosynthetic pathway thus explaining the absence of DABA in H. elongata KS3 cells (Ono et al., 1999). Importantly, the second putative DABA aminotransferase gene located outside of ectABC-ask operon clustering with genes coding for ectoine degradation enzymes were found in the draft genome of Mm. alcaliphilum 20Z.

4.2. DABA acetyltransferase DABA acetyltransferase (EctA) acetylates diaminobutyric acid with acetyl coenzyme A forming g-N-acetyl-a,g-diaminobutyric acid. The enzyme activity was tested at pH 9.0 and 25  C in a 1-ml assay volume containing 50 mM Tris/HCl buffer, 0.1 mM 5,5-dithio-bis-(2-nitrobenzoic acid) (DTNB), 10 mM diaminobutyric acid, 1 mM acetyl-CoA, 200 mM KCl, and enzyme preparations with activity in the range of 10 mU. Formation of 2-thio-5-nitrobenzoic acid as a result of the interaction between 5,5-dithiobis-(nitrobenzoic acid) and sulfhydryl groups of coenzyme A released in the course of the reaction was monitored at l ¼ 412 (e412 ¼ 13.6 mM 1 cm 1) (Reshetnikov et al., 2006). DABA acetyltransferase from Mm. alcaliphilum 20Z was obtained by cloning of ectA into a pET22bþvector (Novagen, USA) and expression in E. coli BL21 (DE3). The His-tagged enzyme was purified on a Ni-NTAagarose column (Qiagen, Germany) in one stage (Reshetnikov et al., 2005). The kinetic properties of the Mm. alcaliphilum DABA acetyltransferase are presented in Table 2.2. Similar methods have been used for obtaining the purified recombinant EctA enzymes from two moderately halophilic methylotrophic bacteria M. thalassica and M. alcalica (Mustakhimov et al., 2008). Comparison of the enzyme properties revealed their correlation with ecophysiologies of host bacteria. Thus, the DABA acetyltransferases from the neutrophilic bacteria H. elongata (Ono et al., 1999) and M. thalassica were more active at lower pH (pH 8.2 or 9.0), than those from the alkaliphilic species Mm. alcaliphilum 20Z and M. alcalica (pH optima  9.5). DABA acetyltransferase from M. thalassica isolated from marine water was considerably inhibited by carbonates while the enzymes from the soda lake isolates

25

Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs

Table 2.2 Some properties of diaminobutyric acid acetyltransferases from methylotrophic bacteria

Properties

a

Mm. alcaliphilum M. alcalica M. thalassica 20Z (Reshetnikov (Mustakhimov (Mustakhimov et al., 2005) et al., 2008) et al., 2008)

pHopt Topt ( C) Molecular mass (SDS-PAGE) (kDa) Mr (gel-filtration) (kDa) Km (DABA) (mM) Km (acetyl-CoA) (mM) Inhibitors (1 mM)

9.5 20 20

9.5 30–35 20

8.5 30–35 20

40 0.465 36.7 Zn2þ, Cd2þ

Optimum KCl Optimum NaCl Stability

0.25 M 0.1–0.2 M Stablea

40 0.375 30 Zn2þ, Cd2þ, Cu2þ 0 0 Stablea

40 0.365 76 Zn2þ, Cd2þ, Cu2þ 0 0 Stablea

The enzyme was stable in 50 mM Tris–HCl, pH 8.5, at least for 1 month at 4–70  C at protein concentration 0.5 mg/ml.

M. alcalica and Mm. alcaliphilum 20Z were not affected, thus corresponding to in situ physiology of these species. Interestingly, Cu2þ at 1 mM completely inhibited the enzyme activity from M. alcalica and 47% of that of M. thalassica. In contrast, no inhibitory effect of Cu2þ was found for the Mm. alcaliphilum 20Z enzyme thus confirming an important role of copper in methane oxidation by the particulate methane monooxygenase of the methanotroph (Murrell et al., 2000). The differential dependence on ionic strength is another intriguing feature of the DABA acetyltransferases. The enzymes from H. elongata and Mm. alcaliphilum 20Z were activated by 0.4 or 0.2 M NaCl whereas the enzymes from M. thalassica and M. alcalica were inhibited by these salts. It might be speculated that the methanol-utilizing bacteria have more effective ion extrusion mechanisms in comparison to methane- or glucose-assimilating bacteria. We hypothesize that low monovalent inorganic ion concentrations are maintained in cytoplasm of the methanol utilizers. Different behavior of the enzyme at high or low ionic strength may be a result of its long-term adaptation in the host species.

4.3. Ectoine synthase The enzyme activity was tested at pH 9.0 and 25  C in a 1-ml assay volume containing 50 mM sodium-phosphate buffer, 20 mM N-a,g-diaminobutyric acid, and 300 mM NaCl and 10 mU of the recombinant EctC. The reaction

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was stopped by boiling the mixture for 5 min and insoluble material was separated by centrifugation for 20 min at 12,000g. The soluble fraction was dried under vacuum and ectoine content was measured by HPLC (Eshinimaev et al., 2007). A preparation of the recombinant ectoine synthase with specific activity of 64 U/mg of protein was obtained from Mm. alcaliphilum 20Z. The molecular mass of the purified ectoine synthase (35 kDa) suggests that the enzyme is a homodimer. The pI is 4.99. The native homogenous ectoine synthase purified from H. elongata with activity of 16 U/mg of protein was purified in the presence of 1 mM N-acetyl-DABA and 2 M NaCl as stabilizing compounds (Ono et al., 1999). The molecular mass of the native enzyme remained unclear because no activity was found after gel-filtration at 0.5 M NaCl. The enzyme was specific to Ng-acetyl-DABA and N-acetyl group in a-position could not be involved in cycling. The protein contained enhanced levels of aspartate and glutamate.

4.4. Aspartate kinase Halotolerant bacteria may use a specialized Ask paralog and the specialization can be achieved by differential regulation accomplished via allosteric control of enzyme activity and/or by transcriptional/translational control of enzyme synthesis (Lo et al., 2009). However, the ectoine-associated Ask enzyme of halophilic/halotolerant bacteria has never been studied. We made several attempts to obtain the recombinant enzyme by cloning of the gene ask either into vector pET30 (Novagen) designed to express a C-terminal His-tagged fusion protein or pET28 (Novagen) designed to express a N-terminal His-tagged fusion protein. However, E. coli BL21 (DE3) transformed with the pET30/ask or pET28/ask plasmids synthesized a non-soluble protein. Co-expression of aspartate kinase with chaperones (GroES/EL) was also accompanied by insoluble protein synthesis. Aspartate kinase in the soluble form (53.3 kDa) was obtained by expression in Methylobacterium extorquens AM1. Gene ask was amplified from pET28/ask or pET30/ask and DNA fragments were cloned into vector pCM160 carrying constitutive promoter of methanol dehydrogenase (Pmax) from M. extorquens AM1 (Marx and Lidstrom, 2001). The resulting plasmid pCMaskN-his (or pCMaskC-his) was transferred into M. extorquens AM1 by conjugation and cells were grown on mineral medium supplemented by 0.5% (v/v) methanol. A single step of immobilized metal ion affinity chromatography on a Ni2þ-NTA column was performed for purification of His6-tagged Ask fusion protein from M. extorquens extract (Mustakhimov et al., 2008). The preparation of the recombinant enzyme with activity 25 mU/mg of protein was obtained. A protein band corresponding to

Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs

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50 kDa was obtained on SDS-PAGE. Analysis of the insoluble fractions did not reveal the corresponding protein. A slightly modified spectrophotometric method (Wampler and Westhead, 1968) was used for testing of aspartate kinase activity by following the ADP production rate in a 1-ml assay volume containing 50 mM Tris–HCl (pH 8.0) buffer, pH 8.0, 5 mM L-aspartic acid (sodium salt), 8 mM ATP (sodium salt), 5 mM phosphoenolpyruvate, 0.25 mM NADH, 1 mg pyruvate kinase and 1 mg lactate dehydrogenase and
1 U of the tested enzyme. The rate of ATP-dependent NADH oxidation at 25  C was monitored at 340 nm on Shimadzu UV-160 recording spectrophotometer (Japan).

5. Conclusion In spite of the significant progress that has been made in understanding the genetic and biochemical basis of ectoine biosynthesis in halophilic/ halotolerant bacteria, numerous important questions remain to be addressed. Although the ectoine biosynthetic pathway is commonly similar in methanotrophic and heterotrophic bacteria with respect to three specific enzymes, the ectoine operon in some cases is supplemented with ectD gene coding for ectoine hydroxylase and/or the ask gene coding for aspartate kinase. The ectoine-associated aspartate kinase occurs only in Gram-negative bacteria and this provides the host methanotrophic strains with enhanced halotolerance. However, the properties of this specialized enzyme in halophiles remain to be explored. Similar to Mm. alcaliphilum 20Z, several other Gram-negative bacteria may harbor the EctR-like transcriptional repressor belonging to the MarR-family of proteins. It is also possible that other osmoregulating genes will be identified. Moreover, this is also complicated by preliminary genomic evidence for the existence of the Mm. alcaliphilum 20Z ectoine degradation pathway found in Halomonas elongata (Schwibbert et al., 2010). Osmoadaptation of aerobic methanotrophs includes, besides the osmoprotective compatible solutes biosynthesis (ectoine, glutamate, and in some case sucrose), other structural–functional mechanisms, for example, changes in phospholipid fatty acids composition and in bioenergetics machinery (Khmelenina et al., 1997). To date, it is not possible to describe the whole regulatory cascade from sensing the osmotic signals by the cell membrane to the resultant metabolic and structural rearrangement events.

ACKNOWLEDGMENTS The work was supported by grants RFBR (10-04-01224a) and CRDF (Rub1-2946-PU-09).

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and emended description of the genus Methylomicrobium. Int. J. Syst. Evol. Microbiol. 58, 591–596. Kempf, B., and Bremer, E. (1998). Uptake and synthesis of compatible solutes as microbial stress responses to high-osmolality environments. Arch. Microbiol. 170, 319–330. Khmelenina, V. N., Kalyuzhnaya, M. G., and Trotsenko, Y. A. (1997). Physiological and biochemical properties of haloalkalitolerant methanotroph. Microbiology (Moscow) 66, 365–370. Khmelenina, V. N., Kalyuzhnaya, M. G., Sakharovsky, V. G., Suzina, N. E., Trotsenko, Y. A., and Gottschalk, G. (1999). Osmoadaptation in halophilic and alkaliphilic methanotrophs. Arch. Microbiol. 172, 321–329. Khmelenina, V. N., Shchukin, V. N., Reshetnikov, A. S., Mustakhimov, I. I., Eshinimaev, B. Ts., Suzina, N. E., and Trotsenko, Y. A. (2010). Structural and functional features of methanotrophs from hypersaline and alkaline lakes. Microbiology (Moscow) 79, 472–482. Ko, W.-Y., Ryan, M. D., and Akashi, H. (2003). Molecular phylogeny of the Drosophila melanogaster species subgroup. J. Mol. Evol. 57, 562–573. Kuhlmann, A. U., and Bremer, E. (2002). Osmotically regulated synthesis of the compatible solute ectoine in Bacillus pasteurii and related Bacillus spp.. Appl. Environ. Microbiol. 68, 772–783. Lo, C.-C., Bonner, C. A., Xie, G., D’Souza, M., and Jensen, R. A. (2009). Cohesion group approach for evolutionary analysis of aspartokinase, an enzyme that feeds a branched network of many biochemical pathways. Microbiol. Molec. Biol. Rev. 73, 594–651. Louis, P., and Galinski, E. A. (1997). Characterization of genes for the biosynthesis of the compatible solute ectoine from Marinococcus halophilus and osmoregulated expression in Escherichia coli. Microbiology (UK) 143, 1141–1149. Marx, C. J., and Lidstrom, M. E. (2001). Development of improved versatile broad-hostrange vectors for use in methylotrophs and other gram-negative bacteria. Microbiology 147, 2065–2075. Murrell, J. C., McDonald, I. R., and Gilbert, B. (2000). Regulation of expression of methane monooxygenases by copper ions. Trends Microbiol. 8, 221–225. Mustakhimov, I. I., Rozova, O. N., Reshetnikov, A. S., Khmelenina, V. N., Murrell, J. C., and Trotsenko, Y. A. (2008). Characterization of the recombinant diaminobutyric acid acetyltransferase from Methylophaga thalassica and Methylophaga alcalica. FEMS Microbiol. Lett. 283, 91–96. Mustakhimov, I. I., Reshetnikov, A. S., Glukhov, A. S., Khmelenina, V. N., Kalyuzhnaya, M. G., and Trotsenko, Y. A. (2010a). Identification and characterization of EctR1, a new transcriptional regulator of the ectoine biosynthesis genes in the halotolerant methanotroph Methylomicrobium alcaliphilum 20Z. J. Bacteriol. 192, 410–417. Mustakhimov, I. I., Reshetnikov, A. S., Khmelenina, V. N., Murrell, J. C., and Trotsenko, Y. A. (2010b). Regulatory aspects of ectoine biosynthesis in halophilic bacteria. Microbiology (Moscow) 79, 583–592. Ojala, D. S., Beck, D. A. C., and Kalyuzhnaya, M. G. (2010). Genetic systems for moderately halo(alkali)philic bacteria of the genus Methylomicrobium. Methods Enzymol. 495. Ono, H., Sawada, K., Khunajakr, N., Toa, T., Yamamoto, M., Hiramoto, M., Shinmyo, A., Takano, M., and Murooka, Y. (1999). Characterization of biosynthetic enzymes for ectoine as a compatible solute in a moderately halophilic eubacterium, Halomonas elongata. J. Bacteriol. 181, 91–99. Peters, R., Galinski, E. A., and Tru¨per, H. G. (1990). The biosynthesis of ectoine. FEMS Microbiol. Lett. 71, 157–162. Prabhu, J., Schauwecker, F., Grammel, N., Keller, U., and Bernhard, M. (2004). Functional expression of the ectoine hydroxylase gene (thpD) from Streptomyces chrysomallus in Halomonas elongata. Appl. Environ. Microbiol. 70, 3130–3132.

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Reshetnikov, A. S., Mustakhimov, I. I., Khmelenina, V. N., and Trotsenko, Y. A. (2005). Cloning, purification and characterization of the diaminobutyrate acetyltransferase from the halotolerant methanotroph Methylomicrobium alcaliphilum 20Z. Biochemistry (Moscow) 70, 878–883. Reshetnikov, A. S., Khmelenina, V. N., and Trotsenko, Y. A. (2006). Characterization of the ectoine biosynthesis genes of haloalkalotolerant obligate methanotroph “Methylomicrobium alcaliphilum 20Z” Arch. Microbiol. 184, 286–297. Reshetnikov, A. S., Khmelenina, V. N., and Trotsenko, Y. A. (2010). Identification of ectoine synthesis genes in a moderate halophilic alphaproteobacterium Methylarcula marina. Microbiology (Moscow) 79, 856–857. Reuter, K., Pittelkow, M., Bursy, J., Heine, A., Craan, T., and Bremer, E. (2010). Synthesis of 5-hydroxyectoine from ectoine: Crystal structure of the non-heme Iron(II) and 2-oxoglutarate-dependent dioxygenase EctD. PLoS One 5(5), e10647. Sauer, T., and Galinski, E. A. (1998). Bacterial milking: A novel bioprocess for production of compatible solutes. Biotechnol. Bioeng. 57, 306–313. Saum, S. H., and Muller, V. (2008). Growth phase-dependent switch in osmolyte strategy in a moderate halophile: Ectoine is a minor osmolyte but major stationary phase solute in Halobacillus halophilus. Environ. Microbiol. 10, 716–726. Schwibbert, K., Marin-Sanguino, A., Bagyan, I., Heidrich, G., Lentzen, G., Seitz, H., Rampp, M., Schuster, S. C., Klenk, H.-P., Pfeiffer, F., Oesterhelt, D., and Kunte, H. J. (2010). A blue print of ectoine metabolism from the genome of the industrial producer Halomonas elongata DSM2581T. Environ. Microbiol. 10.1111/j.14622920.2010.02336.x. Severin, J., Wohlfarth, A., and Galinski, E. A. (1992). The predominant role of recently discovered tetrahydropyrimidines for the osmoadaptation of halophilic eubacteria. J. Gen. Microbiol. 138, 1629–1638. Trotsenko, Y. A., and Khmelenina, V. N. (2002). Biology and osmoadaptation of haloalkaliphilic methanotrophs. Microbiology (Moscow) 71, 123–132. Wampler, D. E., and Westhead, E. W. (1968). Two aspartokinases from Escherichia coli. Nature of the inhibition and molecular changes accompanying reversible inactivation. Biochemistry 7, 1661–1671. Wilkinson, S. P., and Grove, A. (2006). Ligand-responsive transcriptional regulation by members of the MarR family of winged helix proteins. Curr. Issues. Mol. Biol. 8, 51–62.

C H A P T E R

T H R E E

Facultative and Obligate Methanotrophs: How to Identify and Differentiate Them Svetlana N. Dedysh* and Peter F. Dunfield† Contents 1. Introduction 2. Identification of Methanotrophic Capabilities in Novel Isolates 2.1. Culture conditions 2.2. Registration of growth dynamics on methane 2.3. Observation of intracytoplasmic membrane structures 2.4. Detection of genes encoding methane monooxygenase 3. Substrate Utilization Tests 4. Tests for Culture Purity 4.1. Plating on complex organic media 4.2. Phase-contrast and electron microscopy 4.3. Whole-cell hybridization with fluorescent probes 4.4. 16S rRNA gene clone library analysis 4.5. Dilution–extinction growth experiments 4.6. Quantification of methane monooxygenase-coding genes during growth on an alternate substrate References

32 33 33 34 35 35 37 38 38 39 39 41 42 42 43

Abstract Aerobic methanotrophs are metabolically unique bacteria that are able to utilize methane and some other C1-compounds as sole sources of carbon and energy. A defining characteristic of these organisms is the use of methane monooxygenase (MMO) enzymes to catalyze the oxidation of methane to methanol. For a long time, all methanotrophs were considered to be obligately methylotrophic, that is, unable to grow on compounds containing C–C bonds. This notion has recently been revised. Some members of the genera Methylocella, Methylocystis, and Methylocapsa are now known to be facultative methanotrophs, that is, capable of growing on methane as well as on some multicarbon substrates. The diagnosis of * Winogradsky Institute of Microbiology, Russian Academy of Sciences, Moscow, Russia Department of Biological Sciences, University of Calgary, Calgary, Alberta, Canada

{

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00003-6

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2011 Elsevier Inc. All rights reserved.

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Svetlana N. Dedysh and Peter F. Dunfield

facultative methanotrophy in new isolates requires a great degree of caution since methanotrophic cultures are frequently contaminated by heterotrophic bacteria that survive on metabolic by-products of methanotrophs. The presence of only a few satellite cells in a culture may lead to false conclusions regarding substrate utilization, and several early reports of facultative methanotrophy are likely attributable to impure cultures. Another recurring mistake is the misidentification of nonmethanotrophic facultative methylotrophs as facultative methanotrophs. This chapter was prepared as an aid to avoid both kinds of confusion when examining methanotrophic isolates.

1. Introduction At present, aerobic methanotrophic capabilities are recognized in members of two bacterial phyla, the Proteobacteria (the classes Alpha- and Gammaproteobacteria), and the Verrucomicrobia (reviewed in Op den Camp et al., 2009). Bacteria from the as yet unnamed candidate phylum NC10 also perform aerobic methanotrophy after forming free dioxygen from NO, but as yet little is known about these (Ettwig et al., 2010). Most aerobic methanotrophs grow only on methane, plus in some instances on methanol, formate, formaldehyde, and methylamines, and are therefore termed “obligate methanotrophs.” By contrast, facultative methanotrophs are able to use either methane or some multicarbon compound(s) as their sole carbon and energy source. The occurrence of facultative methanotrophy was recently demonstrated in several alphaproteobacterial methanotrophs of the genera Methylocella, Methylocystis, and Methylocapsa (Belova et al., 2011; Dedysh et al., 2005; Dunfield et al., 2010; Im et al., 2011). Members of these genera differ with regard to their physiology and substrate preferences. Methylocella species were the first to be conclusively shown to have a facultative capability. Among the known methanotrophs, these bacteria are very unique. They possess only a soluble methane monooxygenase (sMMO) and lack an extensive intracytoplasmic membrane (ICM) system common to all other methanotrophs. Perhaps not surprisingly given their unique status among methanotrophs, they can also utilize a number of multicarbon compounds (acetate, pyruvate, succinate, malate, and ethanol). These substrates are in fact preferred, and sMMO in Methylocella is repressed if an alternative multicarbon growth substrate is present (Theisen et al., 2005). Later, several facultatively methanotrophic Methylocystis spp. and Methylocapsa aurea were described. Unlike the oddball Methylocella, these are more typical aerobic methanotrophs, possessing a particulate methane monooxygenase enzyme (pMMO) and a well-developed ICM system in which pMMO is bound. The preferred growth substrate of these organisms is methane, but growth

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can also occur on acetate and/or ethanol in the absence of methane. Notably, it has been shown for a Methylocystis that pMMO is expressed in the presence of these alternate substrates, which can be utilized alone or together with methane. Thus, a facultative lifestyle may occur in both pMMO- and sMMOpossessing methanotrophs. It should be noted, however, that the existence of methanotrophs capable of growth on multicarbon compounds was a controversial topic for a long time prior to 2005. Periodic reports of facultative methanotrophs could not be independently verified (reviewed in Dedysh and Dunfield, 2010; Theisen and Murrell, 2005). The most common problems with these studies were: (1) lack of evidence for the genetic and enzymatic machinery for methane oxidation in a target organism and (2) questionable culture purity. In other words, either a nonmethanotrophic facultative methylotroph was erroneously identified as a facultative methanotroph or the study was conducted using a tight syntrophic association between a methanotroph and a facultative methylotroph. In this chapter, we describe our standard procedures for identifying methanotrophic capabilities in a novel isolate, for testing its ability to grow on multicarbon substrates, and for evaluating methanotroph culture purity.

2. Identification of Methanotrophic Capabilities in Novel Isolates 2.1. Culture conditions Both obligate and facultative methanotrophs are cultivated using liquid or solid mineral media with methane as a growth substrate. The use of multicarbon compounds for isolation or laboratory maintenance of facultative methanotrophs is not recommended since these substrates give a selective advantage to heterotrophic bacteria, which overgrow and contaminate methanotrophic cultures. Media that can be used for methanotroph cultivation include: A. Nitrate mineral salts (NMS) medium (Bowman, 2000; Whittenbury et al., 1970) is most widely used for methanotroph cultivation. It contains (g/l distilled water): KNO3, 1; MgSO4
7H2O, 1; Na2HPO4
12H2O, 0.717; KH2PO4, 0.272; CaCl2
6H2O, 0.2; ferric ammonium EDTA, 0.005. The medium pH is 6.8. Some strains prefer an ammonium mineral salts medium (AMS) in which the 1 g of KNO3 is replaced with 0.5 g NH4Cl. B. Dilute nitrate mineral salts (DNMS) medium is NMS medium diluted 1:5 with distilled H2O and containing 1 mM of NaH2PO4–Na2HPO4 buffer corresponding to a certain pH (5.5–7.0) (Dunfield et al., 2003). It is most suitable for methanotrophs from freshwater and salt-free terrestrial environments.

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C. Medium M2 (g/l distilled water) KNO3, 0.25; KH2PO4, 0.1; MgSO4
7H2O, 0.05; CaCl2
2H2O, 0.01; and NaCl, 0.02; pH is 5.5. This medium is suitable for methanotrophs from freshwater wetlands and mildly acidic soils. All these media are supplemented with 0.1% (by volume) of a trace elements stock solution. Several versions have been used. We recommend a mixture containing (in grams per litre) EDTA, 0.5; FeSO4
7H2O, 0.2; H3BO3, 0.03; ZnSO4
7H2O, 0.01; MnCl2
4H2O, 0.003; CoCl2
6H2O, 0.02; CuSO4
5H2O, 0.1; NiCl2
6H2O, 0.002, and Na2MoO4, 0.003. For plating, these media are solidified with agar (Difco) or gellan gum (Gel-Gro; ICN Biomedicals). These media are incubated in a closed vessels containing air supplemented with 10–30% (v/v) CH4 and 5% CO2. Growth under these conditions does not necessarily imply a methanotrophic phenotype in a particular bacterial strain. Growth may be supported by energy sources occurring as contaminants of the media components, on other trace hydrocarbons contained in the methane (especially if of low purity), on the polysaccharides used as gelling agents (agar or phytagel), or on secondary metabolites produced by a methanotroph growing in the medium. To verify a methanotrophic phenotype, growth experiments in liquid media are performed as described below.

2.2. Registration of growth dynamics on methane 1. For growth experiments, 100–500 ml screw-cap serum bottles are used with a headspace/liquid space ratio of 4:1. After inoculation, the bottles are sealed with butyl-rubber stoppers, and methane (10–20%, v/v) is added to the headspace by using a syringe and a sterile filter (0.22 mm). 2. Bottles are inoculated with cells of a target strain to achieve an initial OD410 of 0.01–0.03. Uninoculated controls of the medium used are included as blanks to control for methane leakage and sterility control, and inoculated medium with no added methane is also included. The experiment is carried out in triplicate. After inoculation, bottles are incubated on a rotary shaker (100–150 rpm) at an optimal growth temperature. 3. Gas samples and culture aliquots are taken once every 1–2 days for determination of methane concentrations and OD410 measurements. Methane concentration is measured using a gas chromatograph equipped with a flame ionization detector and a Porapak Q column (if unavailable, a thermal conductivity detector and a Molecular Sieve 5A column may be used). The optical density is measured on a spectrophotometer. Most pMMO-possessing methanotrophs show good growth in these experiments with OD410 reaching 0.8–1.5 within 3–6 days. By contrast, methanotrophs that possess only a soluble MMO may require up to 2–3 weeks until the cultures reach OD410 of 0.2–0.3. In both cases, an

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increase in OD410 is accompanied by a decline in CH4 mixing ratio of the headspace, while no growth is observed in the same medium in the absence of methane.

2.3. Observation of intracytoplasmic membrane structures All pMMO-possessing proteobacterial methanotrophs contain a welldeveloped ICM system. These membranes are arranged in stacks in cells of methanotrophic Gammaproteobacteria or are aligned parallel to cytoplasmic membrane in methanotrophic Alphaproteobacteria. These characteristic ICM are absent from cells of methanotrophs possessing only sMMO: that is, Methylocella spp. The latter methanotrophs instead contain a vesicular membrane system, which is connected to the cytoplasmic membrane. To reveal both kinds of these structures, thin sections are usually prepared using batch cultures grown to the mid- or late-exponential growth phases. Here are the main steps of this procedure: 1. Cells are collected by centrifugation and prefixed with 1.5% (w/v) glutaraldehyde in 0.05 M cacodylate buffer (pH 6.5) for 1 h at 4  C and then fixed in 1% (w/v) OsO4 in the same buffer for 4 h at 20  C. 2. Samples are dehydrated by successive passages through an ethanol series (50%, 70%, 80%, 96%, and 100% (v/v)) and are embedded in a Spurr epoxy resin. 3. Thin sections are cut on a microtome, mounted on copper grids covered with Formvar film, contrasted with 3% (w/v) uranyl acetate in 70% (v/v) ethanol for 30 min, and then stained with lead citrate (2.7%, w/v) at 20  C for 4–5 min. 4. The preparations are examined using an electron microscope.

2.4. Detection of genes encoding methane monooxygenase All aerobic methanotrophs contain one or both of two potential MMO enzymes. pMMO is encoded by a pmoCAB operon, while sMMO is encoded by a more complex set of genes including mmoXYBZDC. The pmoA and mmoX genes encoding the b-subunit of pMMO and the a-subunit of the sMMO hydroxylase, respectively, have been extensively used as functional markers for these bacteria, and large sequence databases are available for design of universal primers. Detection of either pmoA or mmoX in DNA extracted from a new isolate strongly implies that the bacterium is capable of methane oxidation. Currently, a wide variety of primers targeting pmoA and mmoX of different proteobacterial methanotroph groups is available (McDonald et al., 2008). Some are listed in Table 3.1. Here, we report three most useful protocols for the PCR-based screening of novel isolates for the presence of pmoA and mmoX genes:

Table 3.1 PCR primers used for amplification of pmoA and mmoX genes Primer sequence (50 –30 )

Primer

Target

A189f

pmoA, GGNGACTGGGACTTCTGG amoA

A682r mb661r

pmoA, GAASGCNGAGAAGAASGC amoA pmoA CCGGMGCAACGTCYTTACC

A650 882f

pmoA mmoX

1403r mmoX1f mmoX2r mmoX901r mmoXA-f mmoXB-r mmoXmc1

mmoX mmoX mmoX mmoX mmoX mmoX mmoX

mmoXmc2 mmoX mmoXmc3 mmoX

Target organisms

Reference

pMMO-possessing proteobacterial methanotrophs and some nitrifiers Same as above

Holmes et al. (1995)

pMMO-possessing proteobacterial methanotrophs TGGAAGCCATTCCTGCA Same as above GGCTCCAAGTTCAAGGTCGAGC sMMO-possessing methanotrophs TGGCACTCGTAGCGCTCCGGCTCG Same as above CGGTCCGCTGTGGAAGGGCATGAAGCGCGT Same as above GGCTCGACCTTGAACTTGGAGCCATACTCG Same as above ACCCAGCGGTTCCASGTYTTSACCCA Same as above ACCAAGGARCARTTCAAG Same as above TGGCACTCRTARCGCTC Same as above VCGYTCGCCCCARTCRTC sMMO-possessing Beijerinckiaceae methanotrophs VGTCGGGCAGAASGGCAC Same as above CCGGCSGCSCAGAAATAT Same as above

Holmes et al. (1995) Costello and Lidstrom (1999) Bourne et al. (2001) McDonald et al. (1995) McDonald et al. (1995) Miquez et al. (1997) Miquez et al. (1997) Shigematsu et al. (1999) Auman et al. (2000) Auman et al. (2000) Dunfield et al. (2010)

Dunfield et al. (2010) Dunfield et al. (2010)

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A. In most pMMO-possessing methanotrophs, a 525-bp fragment of pmoA gene can be amplified using the primers A189 þ A682 (Table 3.1) and PCR conditions described by Holmes et al. (1995): an initial denaturation step of 96  C for 4 min, followed by 30 cycles of 92  C for 1 min, 56  C for 1 min, 72  C for 45 s, and a final extension of 5 min at 72  C. Please note that these primers are not suitable for pMMO-possessing methanotrophic representatives of the Verrucomicrobia. B. For most sMMO-possessing methanotrophs, the best results in mmoX gene fragment (approx. 1230 bp) amplification can be obtained using the primers mmoXA (166f) þ mmoXD (1401r) and PCR conditions described by Auman et al. (2000): an initial denaturation step of 94  C for 30 s, followed by 30 cycles of 92  C for 1 min, 60  C for 1 min, 72  C for 1 min, and a final extension of 5 min at 72  C. However, this protocol may fail with some Methylocella-like organisms. If this is the case, try the approach listed below. C. The following primer combinations were proven useful for mmoX detection in some Beijerinckiaceae methanotrophs (Dunfield et al., 2010): mmoXA (166f) þ mmoXmc1 (1353r), mmoXA (166f) þ mmoXmc2 (1272r), or mmoXmc3 (786f) þ mmoXmc1 (1353r). The corresponding PCR protocol consists of 35 cycles of denaturation at 94  C for 1 min, primer annealing at 55  C for 1 min, and elongation at 72  C for 1 min with a final extension step of 7 min. Reaction products are then checked for size and purity on 1% agarose gels visualized by staining with ethidium bromide, sequenced, and the resulting nucleotide sequences are compared with those available in public databases. Care must be taken here as the mmoX gene is homologous to genes encoding other alkane monooxygenases, while pmoA is homologous to the amoA gene of ammonia oxidizers. Highest similarity to a known methanotrophic isolate is desired. In summary, observation of methanotroph-specific ultrastructures in cells of a novel isolate, demonstration of growth concurrent with a decline of added methane in closed vessels, and detection of pmoA or mmoX genes in DNA extracts give a solid basis for the identification of a bacterium as a methanotroph. The next step is testing its ability to utilize different C1 and multicarbon compounds as growth substrates.

3. Substrate Utilization Tests The use of multicarbon substrates by novel isolates is tested in the same way as described in Section 2.2 with the only difference that methane is replaced with one of the following compounds at a concentration of 0.05% (w/v): ethanol, mannitol, sorbitol, inositol, glucose, fructose, sucrose, arabinose, lactose, xylose, maltose, raffinose, ribose, galactose, acetate, citrate,

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oxalate, malate, pyruvate, and succinate. Growth is examined after 1 month of incubation and confirmed by comparison to negative controls (growth on the same liquid mineral medium without source of carbon) and positive controls (growth with methane). These experiments may yield two possible results: A. No growth is observed on any of the multicarbon compounds tested, while good growth occurs on methane. This confirms an obligate nature of the examined methanotroph. B. Growth occurs on methane as well as on some multicarbon substrates. This suggests a facultative methanotrophy in a novel isolate. Two different scenarios can be observed in this case: B1: Methane is the preferred growth substrate, which is typical for pMMO-possessing facultative methanotrophs. B2: The isolate grows best on a multicarbon substrate(s), which is a characteristic of Methylocella-like, sMMO-possessing facultative methanotrophs. If the result is “B,” you may either have a facultative methanotroph or a coculture of a methanotroph with another organism. In the latter case one should proceed with tests for culture purity. Any microbiologist should be familiar with such procedures, however, for methanotrophs we prefer those listed below.

4. Tests for Culture Purity 4.1. Plating on complex organic media This is a routine test for the presence of heterotrophic satellites in methanotrophic cultures. The following media can be used for this purpose: A. Standard undiluted and 10-fold-diluted Luria-Bertani agar (1.0% tryptone, 0.5% yeast extract, 1.0% NaCl), R2A, or Nutrient Agar (Difco). B. Standard undiluted and 10-fold-diluted NMS- or M2-agar media (see above) amended with 0.05% (w/v) glucose, fructose, or sucrose and 0.005% (w/v) yeast extract. As a control, the same agar mineral medium is used without any organic substrates. No growth should be observed on the complex organic media. In some cases, weak growth of the same low magnitude can be observed both on agar mineral media with individual sugars and on control plates lacking any organic substrates. However, some facultative methylotrophs may not develop on any of the above listed media and may escape detection. Therefore, this routine

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plating approach is a preliminary test, which is insufficient to prove the purity of a target methanotroph.

4.2. Phase-contrast and electron microscopy Microscopy techniques allow visual examination and comparison of cell morphology and ultrastructure in cultures grown on methane and on an alternative multicarbon substrate. Cells in both cultures are expected to exhibit essentially the same morphology. In many cases, however, the cells fed with different substrates may slightly differ in size (Fig. 3.1A and B). In pMMO-possessing methanotrophs, ICM structures are present in both methane and Cn-substrate-grown cultures, though in the latter these membranes are less extensive and more loosely organized (Fig. 3.1C and D).

4.3. Whole-cell hybridization with fluorescent probes Since a nonmethanotrophic satellite bacterium may display the same cell morphology as the methanotrophic partner in a syntrophic association, the observation of uniform cell morphology by phase-contrast microscopy is insufficient to guarantee culture purity. In our experience, the use of A

B

C

D

ICM

ICM

Figure 3.1 Phase-contrast micrographs (A, B) and electron micrographs of ultrathin sections (C, D) of cells of pMMO-possessing facultative methanotroph Methylocystis sp. strain H2s. Cells grown on methane (A) are slightly larger than those grown on acetate (B); bar, 5 mm. Intracytoplasmic membranes (ICM) are present both in cells grown on methane (C) and in cells collected after three transfers on acetate (D); bar, 0.5 mm.

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whole-cell hybridization with 16S rRNA-targeted, fluorescently labeled oligonucleotide probes is a highly effective way to examine for contamination. This test is performed with methane- and acetate-grown cultures (or any other alternative substrate one desires to test). For each cell preparation, two probes labeled with different fluorescent dyes are applied. For example, a species-specific probe labeled with indocarbocyanine dye (Cy3) can be combined with a group-specific (genus-, type I or type II methanotroph-specific) probe labeled with 5(6)-carboxyfluorescein-N-hydroxysuccinimide ester (FLUOS). Observation of all cells being stained with both fluorescent probes is a strong argument for culture purity. The example of this method application is shown in Fig. 3.2. 1. Cells growing on the same mineral medium with methane or acetate as a growth substrate are harvested in the logarithmic phase by centrifugation and resuspended in 0.5 ml of phosphate-buffered saline (PBS) (g/l: NaCl, 8.0; KCl, 0.2; Na2HPO4, 1.44; NaH2PO4, 0.2; pH 7.0). 2. Cell suspensions are mixed with 1.5 ml of 4% (w/v) freshly prepared paraformaldehyde solution and fixed for 1 h at room temperature. The cells are then collected by centrifugation (6600  g for 1 min) and washed twice with PBS to ensure removal of paraformaldehyde. The resulting pellet is resuspended in 0.5 ml of 50% ethanol-PBS (v/v). 3. Hybridization is performed on Teflon-coated slides rinsed with 70% ethanol and dried. Ideally, slides have 6–8 wells for independent positioning of the samples. One to two microliters of the fixed cell suspension is spread on each well, air-dried, and dehydrated by successive passages through an ethanol series (50%, 80%, and 100% (v/v)) for 3 min each.

Figure 3.2 Whole-cell hybridization in a culture of a facultative methanotroph Methylocella silvestris BL2 grown on acetate as the sole carbon and energy source. Left panel: phase contrast; middle panel: hybridized with the Methylocella genus-specific probe Mcell-1445; right panel: hybridized with the Methylocella silvestris species-specific probe Mcells-1024. The scale bar represents 10 mm. All cells seen in phase contrast hybridized with both probes, indicating that the culture is pure.

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4. A 50-ml polypropylene screw-top Falcon tube containing a slip of Whatman filter paper soaked in hybridization buffer is used as a hybridization chamber as described by Stahl and Amann (1991). The chamber is allowed to equilibrate for at least 30 min at the hybridization temperature. 5. A 9-ml aliquot of hybridization buffer (0.9 M NaCl, 20 mM Tris/HCl, pH 7.2, 0.01% SDS, and formamide concentration corresponding to the probe stringency conditions) is placed on each spot of fixed cells. The slide is transferred to the equilibrated chamber and prehybridized for 30 min. Following prehybridization, 1 ml of fluorescent probe (50 ng ml 1 solution in double-distilled water) is added to each spot and the slide is returned to the hybridization chamber for 1–1.5 h. 6. Slides are washed at the hybridization temperature for 10 min in washing buffer (20 mM Tris/HCl, 0.01% SDS, and NaCl concentration corresponding to the probe stringency conditions), then rinsed with twicedistilled water and air-dried. 7. Each well of the slide is mounted with a drop of Citifluor AF1 antifadent (or with glycerol if Citifluor is missing), covered with a coverslip and viewed with an epifluorescent microscope equipped with the filters for Cy3- and FLUOS-labeled probes.

4.4. 16S rRNA gene clone library analysis This approach is used to prove the homogeneity of isolated methanotrophic strains and to demonstrate the phylogenetic identity of the cultures grown on methane and on alternative multicarbon substrates. 1. A single colony of the target strain is used to inoculate a liquid culture. This culture is grown on methane to the mid-exponential phase. Then cells are collected by centrifugation and divided into two parts, one of which is again provided with methane, while the other grows on acetate (or another multicarbon substrate). 2. After 7–10 days of incubation, the cells are harvested, and genomic DNA is extracted using a mechanical cell disruption procedure. 3. PCR-mediated amplification of 16S rRNA genes is performed with a primer set useful for most members of the domain Bacteria, for example, 9f and 1492r of Weisburg et al. (1991). 4. PCR-amplified 16S rRNA gene products are cloned into E. coli using any of the commercially available cloning kits. Thirty to fifty clones from each of the two constructed 16S rRNA gene clone libraries (from methaneand acetate-grown cells) are randomly selected for the examination by means of (a) sequencing of at least 500–600 bp from the 50 end of the 16S rRNA gene inserts or (b) restriction fragment length polymorphism analysis using digestion by two sets of tetrameric endonucleases (e.g., MspIþRsaI and HhaIþHaeIII, as described by Dedysh et al., 2000).

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Identity of the resulting nucleotide sequences or the respective restriction patterns of the cloned 16S rRNA gene fragments in both clone libraries indicate homogeneity of the target culture.

4.5. Dilution–extinction growth experiments The experiments described in Sections 4.3 and 4.4 may fail to reveal a contaminating heterotrophic bacterium if the latter is numerically inferior to a methanotrophic organism (<1 in 100). A pyrosequencing procedure could be used to obtain a larger number of sequences. Alternatively, to rule out this possibility, a simple dilution growth experiment can be made as follows: 1. Serial (10-fold) dilutions for most-probable-number (MPN) counts are prepared using quadruplicate 20-ml test tubes containing 5 ml of medium. The source culture for the MPN should be grown through multiple transfers on acetate (or other desired substrate) alone. One set of MPN tubes contains acetate for the enumeration of acetate-utilizing bacteria, while another set of tubes is incubated in closed glass desiccators under a headspace containing 10% (v/v) CH4 for the enumeration of methane-utilizing bacteria. 2. Tubes are incubated for 1 month and MPNs are calculated using an Excel-based program (Briones and Reichardt, 1999). If pure, the examined culture should contain equal numbers of methaneutilizing and acetate-utilizing cells, even after multiple transfers of this culture on acetate alone. The same experiment can be made with a source culture grown through multiple transfers on methane.

4.6. Quantification of methane monooxygenase-coding genes during growth on an alternate substrate The increase of methanotroph-specific mmoX or pmoA genes during growth on an alternate substrate such as acetate clearly demonstrates that methanotrophic cells and their genomes are multiplying on a substrate other than methane. 1. Substrate utilization tests should be performed as in Section 3. Periodically samples should be taken for measurement of optical density or direct cell counts. 2. Genomic DNA is extracted from each sample using a mechanical cell disruption procedure. 3. qPCR is performed using a Sybr-Green based method (A TaqMan approach is also feasible). A series of pmoA and mmoX primers targeting

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various groups of methanotrophs have been designed for use in qPCR assays by Kolb et al. (2003). 4. qPCR should be performed to target the 16S rRNA gene and the respective pmoA or mmoX gene of the isolate (Kolb et al., 2003). Variations in extraction efficiency and copy numbers may mean that the resulting gene counts are not equal, however, the growth curves measured via OD or direct cell counting should closely parallel the increase in copy numbers of the 16S rRNA genes and methanotrophspecific functional genes.

REFERENCES Auman, A. J., Stolyar, S., Costello, A. M., and Lidstrom, M. E. (2000). Molecular characterization of methanotrophic isolates from freshwater lake sediment. Appl. Environ. Microbiol. 66, 5259–5266. Belova, S. E., Baani, M., Suzina, N. E., Bodelier, P. L. E., Liesack, W., Dunfield, P. F., and Dedysh, S. N. (2011). Acetate utilization as a survival strategy of peat-inhabiting Methylocystis spp. Environ. Microbiol. Reports 3, 36–46. Bourne, D. G., McDonald, I. R., and Murrell, J. C. (2001). Comparison of pmoA PCR primer sets as tools for investigating methanotroph diversity in three Danish soils. Appl. Environ. Microbiol. 67, 3802–3809. Bowman, J. (2000). The methanotrophs—The families Methylococcaceae and Methylocystaceae. release 3.1. In “The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community,” (M. Dworkin, et al., eds.), 3rd edn. http://link.springer-ny.com/ link/service/books/10125/. Briones, J. A. M., and Reichardt, W. (1999). Estimating microbial population counts by ‘most probable number’ using Microsoft Excel. J. Microbiol. Meth. 35, 157–161. Costello, A., and Lidstrom, M. E. (1999). Molecular characterization of functional and phylogenetic genes from natural populations of methanotrophs in lake sediments. Appl. Environ. Microbiol. 65, 5066–5074. Dedysh, S. N., and Dunfield, P. F. (2010). Facultative methane oxidizers. In “Handbook of Hydrocarbon and Lipid Microbiology,” (K. N. Timmis, ed.), pp. 1967–1976. SpringerVerlag, Berlin. Dedysh, S. N., Liesack, W., Khmelenina, V. N., Suzina, N. E., Trotsenko, Y. A., Semrau, J. D., Abing, A. M., Panikov, N. S., and Tiedje, J. M. (2000). Methylocella palustris gen. nov., sp. nov., a new methane-oxidizing acidophilic bacterium from peat bogs, representing a novel subtype of serine-pathway methanotrophs. Int. J. Syst. Evol. Microbiol. 50, 955–969. Dedysh, S. N., Knief, C., and Dunfield, P. F. (2005). Methylocella species are facultatively methanotrophic. J. Bacteriol. 187, 4665–4670. Dunfield, P. F., Khmelenina, V. N., Suzina, N. E., Trotsenko, Y. A., and Dedysh, S. N. (2003). Methylocella silvestris sp. nov. a novel methanotrophic bacterium isolated from an acidic forest cambisol. Int. J. Syst. Evol. Microbiol. 53, 1231–1239. Dunfield, P. F., Belova, S. E., Vorob’ev, A. V., Cornish, S. L., and Dedysh, S. N. (2010). Methylocapsa aurea sp. nov., a facultatively methanotrophic bacterium possessing a particulate methane monooxygenase. Int. J. Syst. Evol. Microbiol. 60, 2659–2664.

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Ettwig, K. F., Butler, M. K., Le Paslier, D., Pelletier, E., Mangenot, S., Kuypers, M. M. M., Schreiber, F., Dutilh, B. E., Zedelius, J., de Beer, D., Gloerich, J., Wessels, H. J. C. T., et al. (2010). Nitrite-driven anaerobic methane oxidation by oxygenic bacteria. Nature 464, 543–549. Holmes, A. J., Costello, A., Lidstrom, M. E., and Murrell, J. C. (1995). Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132, 203–208. Im, J., Lee, S.-W., Yoon, S., Dispirito, A. A., and Semrau, J. D. (2011). Characterization of a novel facultative Methylocystis species capable of growth on methane, acetate and ethanol. Environ. Microbiol. Reports 10.1111/j.1758-2229.2010.00204.x (in press). Kolb, S., Knief, C., Stubner, S., and Conrad, R. (2003). Quantitative detection of methanotrophs in soil by novel pmoA-targeted real-time PCR Assays. Appl. Environ. Microbiol. 69, 2423–2429. McDonald, I. R., Kenna, E. M., and Murrell, J. C. (1995). Detection of methanotrophic bacteria in environmental samples with the PCR. Appl. Environ. Microbiol. 61, 116–121. McDonald, I. R., Bodrossy, L., Chen, Y., and Murrell, J. C. (2008). Molecular ecology techniques for the study of aerobic methanotrophs. Appl. Environ. Microbiol. 74, 1305–1315. Miquez, C. B., Bourque, D., Sealy, J. A., Greer, C. W., and Groleau, D. (1997). Detection and isolation of methanotrophic bacteria possessing soluble methane monooxygenase (sMMO) genes using the polymerase chain reaction (PCR). Microbial Ecol. 33, 21–31. Op den Camp, H. J. M., Islam, T., Stott, M. B., Harhangi, H. R., Hynes, A., Schouten, S., Jetten, M. S. M., Birkeland, N. K., Pol, A., and Dunfield, P. F. (2009). Minireview: Environmental, genomic, and taxonomic perspectives on methanotrophic Verrucomicrobia. Environ. Microbiol. Reports 1, 293–306. Shigematsu, T., Hanada, S., Eguchi, M., Kamagata, Y., Kanagawa, T., and Kurane, R. (1999). Soluble methane monooxygenase gene clusters fron trichloroethylene-degrading Methylomonas sp. strains and detection of methanotrophs during in situ bioremediation. Appl. Environ. Microbiol. 65, 5198–5206. Stahl, D. A., and Amann, R. (1991). Development and application of nucleic acid probes. In “Nucleic Acid Techniques in Bacterial Systematics,” (E. Stackebrandt and M. Goodfellow, eds.), pp. 205–248. Wiley, New York, NY. Theisen, A. R., and Murrell, J. C. (2005). Facultative methanotrophs revisited. J. Bacteriol. 187, 4303–4305. Theisen, A. R., Ali, M. H., Radajewski, S., Dumont, M. G., Dunfield, P. F., McDonald, I. R., Dedysh, S. N., Miguez, C. B., and Murrell, J. C. (2005). Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol. Microbiol. 58, 682–692. Weisburg, W. G., Barns, S. M., Pelletier, D. A., and Lane, D. J. (1991). 16S ribosomal DNA amplification for phylogenetic study. J. Bacteriol. 173, 697–703. Whittenbury, R., Phillips, K. C., and Wilkinson, J. F. (1970). Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61, 205–218.

C H A P T E R

F O U R

Approaches for the Characterization and Description of Novel Methanotrophic Bacteria John P. Bowman Contents 1. Introduction to Aerobic Methanotroph Characterization for Systematic Research and Other Purposes 2. Broad Level Systematic Diversity 2.1. Environmental relevance and knowledge limitations 2.2. Requirements related to the Bacteriological Code 3. Methodological Approaches 3.1. Enrichment and isolation 3.2. Minimal standards for characterization 3.3. Chemotaxonomy 3.4. Informative sequence-based analyses and genomic comparisons 3.5. Culture collections 4. Concluding Remarks References

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Abstract Methanotrophs are a key group of environmental microorganisms that play an integral role in the global cycling of methane. Modern culture-independent techniques, traditional painstaking culture methods, and genomics-related approaches continue to reveal new information about methanotrophs, including their ecosystem associations, biochemistry, and their systematic and evolutionary nature. The increasing interest in methanotrophic bacteria especially in the context of climate change will likely lead to an increase in cultures available for in-depth studies. This chapter details the rigorous approach needed for successful appraisal and confirmation of novel aerobic methanotrophic bacterial species that can then be made accessible for in vitro research purposes. This includes a guide in which descriptions maintain adherence to the Bacteriological Code; that Tasmanian Institute of Agricultural Research, University of Tasmania, Hobart, Tasmania, Australia Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00004-8

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2011 Elsevier Inc. All rights reserved.

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meet minimal standards; and the mandatory deposition of representative strains in approved culture collections.

1. Introduction to Aerobic Methanotroph Characterization for Systematic Research and Other Purposes Aerobic methanotrophs represents a metabolically allied collection of bacteria that possess the ability to grow on methane as a sole carbon and energy source. To this current point in time, methanotrophy is carried out by specialist bacteria, which have largely dispensed with multicarbon compound catabolism. All known methanotrophs can utilize the metabolic products generated from methane oxidation (methanol, formaldehyde) and sometimes other C1 compounds. Claims of facultative methanotrophy, in which strains have been shown to be able to utilize methane as well as multicarbon compounds, has been controversial (Dedysh et al., 2004) but far from being unprecedented (Dunfield et al., 2010; Stoecker et al., 2006). Methanotrophy that occurs anaerobically is carried out by consortia of specific methanogenic archaea and sulfate-reducing bacteria (Knittel and Boetius, 2009). Since this process is ecologically and biochemically distinct from aerobic methanotrophy it will not be dealt with in this chapter. Furthermore, the syntrophic nature of anaerobic methanotrophy hinders the systematic understanding of the associated microorganisms performing the process and at this stage much research is still being focused on better understanding the biochemical and ecological nature of the process itself.

2. Broad Level Systematic Diversity The aerobic methanotrophic process on the other hand is much better understood and is based on decades of research involving pure cultures, including model strains, such as Methylococcus capsulatus BATH (Ward et al., 2004), coupled to environmental in situ studies. Aerobic methanotrophy is possible due to possession of methane monooxygenase (MMO; EC 1.14.13.25) an oxidoreductase enzyme that comes in two major varieties (Baik et al., 2003). Most cultured bacteria that possess MMO belong to the phylum Proteobacteria and are thus Gram-negative. Most are motile by flagella. No cultured strains have been shown, conclusively at least, to be capable of autotrophic growth using CO2 for carbon or able to use an alternative energy sources such as H2 or CO; however, systematic studies to date tend to follow a well-trodden path in terms of test methods (see Section 3). Recently, extremely acidophilic and mildly thermophilic

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methanotrophs were identified that belong to the phylum Verrucomicrobia (Dunfield et al., 2007; Islam et al., 2008; Pol et al., 2007). As of late 2010, none of these verrucomicrobial isolates were validly described though they have been provisionally proposed as belonging to a genus designated “Methylacidiphilium” (Op den Camp et al., 2009). Methanotrophs belonging to the phylum Proteobacteria are grouped into families that are generally monophyletic (clustered together) or represent closely related paraphyletic (distinct but adjacent) lineages. These distinct groupings suggest methanotrophy arose multiple times following the major divergence of bacteria from archaea (Zerkle et al., 2005) and possibly coevolved with ammonia oxidizers owing to the similarity in ammonia monooxygenase and the copper-containing particulate MMO (pMMO). The members of each of the proteobacterial families possess distinct phenotypic differences and thus the methanotrophs are often termed in generalist subgroupings as type I, type II (and occasionally in the case of Methylococcus and its immediate relatives, “type X”) methanotrophs. Type I methanotrophs include those belonging to the class Gammaproteobacteria and include psychrophilic to mildly thermophilic species and are grouped together in the family “Methylococcaceae” represented by genus Methylomonas and its relatives. Methanotrophs with a preference for mildly thermal environments form a cluster deeper in class Gammaproteobacteria and include genus Methylococcus and Methylocaldum and represent effectively family Methylococcaceae sensu stricto, since Methylococcus is the type genus. The description of a multitude of new species has resulted in the originally described family Methylococcaceae (Bowman et al., 1995) becoming increasingly less monophyletic; however, it is now well recognized that cultured methanotroph diversity is only a fraction of what can be observed using PCR-based and array-based culture-independent techniques (Kip et al., 2010). Type II methanotrophs occur in the closely adjacent families Methylocystaceae and Beijerinckaceae, which belong to the class Alphaproteobacteria. The family Beijerinckiaceae incorporate non-methanotrophic genera (i.e., Beijerinckia, Rhodoblastus); however, recent evidence suggests most of these probably are methylotrophic (Dedysh et al., 2005; Kulichevskaya et al., 2006). The type II methanotrophs are also closely related to the genus Methylobacterium commonly isolated pink-pigmented, methylotrophs able to utilize C1 and multicarbon compounds. It is thus recognized the trait of methylotrophy, especially methanol utilization, is highly relevant to methanotrophs. All methanotrophic genera have traditionally been named with the genus prefix “Methylo-”; however, an exception is Crenothrix polyspora. This microbe is very much a distinction since it suggests that methanotrophs have evolved not only surprising complexity in terms of life cycle and growth strategies but also unsuspected metabolic flexibility, including

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utilization of alternative energy sources. As perhaps the best example of a microorganism that does not follow the standard “methanotroph mold,” C. polyspora is a sheathed, filamentous, iron-accumulating bacterium that has been shown to have a rather distinctive pMMO; forms unusual hexagonal and fibrillar-like inclusion bodies of unknown function; a life cycle and morphology not unlike various cyanobacteria, that is, formation of gonidia (Vo¨lker et al., 1977); and based on radio-labeling studies is able to take up multicarbon substrates (Stoecker et al., 2006). Further (systematic) studies may yet reveal that methanotrophs are diverse, sophisticated, and highly networked microorganisms within the Earth’s biosphere.

2.1. Environmental relevance and knowledge limitations Ecologically aerobic methanotrophs exist at positions in which oxygen and methane is available. Methane is mainly derived from methanogenesis but also occurs due to abiotic processes (i.e., volcanic, extraterrestrial). The densest populations of aerobic methanotrophs overly anoxic zones in which methane is emitted and enters aerobic zones. Such locations include mainly freshwater aquatic environments and soil (Chen and Murrell, 2010). In marine zones, aerobic methanotrophy may occur if the environment is sulfate depleted or where active natural gas seeps or accumulated methane clathrates occur, in general, anaerobic methane oxidation may be otherwise more important (Treude and Ziebis, 2010). Little information is available on marine aerobic methanotrophs and is thus fertile ground for study. In soil ecosystems, it is also noted that indigenous methanotrophs may have high affinity MMO and thus are able to utilize atmospheric methane (currently at about 1.8 ppm). No examples of these “high affinity” methanotrophs are available in pure culture; indeed, it is uncertain whether they exist at all and rather cultured methanotrophs behave differently under conditions of low methane relying on other forms of exogenous carbon (Degelmann et al., 2010). Indeed, the general practice is to always grow methanotrophs under highly methane excess conditions; thus understanding of the full metabolic potential of methanotrophs is currently poorly developed.

2.2. Requirements related to the Bacteriological Code Microbial taxonomy and the nomenclatural framework it rests upon in recent years have increasingly become better defined. As a primer, it is strongly suggested the reader consult the paper “Notes on the characterization of prokaryote strains for taxonomic purposes” (Tindall et al., 2010), which spells out in detail the process and information content expected for the quality expected in novel species descriptions. An additional critical criterion that must be emphasized is strain availability. For a species type strain to be validly described it must be housed in at least two internationally

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recognized culture collections (Ka¨mpfer, 2010). Culture collection recognition is conducted by the Word Federation of Culture Collections ( www. wfcc.info); however, the extant nature of the collection is not verified and thus methanotroph deposition should be performed using only collections that have a reliable history and the associated expertise (see Section 3.5 for more information). Other strains can be described for research purposes; however, only by fulfilling the strict criteria adopted by the International Committee on Systematic Bacteriology (www.the-icsp.org) and the Bacteriological Code (Lapage et al., 1992; freely available at http://www.ncbi.nlm.nih. gov/bookshelf/br.fcgi?book¼icnb) can they be recognized as taxonomically official. A full list of official names is maintained at the LPSN (List of Prokaryotic names with Standing in Nomenclature) website at www.bacterio. cict.fr. The LPSN web resource is an excellent information portal for taxonomic names and their status, type strain designations, culture collections where type strains are found; and verified 16S rRNA sequence data derived from type strains (Pruesse et al., 2007; www.arb-silva.de). The actual process through which the appropriate and exacting isolation and characterization of methanotrophs occurs by is very important, owing to special problems that are a feature of this particular type of bacteria. For successful taxonomic delineation of methanotroph isolation a set of minimal requirements is clearly required.

3. Methodological Approaches The processes by which the taxonomy and thus the essential biological and biochemical features of methanotrophs are obtained are described below in a series of distinct stages: (3.1) enrichment and isolation; (3.2) essential identification and baseline characterization; (3.3) chemotaxonomy; (3.4) sequence analysis; and (3.5) long-term preservation.

3.1. Enrichment and isolation Enrichment is usually required to isolate methanotrophs though direct plating of diluted samples onto agar can also be used where they occur in abundance. The latter applies typically to freshwater sediment and some soils. Primary selection medium is traditionally nitrate mineral salts (NMS) formulated originally by Roger Whittenbury and colleagues (Bowman, 2006; Whittenbury et al., 1970). Ammonia salts are not used as they may competitively inhibit MMO. Methane must be of high purity since as even traces of acetylene will permanently inactivate MMO. The NMS medium can be modified to suit the environments being sampled. For isolation of methanotrophs from saline systems the NMS salt content should be adjusted

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to match the salinity profile of the source samples (Bowman, 2006). Methanotrophs from freshwater and soil samples are usually rather salt-sensitive. For acidic soils the medium has been modified by diluting the medium 1:5 and pH adjusted to approximately pH 4.5–6.0 using a phosphate buffer (Dunfield et al., 2007). Typically for acidic soils soil particles are placed directly on agar plates (Dunfield et al., 2003). Obviously empirical media modification could reveal novel methanotrophs and fundamentally the initial conditions should match the environment being sampled. For enrichments bottles or tubes that are crimped sealed with butyl rubber stoppers are ideal. Agar plates can be incubated in desiccators or anaerobic jars. The atmospheres of the vessels are adjusted by removal of 5–50% of the headspace and replacing with methane. For routine cultivation, the standard level of methane most often used is a 1:4 CH4:air ratio. Since all known pure cultures have a relatively low affinity MMO (be it pMMO or soluble MMO) the actual concentration of the methane atmosphere is not critical except perhaps from a safety point of view. Incubation temperatures should reflect the source environment and can potentially range from 4  C for tundra and Antarctic samples to ca. 60  C for thermal springs and aquifers. The given optimal growth temperature of isolates can be tested empirically. An ideal means to do biokinetic temperature data accurately is described by Ratkowsky et al. (1983). Most polar species described to date have temperature growth ranges typical of “psychrotrophs” (4 to 30–37  C) or psychrophiles (0–22  C) (Wartiainen et al., 2006). The latter are often notoriously slow growing (doubling times 1 day) (Bowman et al., 1997). Most known methanotrophs are mesophiles growing from 10 to 40  C (optimum ca. 30  C) while the species of genus Methylococcus and Methylocaldum grow from 20 to 55  C (optimum ca. 45  C). Species evolved for acidic or alkaline environments do occur (Kalyuzhnaya et al., 2008). Type I methanotrophs include several examples of halophilic strains including salt and soda lakes, coastal and estuarine species (e.g., Methylomicrobium pelagica, Methylohalobius crimeensis; Heyer et al., 2005; Kalyuzhnaya et al., 2008). The general lack of descriptions from more saline water bodies suggests large untapped methanotrophic diversity in many different aquatic ecosystems that could be readily accessible through standard cultivation approaches. The analysis of geochemical signatures could be a very useful first step in exploring sites for the presence of active aerobic methanotrophy. For example the presence of signature fatty acids (see Section 3.3) and aminobacteriohopanepolyols (Zhu et al., 2010) may be useful as a guide for sample selection for subsequent isolation and characterization studies. Agar media can be prepared with normal agar. Agarose or noble agar as a solidifying agent may be ideal during the purification phases though not a requirement. It should be noted some methanotrophs do not grow or do not maintain stable growth on agar-solidified media possibly due to intolerance to the agar (Bowman et al., 1997; Rahalkar et al., 2007). In these cases, liquid

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culture is required or silica gel-solidified media (Bazylinski and Rosenberg, 1980); however, the latter is potentially technically difficult to prepare and has been used infrequently for methanotrophs. Purification of agarophobic methanotrophs requires repeated dilution to extinction in liquid media (Bowman et al., 1997) and so far has rarely been done due to technical difficulties and some element of luck in obtaining pure cultures. The utilization of highthroughput culturing methods (Connon and Giovannoni, 2002; Toledo et al., 2006) has yet to be exploited significantly for methanotroph cultivation and could be ideal approaches for isolation of novel methanotrophs. Colonies that subsequently appear on primary isolation agar plates are without exception contaminated with other non-methanotrophic bacteria. The main culprits are non-methanotrophic methylotrophs and oligotrophic, heterotrophic bacteria such as Brevundimonas, Caulobacter, etc. To obtain pure cultures the most effective method is to add a small amount of readily useable carbon to the medium (typically 0.1% glucose or 0.05% yeast extract) and then to dilute cell suspensions to extinction. The extra carbon stimulates the growth of the contaminants to form distinct colonies otherwise they grow by cross-feeding on metabolites from the methanotrophs and thus cannot be separated from primary growth. Plates need to be incubated for extended periods of time and thus care must be made against fungal contamination. Addition of cycloheximide, nystatin, or natamycin is advisable. Containers holding plates need to be cleaned regularly with ethanol and disinfectant to avoid fungal contamination. Final checks of purity require microscopic examination of cells and colonies for morphological consistency. This process may require repetition before pure cultures can be obtained, if at all. Validation of purity must be performed by plating cultures onto rich media such as R2A agar or 0.1  strength trypticase soy agar (both available from Oxoid and Difco Laboratories). Very rich media such as nutrient agar or full-strength trypticase soy agar are not ideal since oligotrophic contaminants may not grow on these media. The final confirmation of purity requires extra rigor for claims of facultative methanotrophy to be proven owing to past issues with culture purity. This can be achieved by denaturing gradient gel electrophoresis of 16S rRNA PCR products in which case only one PCR product should be observable; and sequence analysis of 16S rRNA, which should reveal a chromatogram with only a low background signal. Confirmation of purity of verrucomicrobial methanotrophs required direct genome sequencing (Pol et al., 2007). In some cases, methanotroph growth is stimulated by the presence of exogenous multicarbon compounds (such as small amounts of tryptone or yeast extract); however, there is a perpetual threat of culture contamination thus it is ideal for cultures to be maintained under methane only conditions. Finally, vitamin requirements are not unprecedented for methanotrophs and should be considered in any enrichment and isolation process though they may potentially complicate purification of strains.

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3.2. Minimal standards for characterization Standard tests for methanotrophs are summarized in Table 4.1 and are linked to special features possessed by certain methanotrophs and thus have known discriminating power. A species description of high quality would involve thorough investigation of all properties listed in Table 4.1 and would ideally involve comparison of multiple strains against suitable reference cultures; however, the requirement to have multiple strains is not obligatory. Popular commercial test strips such as API (Biome´rieux-Vitek) and Biolog are generally not utilized in testing methanotrophs. However, certain salient properties such as enzymes, and responses to nitrogen and phosphate sources as well as response to exogenous carbon sources have not to this point in time been given much attention. Opportunities thus exist to utilize relatively new testing platforms such as the Phenotype MicroarrayTM system from Biolog (http://www.biolog.com/pmMicrobialCells.html) to test for novel properties of both new and old methanotrophic strains. These would advance the taxonomy as well as general physiological understanding of methanotrophs. Fundamentally, the essential basics for validation of methanotrophs, especially if a claim for facultative use of methane is argued includes (a) determination of cellular morphology and importantly intracytoplasmic membrane ultrastructure; (b) determination of the predominant pathway for formaldehyde fixation; (c) means of methane oxidation (i.e., pMMO and/or sMMO); (d) basic cellular and colonial nutritional and physiological properties; (e) chemotaxonomy (see Section 3.3); and (f) 16S rRNA, pmoA, and nifH gene sequence analysis. Many descriptions to date are based on single isolates thus the inherent variation in traits at the species level can generally only be assessed at the genus level.

3.3. Chemotaxonomy Guanosine–cytosine (GþC) ratios of DNA and fatty acid profile are mandatory for novel descriptions of methanotrophs as these data are considered a useful means of differentiation not only of the broad types of methanotrophs but also of individual genera. GþC ratio analysis is now popularly analyzed by HPLC analysis and is considered a rapid, routine analysis procedure (Mesbah et al., 1989). The presence of distinct fatty acids by aerobic methanotrophs are taxonomically useful in distinguishing genera and/or species and has also been adopted as signatures of their presence in environmental samples (Bodelier et al., 2009; Bowman et al., 1991) though limitations to this technique are now clear due to some methanotrophs apparently lacking any designated signature lipids (e.g., Methylovulum miyakonense; Iguchi et al., 2010). Also it has been found that some type II methanotrophs may possess signature fatty acids typically found in type I methanotrophs (Dedysh et al., 2007). Most studies utilize the Sherlock Microbial Identification System

Table 4.1 A list of characteristics designated as minimal standards for characterization of novel aerobic methanotrophs Characteristicsa

Morphology and ultrastructure Cell shape, cell division, capsules Mode of motility Resting cells—type and relation to survival of desiccating conditions and heating Cell wall stability based on lysis in 2% sodium dodecyl sulfate solution Intracytoplasmic membrane (ICM) presence and morphology Spinae (extensions of the S-layer) Intracellular granules—polyhydroxyalkanoate (PHA) and polyphosphate Pigmentation Physiology and biochemistry pH range; NaCl tolerance; temperature range for growth Naphthalene oxidation assay for sMMO in the presence and absence of copper Formaldehyde fixation pathway

Special features that occur in some methanotrophs

Rosettes and budding cell division (Methylosinus spp., some Methylocella spp.); sarcinal packets (Methylosarcina spp.) Polar tufts (Methylosinus spp.) or polar flagella otherwise Cysts (most type I methanotrophs; Methylocystsis spp.); exospores (Methylosinus spp.) Type II methanotrophs are more stable and resist lysis in 2% SDS Absent in Methylocella spp. have a distinct ICM arrangement compared to other methanotrophs Formed by some Methylocystis spp. (can be up to 150 nm) Large PHA granules may occur depending on the strain (possibly can be confused with cysts) some strains may have distinct pigments though it is uncommon Methylocella and Methylocapsa are acidophilic; various type I methanotrophs show preferences for different pH, salinity, and/or temperature Methylocella spp. only possess sMMO

(continued)

Table 4.1 (continued) Characteristicsa

Special features that occur in some methanotrophs

Ribulose monophosphate pathway (type I methanotrophs); Serine pathway (type II methanotrophs) Catalase, cytochrome c oxidase Completeness of the TCA cycle; presence of glyoxylate cycle, glycolysis, and pentose phosphate pathway key enzymes

2-Oxoglutarate dehydrogenase may be absent in some type II methanotrophs

Other enzymatic activity—phosphatases, nitrate reduction, urease Carbon/nitrogen source tests Many strains can use a range of amino acids as nitrogen sources Nitrogen fixation – resting cell acetylene reduction assay; nifH gene sequencing Sequence analysis pmoA, 16S rRNA sequencing a

May vary between strains of the same species

Specific methods for characteristics can be found in the recent methanotroph taxonomic literature: Dedysh et al. (2007), Dunfield et al. (2003, 2010), Heyer et al. (2005), Kalayuzhnaya et al. (2008), and Rahalkar et al. (2007). More general guidelines for taxonomic descriptions and the standard and quality needed for such data is described in Tindall et al. (2010).

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(MIDI Inc., http://www.midi-inc.com) and several laboratories offering paid MIDI analysis services (e.g., DSMZ—German Collection of Microorganisms; http://www.dsmz.de/identification/main.php). Since the transesterified fatty acids are identified by GC in comparison to retention time libraries, the more unusual fatty acids typical of several methanotrophs (such as 16:1o8c, 18:1o8c) may not be accurately identified. To confirm their presence requires specific double-bond directed derivatization (Nichols et al., 1986) and subsequent GC/MS analysis. Certain fatty acid components are also produced at highly variable levels (e.g., affected by O2 availability), especially cyclopropane fatty acids ( Jahnke and Nichols, 1986) and thus there is a need for standardization in culture conditions. Quinone analysis may also have value owing to some methanotrophs possessing quinones that have methylenated isoprenoid chains (Watzinger et al., 2008). Analysis of quinone compounds requires HPLC and MS for confirmation (Tindall et al., 2010). Polar lipid analysis is becoming more popular in bacterial taxonomic characterization and has potential value for methanotrophs as some species produce distinctive patterns of polar lipids (Andreev and Galchenko, 1983). It is required that this type of analysis be performed using two-dimensional thin layer chromatography with appropriate use of indicator spray reagents. Tindall et al. (2010) describes appropriate standards needed for description of fatty acid, quinone, and polar lipid data. Methanotrophs also contain unusual sterol and hopanoid compounds (Cvejic et al., 2000; Elvert and Niemann, 2008). No systematic study has been completed to determine if these lipids vary between species or groups and whether they have taxonomic value. Such information could be valuable as these more unusual lipids could also be useful as environmental biomarkers.

3.4. Informative sequence-based analyses and genomic comparisons DNA:DNA hybridization data is an obligatory element for novel taxa descriptions unless they are distinct in terms of 16S rRNA sequence similarity. The generally accepted threshold is 97%; however, a description must also be supported by other evidence including phenotypic, chemotaxonomic, and other genetic data (Tindall et al., 2010). DNA:DNA hybridization can be performed by a variety of means including the S1 nuclease technique (Christensen et al., 2000), renaturation kinetics (De Ley et al., 1970) or by using photobiotin labeled DNA (Hirayama et al., 1996). The DSMZ offers a paid service to perform DNA:DNA hybridization analyses. It has become the practice to also compare novel strains on the basis of similarity of the pmoA gene though the comparisons may not correlate with 16S rRNA gene sequence data for closely related taxa. The analysis also further validates the nature of methanotrophy in a given strain. Though sMMO genes can be also identified by PCR normally the naphthalene oxidation assay (Brusseau et al., 1990) is performed for cells grown in the

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presence and absence of copper ions. Oxidation leading to the formation of naphthol in the absence of copper and abolishment of this activity in the presence of copper is usually sufficient evidence for sMMO activity. It should be noted some type II methanotrophs, primarily Methylocella spp. have been described that lack pMMO and only form sMMO. Evidence for nitrogen fixation can also be established by PCR detection of nitrogenase nifH genes; however, a resting acetylene reduction assay must be also used to confirm nitrogenase activity and supported by evidence of growth in a medium lacking a combined nitrogen source (Bowman, 2006).

3.5. Culture collections Though methanotrophs have been shown to be stably maintained under a methane atmosphere for several months (Bowman, 2006), many methanotrophic species (especially type I methanotrophs) tend to survive long-term preservation (lyophilized, deep freezing in glycerol suspensions, etc.) sometimes poorly or not at all. Green and Woodford (1992) suggest methanol and polyvinylpyrrolidone could be useful cryoprotectants; however, these chemicals need to be tested with species isolated in recent years. Thus, for long-term maintenance of type strains or other important strains for perpetuity and also to make them publically accessible the strains need to be housed in culture collections that have already extensive methanotroph collections, for example, NCIMB, DSM, and VKM (Table 4.2). As listed in Table 4.2, the available type strains are housed in a disparate range of collections. It would be ideal thus that methanotrophs are distributed as much as possible thus to reduce the chance of being lost, for example, a situation made possible by culture collection extinction brought about potentially by lack of funding. All species that are in the “lost” category are type I methanotrophs thus extra attention is obviously required for novel taxa described in this group due to their apparent sensitivity to freezing.

4. Concluding Remarks The role of methane as a greenhouse gas and current suspected trends of global climate change (e.g., Shakhova et al., 2010) has resulted in much scientific interest and research investigating the significance aerobic methanotrophy plays as a biological sink for methane. One aspect of this research is the discovery and characterization of novel methanotrophic bacteria. The current trend suggests that we still have an undeveloped concept of the diversity of methanotrophs (Kip et al., 2010; Op den Camp et al., 2009; Stoecker et al., 2006) and as mentioned above their inherent physiological capacities are still unplumbed. Thus, cultivation studies coupled to the

Table 4.2

Aerobic methanotroph pure culture location and environmental locations of isolation

Genus name

Extant species and type strain location

Type I methanotrophs Methylomonas M. methanica ATCC, NCIMB, VKM Methylobacter M. luteus ATCC, NCIMB; M. psychrophilus VKM, M. tundripaludum ATCC, DSM, M. whittenburyi ATCC, NCIMB Methylomicrobium M. agile ATCC NCIMB; M. albus ATCC, NCIMB, VKM; M. pelagicus NCIMB; M. alcaliphilum NCIMB, VKM; M. japanense NBRC, FERM-P, VKM; M. kenyense NCCB, NCIMB, VKM Methylosoma M. difficile DSM, JCM Methylovulum M. miyakonense NBRC, DSM Methylosarcina M. fibrata ATCC, DSM; M. lacus ATCC, JCM; M. quisquiliarum ATCC, DSM Methylohalobius M. crimeensis ATCC, DSM Methylothermus M. thermalis VKM Methylococcus group Methylococcus M. capsulatus ATCC; M. thermophilus NCIMB Methylocaldum

M. gracile NCIMB, VKM; M. szegediense NCIMB

Known environmental associations

Terrestrial (soil, water), groundwater Temperate and polar terrestrial (soil, water) Terrestrial (soil, water), freshwater and soda lakes, estuarine, seawater

Freshwater lakes Forest soil

Hypersaline lake (Crimea, Russia) Hot spring (Japan) Terrestrial (soil, water); warm spring, ground water Warm springs, groundwater (continued)

Table 4.2

(continued)

Genus name

Extant species and type strain location

Type II methanotrophs Methylosinus M. sporium ATCC, NCIMB; M. trichosporium ATCC, NCIMB Methylocystis M. echinoides IMET, VKM; M. heyeri DSM, VKM; M. hirsuta ATCC, DSM; M. parvus ATCC, DSM, NCIMB, VKM; M. rosea ATCC, DSM Methylocapsa M. acidiphila DSM, NCIMB Methylocella M. palustris ATCC; M. silvestris DSM, NCIMB; M. tundra DSM, NCIMB

Known environmental associations

Terrestrial (soil, water), groundwater Terrestrial (soil, water), groundwater, tundra soil Acidic peat/sphagnum moss soils Acidic peat/sphagnum moss soils, tundra soils

ATCC, American Type Culture Collection, Manassas, VA, USA (www.atcc.org); DSM, German Culture Collection, Braunschweig, Germany (www.dsmz.de); NCIMB, National Collection of Industrial and Marine Bacteria, Aberdeen, Scotland, UK (www.ncimb.com); VKM, All-Russian Collection of Microorganisms, Russian Academy of Sciences, Institute of Biochemistry and Physiology of Microorganisms, Pushchino, Moscow Region, Russia (www.vkm.ru); JCM, Japan Collection of Microorganisms, Institute of Physical and Chemical Research (RIKEN), Wako, Saitama, Japan (www.jcm.riken.go.jp); NCCB, Netherlands Culture Collection of Bacteria (formerly Phabagen and LMD), Utrecht University, Utrecht, the Netherlands (www.cbs.knaw.nl/nccb); FERM-P, Fermentation Research Institute, Agency of Industrial Science and Technology, Chiba, Japan; NBRC, NITE Biological Resource Center, Department of Biotechnology, National Institute of Technology and Evaluation, Chiba, Japan (www.nbrc.nite.go.jp). These species no longer appear to be available due to loss of the cultures or due to extinction of the culture collections in which they were housed: Methylobacter marinus; Methylococcus chroococcus; Methylococcus mobilis; Methylomonas aurantiaca; Methylomonas fodinarum; Methylomonas scandinavica; Methylocaldum tepidum; Methylosphaera hansonii. The species Crenothrix polyspora is not available as a pure culture (www.bacterio.cict.fr).

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increasing knowledge capture possible with ‘omics-based technologies, will likely result in methanotroph systematic biological knowledge having periodic and potentially exciting updates. This is especially possible given the very recent appearance of high-throughput proteomics technologies (Yates et al., 2009) that moves us beyond the inert nature of genome sequence data to the dynamic mapping of physiological information in whole-genome contexts. To galvanize this and maximize the utility of novel strains to scientific research, an excellent platform requires to be maintained for the systematic of methanotrophs and stable and reliable repositories for strains.

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C H A P T E R

F I V E

Methylococcus capsulatus (Bath): From Genome to Protein Function, and Vice Versa Odd A. Karlsen, Frode S. Berven,1 June I. Bagstevold, Øivind Larsen,2 and Harald B. Jensen Contents 1. Introduction 2. Mapping of the Outer Membrane Proteome of M. capsulatus 2.1. Prediction of OMPs from the predicted M. capsulatus proteome 2.2. Experimental identification of (predicted) OMPs. Subfractionation and 2DE analysis of the M. capsulatus proteome 2.3. Identifying unannotated proteins in the M. capsulatus genome 3. Inner Membrane Protein Complexes Studied with Blue-Native Polyacrylamide Gel Electrophoresis (BN-PAGE) 3.1. Detergent extraction of the M. capsulatus inner membrane protein complexes 3.2. BN-PAGE of Triton X-100 solubilized membranes 3.3. Large-scale 2D BN-/SDS-PAGE 4. Visualization and Identification of M. capsulatus c-type Heme Proteins 4.1. Heme staining of M. capsulatus c-type heme proteins Acknowledgments References

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Abstract The genome sequence of Methylococcus capsulatus (Bath), considered a model methylotroph, was published in 2004 [Ward, N., et al. (2004). Genomic insights into methanotrophy: the complete genome sequence of Methylococcus Department of Molecular Biology, University of Bergen, Bergen, Norway Present address: Proteomic Unit (PROBE), Department of Biomedicine, University of Bergen, Bergen, Norway 2 Present address: Uni Environment, Bergen, Norway 1

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00005-X

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2011 Elsevier Inc. All rights reserved.

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capsulatus (Bath). PLoS Biol. 2, e303]. In the postgenomic era, the challenge is to determine the gene function, and to this end, genomics must be complemented with proteomic approaches. This chapter describes some experimental and computational approaches we have used and developed for the exploration of the genome and proteome of M. capsulatus (Bath).

1. Introduction As the first complete genome sequence from an obligate methanotroph, the genome sequence of Methylococcus capsulatus (Bath) was published in 2004 (Ward et al., 2004). Analysis of the 3.3-Mb genome allowed the annotation of 3113 open-reading frames (ORFs), with about 1766 proteins similar to proteins of known function and role category, 514 conserved hypothetical proteins, and 504 hypothetical proteins. The genome sequence confirmed previous studies on carbon assimilation (Strom et al., 1974; Taylor et al., 1981), nitrogen fixation (Murrell and Dalton, 1983), hydrogenase (Hanczar et al., 2002), and the presence of particular and soluble methane monooxygenase (Stainthorpe et al., 1989, 1990; Stolyar et al., 1999). More surprisingly was the presence of two ORFs with high sequence similarity to 2-oxyglutarate dehydrogenase, one of the key enzyme activities of the tricarboxylic acid cycle not found in type I methanotrophs (Davey et al., 1972). The genome is continuously updated (NCBI GenBank Accession number AE017282). The genome sequencing uncovered redundancy in many pathways: some 200 linage-specific gene duplications were found covering both methane oxidation, carbon assimilation, amino acid biosynthesis, energy metabolism, transport, regulation, and environmental sensing (Ward et al., 2004). Thus, in addition to the previously known redundancy for methane monooxygenase (Stanley et al., 1983), all of the other steps involved in methane oxidation to CO2 are covered by redundant enzyme systems (Ward et al., 2004). The high content of duplicated ORFs, the relatively large number of c-type cytochromes (Karlsen et al., 2008), and membranemodifying components, including sterols (Bird et al., 1971), hopanoids ( Jahnke et al., 1992), and trans fatty acids (Loffler et al., 2010), are consistent with an organism able to rapidly adapt to varying growth conditions. It has become clear that the large number of ORFs predicted from the increasing number of sequenced genomes alone has limited value in relation to a complete understanding of gene function and cellular physiology. The genome sequence is the blueprint of the organism and its theoretical (metabolic) capabilities. When entering the postgenomic era, the challenge is to determine the gene function, and genomics must therefore be complemented with proteomic approaches. Thus, it is necessary to examine the

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expression levels of the genes and protein responses to biological and environmental stimuli, posttranslational modifications (PTMs), and the interactions of these molecules in the cell. Computational algorithms that have been developed for identifying functional genes in bacterial genomes have recently been reviewed (Gao and Chen, 2010; Poptsova and Gogarten, 2010). The use of computational analysis for exploring the genome has proved very useful for the prediction of genes and the functions, location, and structure of their protein products. Such analysis includes sequence alignment, chromosomal location, structural prediction (Gao and Chen, 2010), and comparative genome analysis (Poptsova and Gogarten, 2010). These analyses can be the fundament for either generating hypotheses that must be experimentally verified or for supporting the conclusions and findings of the experimental work. In the following sections, we describe the experimental and computational approaches we have used and developed for the exploration of the M. capsulatus genome and proteome. Computational tools, including some developed in-house, for the prediction of outer membrane proteins (OMPs) were combined with subproteome fractionation and proteomics methods for the identification of transmembrane b-barrel proteins and lipoproteins. We show how proteomics, in combination with an ORF library, can be used in order to identify c-type cytochromes, also those located to the M. capsulatus cellular surface. We also demonstrate the use of Blue-Native PAGE to reveal protein complexes in the inner membrane of M. capsulatus (Bath), which can aid in decoding the functional proteome.

2. Mapping of the Outer Membrane Proteome of M. capsulatus Integral b-barrel proteins constitute a major protein type in the OM of Gram-negative bacteria. These types of proteins have diverse functions such as passive nutrient intake, active ion transport, membrane anchors, and enzymes (Schulz, 2000). Another group of OMPs is the lipoproteins that are attached to the OM through lipid anchors. Peripheral proteins may be associated to the outer and inner leaflet of the OM, or they can be associated with OMPs through protein–protein interactions, without themselves being in direct contact with the OM. In the following sections, we focus on the prediction of integral b-barrel proteins and lipoproteins from the genome sequence, and experimental identification of such proteins. Most of the tools and approaches we present here are generic and are applicable to most Gram-negative bacteria. The strategies for subproteome fractionation may, however, vary from one bacterial species to another.

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2.1. Prediction of OMPs from the predicted M. capsulatus proteome 2.1.1. Prediction of b-barrel proteins A large number of software tools that predict integral b-barrel OMPs have been developed, and an overview of most of these tools is given elsewhere (Remmert et al., 2009). Some of them can only process one sequence at the time, while others can process thousands of sequences simultaneously. The latter category includes the b-barrel Outer Membrane protein Predictor (BOMP) and HHomp (Berven et al., 2004; Remmert et al., 2009) and is preferred for the prediction of the entire b-barrel subproteome. In addition to b-barrel prediction, the positive predictions are analyzed for the presence of signal sequences, and this is an indication of transport of the protein to the membranes or the periplasm, or secretion. BOMP is based on two separate components to recognize integral b-barrel proteins: one component calculates an integral b-barrel score of the sequence based on the extent to which the sequence contains stretches of amino acids typical of transmembrane b-strands, while the other component searches for a C-terminal pattern typical of many integral b-barrel proteins. The following steps describe the use of BOMP to predict b-barrel OMPs in M. capsulatus, and additional experimental analysis that may support the predictions. 1. Go to the BOMP webpage: http://services.cbu.uib.no/tools/bomp and upload a file with the protein sequences in FASTA format. 2. If BLASTp information is desired for the dataset, check this option and set the E-value (typically, 1e
10). This will increase the run time to some extent and will also provide additional information to the predictions, and therefore, we recommend that this option is used. 3. The predictions will be given as categories 1–5 with increasing probability of being a b-barrel protein with higher category; Table 2 in Berven et al. (2004) gives information about the meaning of the categories. 4. Generally, sequences ending up in categories 4 and 5 are considered to be b-barrel OMPs with very high probability. When submitting all the predicted protein encoding sequences from 10 genomes of Gram-negative bacteria, none of the sequences predicted in categories 4 and 5 was confirmed to have localization other than the OM. 5. The output from BOMP is a list of predicted b-barrel OMPs and the category to which they belong. In addition, a FASTA file of these sequences is created, and this list can be submitted to other prediction programs to further increase the reliability of the predictions. 6. We have typically used SignalP (http://www.cbs.dtu.dk/services/SignalP/) and secondary structure prediction programs like PSIPRED (http://bioinf. cs.ucl.ac.uk/psipred/) to obtain more information about the predicted b-barrel OMPs, but other b-barrel OMPs predictors can also be used, as for example, HHomp (Berven et al., 2004; Remmert et al., 2009).

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2.1.2. Prediction of lipoproteins Lipoproteins belong to another important category of proteins of the OM. A couple of software tools have been developed to predict such proteins from larger genome data, including Lipo (http://services.cbu.uib.no/tools/ lipo) developed by us (Berven et al., 2006), and LipoP (http://www.cbs.dtu. dk/services/LipoP/; Juncker et al., 2003). Lipo recognizes lipoproteins based on a lipo-box pattern that was extracted from 183 Gram-negative lipoproteins collected from Swiss-Prot. The predicted lipoproteins were divided into three categories: high, medium, and low, based on how frequent each amino acid appears in each position in the lipo-box in the 183 training set proteins. In the following sections, we will describe our approach for the prediction of lipoproteins in M. capsulatus using Lipo. 1. Go to the Lipo webpage: http://services.cbu.uib.no/tools/lipo and upload a file with the protein sequences in FASTA format. 2. The predictions will be given as categories high, medium, low, indicating the likelihood of a correct prediction. 3. The output from Lipo is a list of predicted lipoproteins and the category to which they belong, and whether the lipoprotein is likely to be located in the IM or OM. In addition, a FASTA file of these sequences is created, and this list can be submitted to other prediction programs to further increase the reliability of the predictions. 4. The predictions in the high-scoring category had a precision of 92%, medium 73.7%, whereas the precision of the low-scoring category was only 13.3%. The number of lipoproteins found in the low category was only 5%, so if high confidence is more important than the possibility of losing some lipoproteins in the prediction, the low-category predictions can be excluded. 5. We have typically used SignalP (http://www.cbs.dtu.dk/services/SignalP/) to obtain more information about the predicted lipoproteins, but other lipoprotein predictors can also be used if increased confidence is desired, as for example, LipoP ( Juncker et al., 2003), DOLOP (http://www.mrc-lmb. cam.ac.uk/genomes/dolop/; Babu et al., 2006; Madan Babu and Sankaran, 2002), and PROSITE (http://au.expasy.org/prosite/).

2.2. Experimental identification of (predicted) OMPs. Subfractionation and 2DE analysis of the M. capsulatus proteome 2.2.1. Subfractionation M. capsulatus cells are fractionated according to the protocols outlined in a separate chapter in this issue (chapter eleven). The resulting protein fractions are enriched in soluble proteins, inner membrane proteins, OMPs, membrane-associated proteins, and surface-associated proteins, respectively.

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2.2.2. 2DE analysis The different protein fractions residing from the fractionation procedure were analyzed by 2DE. By this approach, it is relatively easy to get an overview of the presence and relative amounts of each protein in the different fractions. Most proteins are rarely present exclusively in one fraction, but when looking at the spot abundances from the 2DE gels, it is possible to determine in which fraction the protein is most abundant. This comparison reveals information about the localization of the proteins in the cell and gives indications on which type of OMP that has been identified, for example, surface associated, integral b-barrel protein, membrane associated, etc. The 2DE separation of M. capsulatus OMPs is described in detail in Berven et al. (2003). Protein spots enriched in the OM fractions were analyzed by mass spectrometry and/or N-terminal sequencing revealing their identity, as described elsewhere (Berven et al., 2003, 2006). A representative combined 2DE gel of the enriched OMP fraction is shown in Fig. 5.1. The numbered protein spots have been identified and are given in Table 5 in Berven et al. (2006). 2.2.3. Multiple spots When analyzing the OMPs of M. capsulatus (and in some instances other protein fractions), we observed that many spots in the gel corresponded to the same protein species. The multiple spot patterns could be horizontal as well as vertical. It is not always easy to distinguish between PTMs and pI

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Figure 5.1 2DE gel separation of M. capsulatus OMPs. A 2DE gel showing the OMP fraction of M. capsulatus separated using six overlapping narrow range IPG strips with overlapping pH intervals, followed by separation of each strip using 12.5% SDS-PAGE. The six resulting gel images were combined to give one large 2DE gel. All strips were loaded with equal protein amounts. Spots were detected by silver staining. Spot numbers refer to Table 5 by Berven et al. (2006). The figure is reproduced from Fig. 1 in the same article with kind permission from Springer ScienceþBusiness Media.

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conformational equilibria of proteins (Berven et al., 2003). The origin of horizontal spot patterns could be caused by PTMs giving charge alterations, as well as conformational equilibria arising in vitro during the 2DE procedure. The 2DE-based protocol outlined in Section 2.2.3.1 can be followed in order to reveal if the latter is the case. Figure 5.2 shows the result of using this procedure to analyze a train of polypeptide spots from the OMP fraction, spots 1–3 in Fig. 5.1, all representing the MopG protein, see Table 5 in Berven et al. (2006). It is clear from the figure that the entire original spot pattern can be regenerated from (any) one of the excised spots. Due to the great stability of b-barrel proteins (Schulz, 2000), oligomers, folded monomers, and denatured monomers can be found in varying amounts in 2DE, due to incomplete denaturation (Berven et al., 2003; Exner et al., 1995; Fjellbirkeland et al., 1997). An example can be seen in Fig. 5.1, spots 11–17, which all represent the OMP MopB (see Table 5 in Berven et al., 2006). See the 1DE procedure outlined in Section 2.2.3.2 in order to investigate such a case. 2.2.3.1. Analyzing (in vitro-generated) horizontal multiple spot patterns in 2DE gels

1. Analyze the protein sample using 2DE (Berven et al., 2003), and load as much protein as possible. 2. Excise one spot from the horizontal train of spots observed in the Coomassie Brilliant Blue R-250 stained 2-DE gels. 3. Elute the excised polypeptide e.g. by using the model 422 Electro-Eluter from Bio-Rad, utilizing a volatile electrode buffer, according to the manufacturer’s protocol. pH 4.3 Original spot pattern Re-run spot 1 Re-run spot 2

Train 3 1

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Figure 5.2 2DE reapplication of 2DE separated polypeptides. Polypeptide spots of MopG, labeled as train 3 in the figure, were excised from Coomassie Brilliant Blue-stained 2-DE gels and resubjected to a new 2DE separation. The polypeptide pattern observed after the first 2DE separation is shown in the upper panel. The spot numbers indicate the polypeptide spot that was resubjected to 2DE. Narrow-range IPG-strips (18 cm) pH 4.0–5.0 were used in the first-dimensional separation and SDS-polyacrylamide gels (12.5%) in the second-dimension electrophoresis. The gels were stained with silver nitrate. The figure is reproduced from Fig. 1 in Berven et al. (2003) with kind permission from Wiley-VCH verlag Gmbh & Co. KGaA.

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4. Precipitate the eluted polypeptide using 80% (v/v) acetone, allow at least 1 h on ice, resuspend in standard IEF buffer and resubject the polypeptide to 2DE separation, but without the use of an alkylating agent. 5. After the second 2DE separation, the gel is typically silver-stained in order to account for the inevitable loss of material during sample preparation and the second 2DE run, and since one original spot can split up to many isoforms. 6. If a similar spot pattern as the original one is generated from one single excised spot, then it is likely that the original spot pattern was not generated by in vivo PTMs, but rather generated during the 2DE analysis procedure. 2.2.3.2. Analyzing structural isoforms of OMPs resulting in spots with identical pI but different apparent MW in 2DE gels Analyze the protein sample on 2DE, excise and elute relevant polypeptide spots as above (Section 2.2.3.1, 1–4), but resuspend in SDS sample buffer and heat each polypeptide for at least 5 min at 95  C before 1D SDS-PAGE. Most proteins, also the very stable outer membrane b-barrel proteins, are now found as denatured monomers.

2.3. Identifying unannotated proteins in the M. capsulatus genome 2.3.1. Generation and use of an ORF library A database is generated from the total genome sequence using GLIMMER (http://cbcb.umd.edu/software/glimmer/). All the theoretically possible ORFs with a length of more than 100 bases are generated, resulting in a database of 31,490 sequences. Thus, we are sure that all putative ORFs, also the unannotated ones, are included in the database. A refined database based on proteins containing the c-type heme motif, CxxCH, can then be constructed from the ORF library. All mass spectra obtained from excised polypeptide spots/bands, where no identification was obtained, were searched against the ORF library. This gave us the possibility to identify potentially important proteins that would not have been discovered if the original annotated database had been used (Berven et al., 2006; Karlsen et al., 2008).

3. Inner Membrane Protein Complexes Studied with Blue-Native Polyacrylamide Gel Electrophoresis (BN-PAGE) As the number of sequenced genomes increases rapidly, the challenge is to assign a function to each protein and elucidate its interactions with other proteins and macromolecules in the cell. BN-PAGE was initially

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developed to study protein–protein interactions of membrane proteins, which due to their hydrophobic character, have been difficult to study with other methods (Schagger and von Jagow, 1991). BN-PAGE has later also been applied for studying soluble protein complexes (CamachoCarvajal et al., 2004), and has proved to be an efficient method for a highresolution separation of multiprotein complexes, maintaining their native conformation and providing information of their relative abundance and exact composition. In the following sections, we describe (unpublished results) how to empirically find the best-suited detergent concentrations for efficient extraction of inner membrane protein complexes, and we present the combined BN/SDS-PAGE procedure to resolve protein complexes present in inner-membranes of M. capsulatus. Proteins of interest can be identified with MS, and these findings can be compared to genomic structure to provide information about gene clusters and operons, and composition of the protein complexes and possibly, on their biological role.

3.1. Detergent extraction of the M. capsulatus inner membrane protein complexes To study membrane proteins and their complexes by BN-PAGE, it is crucial to extract them from their biological surroundings. Mild detergents are used in order to substitute the hydrophobic environment of the lipid bilayer and transfer the membrane proteins into the aquatic phase. It is important to notice that protein complexes differ substantially in stability. Detergent solubilization must therefore be tested empirically with different detergents at different concentrations and incubation times to ensure that protein–protein interactions are maintained (Reisinger and Eichacker, 2006). Triton X-100 has proved to efficiently solubilize the inner membrane of M. capsulatus, as described by (Fjellbirkeland et al., 1997) and elsewhere in this issue. Total membranes of M. capsulatus are incubated with different concentrations of Triton X-100 and subsequently analyzed by BN-PAGE to find a suitable concentration of detergent for efficient extraction of the inner membrane protein complexes. The same general procedure can be applied to membranes isolated from other bacterial species, and the type of detergent can be changed (e.g., BOG, DDM, Digitonin; Reisinger and Eichacker, 2006). 1. A total membrane fraction from M. capsulatus is obtained, as described in chapter eleven in this issue. 2. Samples containing 100 mg of total membranes are centrifuged at 100,000g for 1 h at 4  C. The pellets are resuspended in 17.5 ml BNPAGE sample buffer (750 mM e-amino caproic acid, 50 mM Bis–Tris, pH 7.0, and 0.5 mM EDTA–Na2) and incubated with Triton X-100 in concentrations ranging from 0.25% to 2.5% (w/v). 3. The samples are solubilized for 30 min at 4  C and centrifuged as above for separation of the insoluble material from the solubilized protein complexes.

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4. The supernatants are gently mixed with 1 ml BN-PAGE loading buffer (750 mM e-amino caproic acid and 5% Coomassie G-250) and BN-PAGE is performed, as described in Section 3.2. Coomassie G-250 renders the proteins with negative charges prior to the electrophoresis.

3.2. BN-PAGE of Triton X-100 solubilized membranes 1. A NativePAGETM gel cassette (4–16%, Invitrogen) is prepared as described by the manufacturer. The BN-PAGE gel can also be cast in the lab, as described elsewhere (Reisinger and Eichacker, 2006). 2. Cathode buffer (50 mM Tricine, 15 mM Bis–Tris, pH 7.0, 0.02% Coomassie G-250) is added to the wells prior to protein loading of the Triton X-100 extracted samples. A molecular mass marker used for native gels is applied to one of the lanes (e.g., NativeMark Unstained Protein Standard (Invitrogen)). 3. Cathode buffer and anode buffer (50 mM Bis–Tris, pH 7.0) are added to the inner and outer buffer chamber, respectively. Both buffers are prechilled to 4  C. 4. The electrophoresis is carried out at 4  C with a constant voltage of 150 V for 1 h, which subsequently is increased to 250 V until the blue dye-front reaches the bottom of the gel. The blue cathode buffer is replaced with a colorless cathode buffer (lacking Coomassie) when the Dye-front has migrated approximately one-third of the gel. Proteins are visualized by staining with Coomassie R-250 (Fig. 5.3).

3.3. Large-scale 2D BN-/SDS-PAGE BN-PAGE is often combined with SDS-PAGE in a second dimension, a variant of the common two-dimensional isoelectric focusing/SDS-PAGE. In order to identify the subunits of the protein complexes that were maintained during the BN-PAGE, a lane from the BN-gel is treated with a denaturing solution containing SDS and reducing agents, and placed on top of the second-dimension stacking gel. As a result, subunits comprising the same complex will dissociate and be separated based on their individual molecular mass in the second-dimension electrophoresis. Due to comigration in the first dimension, subunits will appear as spots along a straight vertical line (Reisinger and Eichacker, 2006). 3.3.1. The first-dimension BN-PAGE 1. An M. capsulatus total membrane faction, corresponding to 400 mg of proteins, is centrifuged for 100,000g for 1 h at 4  C.

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% Triton X-100 kDa 0.25 0.5 1.0 1.5 2.0 1048

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Figure 5.3 Empirical testing of Triton X-100 extraction of inner membrane protein complexes. Approximately 100 mg of proteins from the enriched total membrane fraction were solubilized using Triton X-100 concentrations ranging from 0.25% to 2.5% (w/v), and subsequently analyzed with BN-PAGE. An increased efficiency of protein extraction and electrophoretic mobility shift toward protein complexes of higher molecular weight are observed when a 1% Triton X-100 concentration is used for protein extraction. This is especially evident for the protein band migrating with a molecular mass between 242 and 480 kDa, as indicated in the figure, suggesting that a Triton X-100 concentration of approximately 1% is suitable for the extraction of the inner membrane proteins. The Native Mark Unstained Protein Standard (Invitrogen) is shown to the left. The gel is stained with Coomassie R-250.

2. Resuspend the pellet in 70 ml BN sample buffer (as described above) and add Triton X-100 to a final concentration of 1.25%. 3. Solubilize the sample for 30 min at 4  C. 4. The sample is centrifuged at 100,000g for 1 h at 4  C to remove the membranes from the soluble protein complexes. The supernatant is gently mixed with 5 ml of BN loading buffer (as described above). The BN-PAGE electrophoresis is performed as described (Section 3.2) in a large polyacrylamide gel electrophoresis system (typically, a gel size of 18–16 cm) with a BN gradient gel of desired concentration. The power program is divided into two phases: Phase I 1000 V, 6 mA, 24 W, 45 min; Phase II: 1000 V, 12 mA, 24 W, 2 h. The electrophoresis is carried out at 4  C. 3.3.2. The second-dimension SDS-PAGE 1. The lane from the BN-PAGE electrophoresis containing the separated protein complexes is excised from the BN polyacrylamide gel.

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2. The gel is transferred to a small compartment and incubated for 20 min with 2% SDS and 2% b-mercaptoethanol while rotating. The equilibrated lane is placed horizontally on top of the second-dimension polyacrylamide gel (uniform polyacrylamide concentration of 12.5% with a 4% stacking gel). 3. A molecular mass marker (denatured) is added onto a small piece of Whatman 3MM paper and placed beside the first-dimension lane. 0.5% agarose in Tris-glycine electrophoresis buffer is pored over the lane and the molecular mass marker to seal their position. Care is taken to avoid trapping air between the lane and the stacking gel. The electrophoresis is carried out using the following power program: Phase I: 1000 V, 30 mA, 30 W, 30 min; Phase II: 1000 V, 8 mA, 30 W, 14 h. The electrophoresis unit is cooled to 15  C during the electrophoresis. The resulting gel can be stained with e.g. Colloidal Coomassie or electrotransferred to a membrane for protein immunoblot analyses (Fig. 5.4). Subunits of a protein complex are observed as spots aligned in a vertical line. These spots can be excised from the polyacrylamide gel and identified by mass spectrometry. Reverse genetics toward the M. capsulatus genome provides information about genomic organization (operon), other putative interaction partners, and possibly biological function (e.g., see Fig. 5.4).

4. Visualization and Identification of M. capsulatus c-type Heme Proteins The genome sequencing of M. capsulatus revealed an unexpected large complement of c-type cytochromes; 57 proteins containing a c-type hemebinding motif (CxxCH) were identified during the genome annotation (Ward et al., 2004). This is in clear contrast to Escherichia coli where only seven c-type cytochrome-encoding genes are present in the genome (Methe et al., 2003). The large number of proteins with c-type heme-binding motifs makes M. capsulatus resemble some facultative or strictly anaerobic bacteria that contain numerous c-type cytochromes, such as the dissimilatory metal reducing bacteria Shewanella oneidensis MR1 and Geobacter sulfurreducens, with 42 and 111 predicted c-type cytochromes, respectively (Heidelberg et al., 2004; Methe et al., 2003). Recently, several of the multi c-type heme proteins of M. capsulatus were found located on the cellular surface (Karlsen et al., 2008). This is not commonly observed in bacteria, but a feature of dissimilatory metal-reducing bacteria that utilize an extracellular electron acceptor (Myers and Myers, 2003). These findings imply that redox reactions involving c-type cytochromes can also take place at this cellular localization in methanotrophic bacteria. In the following sections, we outline a protocol for visualizing the

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Figure 5.4 Two-dimensional BN/SDS-PAGE of M. capsulatus inner membrane proteins. (A) Approximately 400 mg of enriched total membrane fraction was solubilized with 1.25% Triton X-100, and separated in a 5–15% BN gradient gel, prior to the second-dimension electrophoresis, using a uniform 12.5% SDS-polyacrylamide gel. Subunits comprising heteromeric protein complexes during the first-dimension BNPAGE appeared along vertical lines in the 2D-gel. Spots 1–3 were identified by MS as the ATP synthase F1a-, b-, and g-subunit, respectively, originating from a protein complex migrating with a molecular mass corresponding to  500–650 kDa. The other polypeptide spots in the same vertical line has not yet been identified, but they correspond well in molecular masses to the ATP synthase subunits F1d (19 kDa), F1e (15 kDa), F0a (29 kDa), F0b (17 kDa), and F0c (8 kDa), as predicted from their encoding genes in the M. capsulatus (Bath) genome. Thus, the ATP synthase complex migrates in the BN-PAGE with an apparent molecular weight that is in line with the expected molecular mass predicted for the M. capsulatus ATP synthase complex, with a possible stoichiometric arrangement of F1a3:b3:g:d:e and F0a:b2:c10–14 as demonstrated for Escherichia coli (Pedersen and Amzel, 1993). Depending on the number of c subunits in the complex, which may vary among different bacteria, the complex in M. capsulatus (Bath) should be about 531–565 kDa.

complement of c-type cytochromes in cellular fractions of M. capsulatus, and how they can be identified by mass spectrometry combined with a c-type heme database constructed from the genome.

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4.1. Heme staining of M. capsulatus c-type heme proteins These methods take advantage of the intrinsic peroxidase activity of the covalently bound heme group of denaturated c-type cytochromes. The procedures described below are based on Dorward (1993), Thomas et al. (1976), and Vargas et al. (1993). 4.1.1. c-type heme staining on protein transfer membranes 1. SDS-PAGE is carried out without the addition of reducing agents. 2. Proteins are electrotransferred to a nitrocellulose/PVDF membrane at 100 V for 1 h. 3. The membrane is washed twice in Tris-buffered saline for 5 min. 4. The membrane is subsequently incubated with ECL as described by the manufacturer (GE Healthcare) and exposed to a Hyperfilm ECL (GE Healthcare) in the dark for 5 to 30 min, depending on the abundance of c-type heme proteins present in the sample. The membranes can be stained for total protein content with Amidoschwarz (nitrocellulose membrane) or Coomassie Blue (PVDF membrane) (Dunn, 1999). 4.1.2. In-gel c-type heme staining 1. SDS-PAGE is carried out without the addition of reducing agents. 2. Tetramethylbenzidin (TMBZ) is solubilized in methanol to a final concentration of 6.3 mM. 3. The polyacrylamide gel is covered with a solution consisting of freshly prepared TMBZ solution and 0.25 M sodium acetate (pH 5.0) in a 3:7 ratio, and incubated for 1–2 h in the dark with a gentle stirring every 10–15 min. 4. H2O2 is added to a final concentration of 30 mM. The staining of the c-type heme proteins should be visible within a few minutes and increases in intensity over the next 30 min. 5. The gel is placed in isopropanol and 0.25 M sodium acetate (pH 5.0) at a ratio of 3:7 in order to remove any precipitated TMBZ, to clear the gel background, and to increase the staining intensity. The acetate-buffered 30% isopropanol solution is replaced twice. 4.1.3. Identification of c-type heme proteins Figure 5.5 shows protein samples extracted from the M. capsulatus surface under different copper concentrations in the growth medium following staining for c-type heme with the protocols outlined above. A parallel Coomassie Blue stained gel of the same samples is used to locate the c-type heme-positive bands. These proteins can be excised from the gel, trypsinated and identified with mass spectrometry analyses. A refined database based on

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A kDa 181.8 115.5 82.2 64.2 48.8

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Figure 5.5 c-type heme staining of the M. capsulatus surfaceome obtained from cultures grown at different copper concentrations. Cultures were grown with copper concentrations corresponding to  0 (lane 1), 0.8 (lane 2), 1.6 (lane 3), 5 (lane 4), and 10 mM (lane 5) copper. (A) A 12.5 % polyacrylamide gel was used and stained with CBB. (B, C) c-type heme staining of (A) using ECL on nitrocellulose membranes (B) and in-gel staining (C). Arrowheads indicate gel bands analyzed by MS, and the numbers correspond to the numbers given in Table 2 in Karlsen et al. (2008). The figure was adopted from Karlsen et al. (2008) with kind permission from John Wiley & Sons, Inc.

proteins containing the c-type heme motif, CxxCH, is constructed from the ORF library, as described in Section 2.3.1. The CxxCH consensus sequence contains the two cysteines that covalently attach the heme group with thioether bonds, and the histidine that is one of the two axial ligands coordinating the heme iron (Barker and Ferguson, 1999). This arrangement is shared by all known c-type cytochromes, and this database will therefore comprise the full complement of putative c-type heme proteins.

ACKNOWLEDGMENTS We thank Johan Lillehaug for his participation and continuous support during our work. This work was supported in parts by grants from the Norwegian Research Council, The Meltzer Foundation, SUP140785/420 (GABI), and Norferm DA.

REFERENCES Babu, M. M., Priya, M. L., Selvan, A. T., Madera, M., Gough, J., Aravind, L., and Sankaran, K. (2006). A database of bacterial lipoproteins (DOLOP) with functional assignments to predicted lipoproteins. J. Bacteriol. 188, 2761–2773. Barker, P. D., and Ferguson, S. J. (1999). Still a puzzle: Why is haem covalently attached in c-type cytochromes? Structure 7, R281–R290.

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Berven, F. S., Karlsen, O. A., Murrell, J. C., and Jensen, H. B. (2003). Multiple polypeptide forms observed in two-dimensional gels of Methylococcus capsulatus (Bath) polypeptides are generated during the separation procedure. Electrophoresis 24, 757–761. Berven, F. S., Flikka, K., Jensen, H. B., and Eidhammer, I. (2004). BOMP: A program to predict integral beta-barrel outer membrane proteins encoded within genomes of Gramnegative bacteria. Nucleic Acids Res. 32, W394–W399. Berven, F. S., Karlsen, O. A., Straume, A. H., Flikka, K., Murrell, J. C., Fjellbirkeland, A., Lillehaug, J. R., Eidhammer, I., and Jensen, H. B. (2006). Analysing the outer membrane subproteome of Methylococcus capsulatus (Bath) using proteomics and novel biocomputing tools. Arch. Microbiol. 184, 362–377. Bird, C. W., Lynch, J. M., Pirt, F. J., and Reid, W. W. (1971). Steroids and squalene in Methylococcus capsulatus grown on methane. Nature 230, 473–474. Camacho-Carvajal, M. M., Wollscheid, B., Aebersold, R., Steimle, V., and Schamel, W. W. (2004). Two-dimensional Blue native/SDS gel electrophoresis of multi-protein complexes from whole cellular lysates: A proteomics approach. Mol. Cell. Proteomics 3, 176–182. Davey, J. F., Whittenbury, R., and Wilkinson, J. F. (1972). The distribution in the methylobacteria of some key enzymes concerned with intermediary metabolism. Arch. Mikrobiol. 87, 359–366. Dorward, D. W. (1993). Detection and quantitation of heme-containing proteins by chemiluminescence. Anal. Biochem. 209, 219–223. Dunn, M. J. (1999). 2-D proteome analysis protocols. In “Methods in Molecular Biology, Vol. 112,” (A. J. Link, ed.).Humana Press Inc., Totowa. Exner, M. M., Doig, P., Trust, T. J., and Hancock, R. E. (1995). Isolation and characterization of a family of porin proteins from Helicobacter pylori. Infect. Immun. 63, 1567–1572. Fjellbirkeland, A., Kleivdal, H., Joergensen, C., Thestrup, H., and Jensen, H. B. (1997). Outer membrane proteins of Methylococcus capsulatus (Bath). Arch. Microbiol. 168, 128–135. Gao, J., and Chen, L. L. (2010). Theoretical methods for identifying important functional genes in bacterial genomes. Res. Microbiol. 161, 1–8. Hanczar, T., Csaki, R., Bodrossy, L., Murrell, J. C., and Kovacs, K. L. (2002). Detection and localization of two hydrogenases in Methylococcus capsulatus (Bath) and their potential role in methane metabolism. Arch. Microbiol. 177, 167–172. Heidelberg, J. F., Seshadri, R., Haveman, S. A., Hemme, C. L., Paulsen, I. T., Kolonay, J. F., Eisen, J. A., Ward, N., Methe, B., Brinkac, L. M., Daugherty, S. C., Deboy, R. T., et al. (2004). The genome sequence of the anaerobic, sulfate-reducing bacterium Desulfovibrio vulgaris Hildenborough. Nat. Biotechnol. 22, 554–559. Jahnke, L. L., Stan-Lotter, H., Kato, K., and Hochstein, L. I. (1992). Presence of methyl sterol and bacteriohopanepolyol in an outer-membrane preparation from Methylococcus capsulatus (Bath). J. Gen. Microbiol. 138, 1759–1766. Juncker, A. S., Willenbrock, H., Von Heijne, G., Brunak, S., Nielsen, H., and Krogh, A. (2003). Prediction of lipoprotein signal peptides in Gram-negative bacteria. Protein Sci. 12, 1652–1662. Karlsen, O. A., Lillehaug, J. R., and Jensen, H. B. (2008). The presence of multiple c-type cytochromes at the surface of the methanotrophic bacterium Methylococcus capsulatus (Bath) is regulated by copper. Mol. Microbiol. 70, 15–26. Loffler, C., Eberlein, C., Mausezahl, I., Kappelmeyer, U., and Heipieper, H. J. (2010). Physiological evidence for the presence of a cis-trans isomerase of unsaturated fatty acids in Methylococcus capsulatus Bath to adapt to the presence of toxic organic compounds. FEMS Microbiol. Lett. 308, 68–75.

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Madan Babu, M., and Sankaran, K. (2002). DOLOP—Database of bacterial lipoproteins. Bioinformatics 18, 641–643. Methe, B. A., Nelson, K. E., Eisen, J. A., Paulsen, I. T., Nelson, W., Heidelberg, J. F., Wu, D., Wu, M., Ward, N., Beanan, M. J., Dodson, R. J., Madupu, R., et al. (2003). Genome of Geobacter sulfurreducens: Metal reduction in subsurface environments. Science 302, 1967–1969. Murrell, J. C., and Dalton, H. (1983). Nitrogen fixation in obligate methanotrophs. J. Gen. Microbiol. 129, 3481–3486. Myers, C. R., and Myers, J. M. (2003). Cell surface exposure of the outer membrane cytochromes of Shewanella oneidensis MR-1. Lett. Appl. Microbiol. 37, 254–258. Pedersen, P. L., and Amzel, L. M. (1993). ATP synthases. Structure, reaction center, mechanism, and regulation of one of nature’s most unique machines. J. Biol. Chem. 268, 9937–9940. Poptsova, M. S., and Gogarten, J. P. (2010). Using comparative genome analysis to identify problems in annotated microbial genomes. Microbiology 156, 1909–1917. Reisinger, V., and Eichacker, L. A. (2006). Analysis of membrane protein complexes by blue native PAGE. Proteomics 6(Suppl. 2), 6–15. Remmert, M., Linke, D., Lupas, A. N., and Soding, J. (2009). HHomp—Prediction and classification of outer membrane proteins. Nucleic Acids Res. 37, W446–W451. Schagger, H., and von Jagow, G. (1991). Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal. Biochem. 199, 223–231. Schulz, G. E. (2000). beta-Barrel membrane proteins. Curr. Opin. Struct. Biol. 10, 443–447. Stainthorpe, A. C., Murrell, J. C., Salmond, G. P., Dalton, H., and Lees, V. (1989). Molecular analysis of methane monooxygenase from Methylococcus capsulatus (Bath). Arch. Microbiol. 152, 154–159. Stainthorpe, A. C., Lees, V., Salmond, G. P., Dalton, H., and Murrell, J. C. (1990). The methane monooxygenase gene cluster of Methylococcus capsulatus (Bath). Gene 91, 27–34. Stanley, S. H., Prior, S. D., Leak, D. J., and Dalton, H. (1983). Copper stress underlies the fundamental change in intracellular location of methane mono-oxygenase in methane utilizing organisms: Studies in batch and continuous cultures. Biotechnol. Lett. 5, 487–492. Stolyar, S., Costello, A. M., Peeples, T. L., and Lidstrom, M. E. (1999). Role of multiple gene copies in particulate methane monooxygenase activity in the methane-oxidizing bacterium Methylococcus capsulatus Bath. Microbiology 145(Pt 5), 1235–1244. Strom, T., Ferenci, T., and Quayle, J. R. (1974). The carbon assimilation pathways of Methylococcus capsulatus, Pseudomonas methanica and Methylosinus trichosporium (OB3B) during growth on methane. Biochem. J. 144, 465–476. Taylor, S. C., Dalton, H., and Dow, C. S. (1981). Ribulose-1,5-bisphosphate carboxylase/ oxygenase and carbon assimilation in Methylococcus capsulatus (Bath). J Gen Microbiol. 122, 89–94. Thomas, P. E., Ryan, D., and Levin, W. (1976). An improved staining procedure for the detection of the peroxidase activity of cytochrome P-450 on sodium dodecyl sulfate polyacrylamide gels. Anal. Biochem. 75, 168–176. Vargas, C., McEwan, A. G., and Downie, J. A. (1993). Detection of c-type cytochromes using enhanced chemiluminescence. Anal. Biochem. 209, 323–326. Ward, N., Larsen, O., Sakwa, J., Bruseth, L., Khouri, H., Durkin, A. S., Dimitrov, G., Jiang, L., Scanlan, D., Kang, K. H., Lewis, M., Nelson, K. E., et al. (2004). Genomic insights into methanotrophy: The complete genome sequence of Methylococcus capsulatus (Bath). PLoS Biol. 2, e303.

C H A P T E R

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Functional Metagenomics of Methylotrophs Marina G. Kalyuzhnaya,* David A. C. Beck,†,‡ and Ludmila Chistoserdova‡ Contents 1. Introduction 2. Enrichment for Specific Functional Types Using SIP 2.1. Sample collection and cell labeling 2.2. DNA extraction, isopycnic centrifugation, and labeled DNA recovery 3. Creating a Metagenomic Database and Linking It to Functionality 3.1. DNA sequencing and assembly 3.2. Gene-centric analysis 3.3. Organism-centric analysis 4. Ultrashort Read-Based Metatranscriptomics 4.1. Principle and strategy 4.2. RNA isolation 4.3. mRNA enrichment and (optional) cDNA synthesis 4.4. Data processing 4.5. Metatranscriptome coverage and specificity 5. Conclusions and Future Perspectives Acknowledgments References

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Abstract It is widely recognized that most microbes in the biosphere remain uncultured and unknown. In the recent few years, whole genome shotgun (WGS) sequencing of environmental DNA (metagenomics) has revolutionized the field of environmental microbiology by allowing one to tap into the genomic content of microbial communities in specific ecological niches, deducing information on their biochemical potentials. However, ascribing specific functions to specific organisms remains very difficult in most cases, due to low sequence coverage and the lack of * Department of Microbiology, University of Washington, Seattle, Washington, USA eScience Institute, University of Washington, Seattle, Washington, USA Department of Chemical Engineering, University of Washington, Seattle, Washington, USA

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Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00006-1

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2011 Elsevier Inc. All rights reserved.

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sequence assembly that result from metagenomics of complex microbial communities. Therefore, methods that link specific biogeochemical processes to specific members of such complex natural communities are urgently needed. We have developed and implemented a functional metagenomics approach that allows such a connection via substrate-specific stable isotope labeling, followed by WGS sequencing of the labeled DNA to describe bacterial populations involved in metabolism of single-carbon compounds in a freshwater lake. We also developed a pipeline for community transcript analysis based on ultrashort read highthroughput sequencing of messenger RNA, matching these to a specific scaffold. The methodologies described in this chapter can be applied in a wide variety of ecosystems for targeting methylotrophs as well as other functional guilds of microbes.

1. Introduction Metagenomics is a fast growing and diverse field within environmental biology directed at obtaining knowledge on genomes of environmental microbes, without prior cultivation, as well as of entire microbial communities. When applied to communities of low complexity, exemplified by the communities of the acid mine drainage biofilm (Tyson et al., 2004) or the symbionts of a marine oligochaete (Woyke et al., 2006), the metagenomics approach, even at a modest sequencing effort, allows for sequence assembly. Thus, analysis of almost complete genomes of the dominant species in these communities can be carried out, including accurate metabolic reconstruction and even detection of strain-specific genomic variants. However, the situation is quite different when metagenomics is applied to communities of high complexity, such as the communities of marine habitats or soils (Rusch et al., 2007; Tringe et al., 2005; Venter et al., 2004). In these cases, significantly larger sequencing efforts resulted in very fragmented assemblies even for the most abundant species, with most of the datasets being represented by singleton sequencing reads. While gene-centric analysis (Tringe et al., 2005) can be conducted on the non-assembled metagenomic data and predictions on the major metabolic pathways can be made, the specific metabolic capabilities are hard or impossible to place into the context of individual species. This approach is especially vulnerable when used with short sequence reads produced by the next-generation sequencing technologies such as 454 Roche (Gomez-Alvarez et al., 2009; Wommack et al., 2008). One approach to directly link a function in the environment to a specific guild performing this function is through the use of a technique known as stable isotope probing (DNA-SIP). This involves feeding the natural population a substrate of interest, labeled by a heavy isotope (e.g., 13C), followed by characterization of the heavy fraction of communal DNA that should be enriched in DNA of microbes that actively metabolize the labeled substrate.

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This technique has been effective in identifying microbes involved in specific biogeochemical transformations such as methylotrophy, phenol degradation, glucose metabolism, etc. (Chen and Murrell, 2010; Friedrich, 2006; Madsen, 2006; Radajewski et al., 2000). Typically, small amounts of DNA are isolated in these experiments, and the DNA is used for phylogenetic profiling and detection of key functional genes, after PCR amplification (Friedrich, 2006). A modification of this approach has been described involving a multiple displacement amplification step, followed by metagenomic library construction and screening for specific marker genes (Chen and Murrell, 2010; Chen et al., 2008). However, we demonstrated that it is possible to scale up the SIP protocol to obtain amounts of DNA sufficient to enable the whole genome shotgun (WGS) sequencing approach, and we applied such an approach to characterize methylotroph communities of Lake Washington (Kalyuzhnaya et al., 2008). Methylotrophy is an important part of the global carbon cycle on this planet (Guenter, 2002; Hanson and Hanson, 1996). Identities of methylotrophs involved in utilization of specific C1 substrates (such as methane, methanol, methylated amines, etc.) in a variety of environments have previously been assessed by both culture-reliant and culture-independent methods (Chistoserdova et al., 2009), the former providing important models for understanding the specific biochemical pathways enabling methylotrophy, and the latter providing insights into species richness within specific functional groups. However, while genomic data for some model methylotrophs are now available (Chistoserdova et al., 2009), these may not represent major players in specific functional guilds. At the same time, current methods for environmental detection provide little insight into the genomic structure of uncultivated methylotrophs. The goal of the functional metagenomics approach is twofold: to reduce the complexity of a community and to directly link specific substrate repertoires of the community to specific functional guilds. The schematic of the enrichment and sequencing flow is shown in Fig. 6.1.

2. Enrichment for Specific Functional Types Using SIP 2.1. Sample collection and cell labeling 1. Samples are collected using an appropriate devise and transported to the laboratory on ice. Samples of Lake Washington sediment used here to demonstrate the method were collected as previously described (Kalyuzhnaya et al., 2004). 2. Microcosms are set up in conditions that mimic the in situ conditions, including substrate concentrations that should approximate the in situ

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12C-DNA 13C-DNA

13C-Methane 13C-Methanol

13C-Methylamine

13C-Formaldehyde 13C-Formate

Analysis Major players

Figure 6.1 General strategy for functional metagenomics exemplified by SIP-based enrichment for methylotrophic functional types.

concentrations. However, they should be high enough to allow for efficient labeling. In our case, each microcosms contained 10 ml of the sediment, 90 ml of Lake Washington water filtered through 0.22 mm filters (Millipore), and one of following 13C-labeled substrates: methane (50% of air), methanol (10 mM), methylamine (10 mM), formaldehyde (1 mM), and formate (10 mM; Kalyuzhnaya et al., 2008). 3. Samples are incubated, preferably at the in situ temperature, for the duration of time that allows some of the DNA to become labeled with 13 C (in Kalyuzhnaya et al., 2008, 3–14 days, dependent on the substrate, at room temperature with shaking were required). Other microcosm incubations were conducted at an in situ temperature (8  C), and lower concentrations of substrates were used, with similar results but requiring longer duration (not shown).

2.2. DNA extraction, isopycnic centrifugation, and labeled DNA recovery 1. Total DNA is extracted using an appropriate protocol for a given sample. PowerSoilR DNA isolation kit (MO Bio Laboratories, Inc., Carlsbad, CA, USA) produces good results with soil and sediment samples. 2. DNA is prepared for CsCl-ethidium bromide density gradient ultracentrifugation as previously described (Radajewski et al., 2000) and centrifuged at 265,000g (Beckman VTi 65 rotor) for 16 h at 20  C. DNA fractions are visualized in UV, and 13C-DNA fractions are collected using 19-gauge needles (Nercessian et al., 2005). DNA preparations may be subjected to a second round of CsCl-ethidium bromide density gradient ultracentrifugation, followed by a standard DNA purification

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procedure (e.g., Sambrook et al., 1989). Optionally, the fractions may be collected and analyzed for comparison.

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C-DNA

3. Creating a Metagenomic Database and Linking It to Functionality 3.1. DNA sequencing and assembly Shotgun libraries are constructed, one per each SIP microcosm. In our original study that utilized the Sanger technology (Kalyuzhnaya et al., 2008), they were constructed in the pUC18 vector (1–3 kb inserts) and vector inserts were sequenced with BigDye Terminators v3.1 and resolved with ABI PRISM 3730 (ABI) sequencers at the Joint Genome InstituteProduction Genomics Facility ( JGI-PGF; Walnut Creek, CA, USA). A total of 344,832 reads comprising 255 Mb of Phred Q20 sequence were generated. Sequences were stringently quality- and vector-trimmed using LUCY (Chou and Holmes, 2001), and the trimmed sequences were both by the individual sample and by en masse (combined assembly) using the PGA assembler. Assembly statistics for the datasets described by Kalyuzhnaya et al. (2008) are shown in Table 6.1. Sequences were automatically annotated and loaded into the JGI’s IMG/M system (http://img.jgi.doe.gov/cgi-bin/m/main.cgi). Sequence coverage and degree of assembly depended on the sequencing effort applied and on the species richness and evenness of the enriched communities. These draft quality assemblies were manually validated and used for all downstream analyses. Currently, the Sanger technology is being replaced by the next-generation sequencing technologies, the longer read length and the higher quality of the read being traded off for the higher throughput and the lower cost (Ansorge, 2009; Lapidus, 2009). With the data generated by these new technologies, new assembly tools and new quality controls are being developed (Lapidus, 2009; Miller et al., 2010). Concurrently, new sequencing technologies are in development, and when they enter the market, new bioinformatics tools will need to be implemented to process the next wave of genomic data types.

3.2. Gene-centric analysis 3.2.1. Phylogenetic markers The community complexity and content can be evaluated via profiling of 16S rRNA genes and other phylogenetic markers. In the case we describe here, community complexity was significantly reduced in microcosms exposed to each of the C1 substrates compared to the complexity of the non-enriched

Table 6.1 Summary of sequencing, assembly and gene prediction statistics

Assembly statistics Number of reads Average read length (bp) Trimmed read length (Mbp) Non-redundant sequence (bp) Percent of reads in contigs Total contigs (> 2 kb) Total singlets Average sequence coverage (x) Highest sequence coverage (x) Average size of contigs (bp) Largest contig (bp) GC content (%) Gene predictions Protein coding genes Genes in COGs Genes in Pfams Predicted enzymes Number of 16S rRNA genes Number of tRNA genes

Methane

Methanol

Methylamine

Formaldehyde

Formate

Combined

Methylotenera

71,808 792 56.85 52.16 10.2 2797 59,417 1.6 7.0 1418 6174 58.9

67,200 797 53.53 50.25 10.0 2871 56,408 1.6 4.8 1288 5913 59.5

83,712 709 59.34 37.23 55.5 7558 29,217 1.9 20.4 2065 20,771 53.0

80,640 712 58.91 57.62 7.3 2583 69,104 1.7 6.4 1166 4714 57.9

41,472 638 26.45 17.57 34.3 3618 18,857 1.9 4.7 1265 6276 65.8

344,832 741 255.08 211.47 27.6 25,877 215,581 1.7 23.1 1593 22,407 58.3

NA NA NA 11.16 100 4078 0 2.1 20.4 2736 15,820 46.2

81,076 43,456 28,090 3089 12 405

77,229 40,773 26,494 3047 12 412

54,340 33,643 23,586 5005 10 376

89,729 46,032 29,375 3065 18 504

28,700 17,112 10,585 1417 5 121

321,503 174,344 115,228 16,780 61 1728

12,719 10,082 8543 3264 3 181

This table is reproduced from the original publication (Kalyuzhnaya et al., 2008). The data currently posted in the JGI’s IMG/M interface differ slightly as a result of a more recent reanalysis by JGI.

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community that we estimate to be over 5000 species (Kalyuzhnaya et al., 2008), and shifted toward specific functional guilds that included both bona fide methylotrophs (Methylobacter tundripaludum, Methylomonas sp., Methylotenera mobilis, Methyloversatilis universalis, Ralstonia eutropha) and organisms only distantly related to any cultivated species, implicating the latter in environmental cycling of C1 compounds. The closest relatives of these included Verrucomicrobia, Nitrospirae, Planctomycetes, Acidobacteria, Cyanobacteria, and Proteobacteria. These data were supported by data on phylogenetic profiling of each metagenomic dataset, based on top BLAST hit distribution patterns (these data can be viewed as part of the IMG/M interface). 3.2.2. Functional genes Enrichment for specific functional genes can be addressed in a similar way, using known genes/proteins as queries in BLAST analyses. We used proteins involved in the reactions of the tetrahydromethanopterin pathway for C1 transfer that is a hallmark pathway in methylotrophs to demonstrate the function-relevant enrichment of the microcosm datasets (Table 6.2). In cases of highly divergent enzymes, multiple queries are required. For example, we used peptide sequences of fae and fae homologs belonging to different phylogenetic groups (Proteobacteria, Planctomycetes, and Archaea) to identify multiple and extremely divergent fae and fae-like sequences in our datasets (Fig. 6.2).

3.3. Organism-centric analysis 3.3.1. Characterizing genomes at high sequence coverage Genomes of individual organisms or populations of closely related strains may be present in a metagenome at high sequence coverage. Estimates of coverage for each organism can be initially gained from the coverage of individual 16S rRNA genes present in a metagenome. For example, Kalyuzhnaya et al. (2008) found that 16S rRNA genes belonging to M. mobilis were covered at up to 20. In such cases, it is reasonable to assume that complete or nearly complete genomes of respective strains may be present in a metagenomic dataset, and these can be extracted using one of the available binning tools (McHardy et al., 2007; Teeling et al., 2004; Tyson et al., 2004). In the metagenomic study of Lake Washington methylotroph populations, a composite genome of M. mobilis totaling slightly over 11 Mb was extracted from the methylamine microcosm metagenome using a compositional binning method PhyloPythia (McHardy et al., 2007; genome statistics are shown in Table 6.1). Genome completeness can be validated by examination of the presence of key metabolic and housekeeping genes (e.g., in Kalyuzhnaya et al., 2008). With satisfactory results, metabolism of an individual organism or a population of closely related strains (which will not necessarily be distinguished between by the

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Table 6.2 Representation of genes involved in tetrahydromethanopterin-linked formaldehyde oxidation in datasets generated in this work, compared to a soil metagenome (Tringe et al., 2005)

Dataset size (Mb)

a

Lake Washington sediment Minnesota farm soil Methane Methanol Methylamine Formaldehyde 100

52

50

37

Gene

Number of copies (coverage score, X)a

16S rRNA fae mtdB/mtdC mch fhcA fhcB fhcC fhcD mptG afp orf5 orf7 orf9 orf17 orf19 orf20 orf21 orf22 orfY

23 (11.5) 3 (1.5) 1 (0.5) 7 (3.5) 5 (2.5) 4 (2.0) 1 (0.5) 2 (1.0) 7 (3.5) 1 (0.5) 4 (2.0) 3 (1.5) 8 (4.0) 3 (6.0) 3 (1.5) 7 (3.5) 6 (3.0) 4 (2.0) 3 (1.5)

12 (13.6) 13 (8.1) 12 (9.7) 2 (2.3) 13 (11.7) 7 (6.3) 5 (2.5) 4 (3.0) 3 (1.5) 3 (1.5) 7 (4.3) 1 (0.5) 7 (5.0) 8 (5.1) 5 (3.8) 10 (7.1) 5 (3.4) 4 (2.0) 5 (3.8)

12 (18.8) 7 (2.9) 6 (3.0) 2 (1.0) 8 (5.1) 6 (5.2) 6 (3.7) 3 (2.2) 7 (4.3) 2 (2.0) 5 (4.2) 3 (1.5) 7 (6.0) 3 (4.1) 0 (0.0) 6 (3.0) 3 (1.5) 2 (1.0) 2 (1.0)

10 (42.5) 27 (45.0) 9 (12.4) 7 (10.1) 16 (18.1) 5 (7.0) 14 (17.6) 16 (21.8) 11 (16.1) 9 (15.7) 8 (13.6) 10 (15.9) 18 (23.5) 10 (13.7) 6 (10.5) 10 (19.1) 7 (20.8) 4 (6.6) 15 (18.9)

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18 (21.4) 11 (5.5) 4 (2.6) 3 (2.4) 6 (5.1) 3 (1.5) 4 (3.2) 4 (2.0) 4 (2.6) 0 (0.0) 2 (1.0) 3 (1.5) 5 (2.5) 2 (1.0) 2 (1.0) 2 (1.7) 2 (1.0) 0 (0.0) 3 (2.4)

Coverage score is calculated as a sum of average contig coverage (X) for each gene. For singleton reads, coverage was arbitrarily counted at 0.5X.

binning methods) can be reconstructed, and genome-wide comparisons may be carried out with other complete or composite genomes. For example, comparing the composite genome of M. mobilis to the complete genome of a close relative M. flagellatus, we were able to uncover examples of highly conserved parts of metabolism, including methylotrophy, as well as of non-conserved parts of metabolism, including non-homologous replacements in common biochemical pathways (Kalyuzhnaya et al., 2008). More recently we obtained the ultimate proof of the precision of the predictions derived from the composite M. mobilis genome (that we dubbed “high-resolution metagenomics”), by completely sequencing a genome of the type strain M. mobilis JLW8 and comparing it to the composite genome extracted from the metagenome (unpublished data).

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Betaproteobacterial Fae

Planctomycetal

Methylamine-specific

Gammaproteobacterial

Fae4

Archaeal

Fae2 Fae3

Figure 6.2 Phylogenetic diversity of Fae peptides identified in metagenomic datasets described in Kalyuzhnaya et al. (2008; only complete or nearly complete sequences were included). Red, methylamine microcosm, green, methane microcosm, blue, methanol microcosm, yellow, formaldehyde microcosm, purple, formate microcosm. Fae, formaldehyde activating enzyme. Fae2-4, homologs of Fae with no demonstrated function. The latter is only found in organisms not possessing H4MPT-linked C1 transfer pathway.

3.3.2. Characterizing genomes at low sequence coverage Binning tools such as PhyloPythia can also be applied to extract less covered genomes. However, in these cases, such approaches will unlikely produce complete genomes of individual organisms or populations. We found it helpful to supplement binning with the so-called protein recruitment technique. In this case a reasonably closely related genome sequence is required, as a reference. A convenient tool for protein recruitment is Phylogenetic Profiler that is part of the IMG/M package. Using the combination of PhyloPythia and Phylogenetic Profiler, we were able to extract an almost complete genome of and reconstruct metabolism of an uncultivated strain of M. tundripaludum and compare it to the genome of Methylococcus capsulatus (Kalyuzhnaya et al., 2008). If a very closely related genome is available as a reference, DNA recruitment can be used in place of protein recruitment (Rusch et al., 2007). Using this technique, we recovered a large portion of a R. eutropha genome (Kalyuzhnaya et al., 2008).

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4. Ultrashort Read-Based Metatranscriptomics 4.1. Principle and strategy For large-scale metatranscriptomics experiments, the next-generation sequencing technologies (Ansorge, 2009; Lapidus, 2009) are especially attractive as assembly is not a prerequisite for transcript analysis. The few metatranscriptomic studies published so far (Frias-Lopez et al., 2008; Gilbert et al., 2008; Urich et al., 2008) employed the 454 sequencing technology, as this technology produces reads of sufficient length to allow for functional predictions based on a single read. These reads are then processed in a genecentric way, overall producing very low resolution data limited to general function assignment for a fraction of reads while many of the reads remain unannotated. A much higher resolution is achieved when the 454 reads can be aligned with a genome of a close relative, presenting a principally different approach, similar to the method that is becoming increasingly popular as applied to single-organism transcriptomics, known as RNA-seq (Sorek and Cossart, 2010; van Vliet, 2009). However, the diversity of most communities is not well covered by sequenced genomes. Our own approach was to match transcripts produced by the Illumina technology (unpaired 75 bp reads) to the existing metagenomic scaffolds derived from the same environment. As the metagenome we employed as a scaffold has been generated from samples specifically enriched for methylotroph functional types (Kalyuzhnaya et al., 2008), as a proof of principle, for the transcriptomics experiments, we carried out enrichments using microcosm incubations, similar to the ones described above. However, only unlabeled substrates were used, and formate was omitted from the transcriptomics experiments. The resulting data were then pooled together. Obviously, the (incomplete) metagenomic scaffolds differ from the (complete, finished) scaffolds representing single genomes, in terms of both sequence coverage and sequence quality. However, we argue that when both a metagenome and a metatranscriptome originate from the same environment/condition, they should have a significant overlap. Most importantly, metatranscriptomes may be repeatedly matched to metagenomes after sequence space expansion (additional sequencing) and/or other iterations, such as an improved assembly.

4.2. RNA isolation The RNA extraction was performed as previously described (Nercessian et al., 2005) with the following modifications. Microcosm samples (0.5 g) were resuspended in 0.75 ml of RNA extraction buffer (0.15 mM NaH2PO4/Na2HPO4 buffer, pH 7.5; 5% CTAB, 1 M NaCl, 2% SDS;

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and 2% N-lauroylsarcosine sodium salt). The DNase I treatment was carried out using the DNAfree kit (BioLabs, Ambion) in accordance with the manufacturer’s instructions. The RNA samples were further purified using the RNeasy columns (Invitrogen, USA). An additional DNase I treatment was carried out directly on the RNeasy columns using the RNase-Free DNase Set (QIAGEN) in accordance with manufacturer’s instructions. The integrity of the RNA preparations was tested on a Bioanalyzer 2100 (Agilent), using Agilent RNA 6000 nano-kit, as suggested by the manufacturer.

4.3. mRNA enrichment and (optional) cDNA synthesis The content of ribosomal RNA was reduced in two steps: first using the MicroExpress Bacterial mRNA Purification Kit (Ambion) and then using the mRNA-ONLYTM Prokaryotic mRNA Isolation Kit (Epicentre Biotechnologies). The resulting mRNA-enriched samples (10 mg) were subjected to first-strand cDNA synthesis using 10 mM random hexamers (Qiagen) and the Omniscript Reverse Transcriptase (Qiagen). The double-stranded DNA was synthesized from single-stranded cDNA using the Exo-Klenow enzyme (Ambion) and standard conditions (Sambrook et al., 1989). The ds-cDNA samples were purified using the QIAquick PCR Purification Kit (Qiagen). The cDNA-synthesis steps can be omitted and the RNA sampled can be submitted to a sequencing facility where a sequencing platform-specific chemistry would be utilized to produce cDNA.

4.4. Data processing The pipeline we used to process and analyze the ultrashort read data is comprised of six steps: (1) the raw reads are subjected to quality assessment and (2) complexity filtering; (3) the reads are then aligned to specific scaffolds in nucleotide space with some allowances for incomplete, inaccurate, or drifting scaffolds; (4) the resulting alignments are postprocessed from the aligner specific format into a unified standard format and (5) imported into a relational database (RDBMS) for unification against scaffold annotations; (6) finally, analyses are performed using a variety of tools and languages including R Development Core Team (2010). 4.4.1. Read quality assessment The quality of raw reads is assessed by examination of two metrics: (a) the frequency of occurrence for the four nucleic acid residues at each position, and (b) the average quality score at each position. In the case of (a), we expect a quality set of reads to have nucleotides A, T, C, and G with populations that roughly agree with the overall GC content of our

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metagenome. In addition, we expect to have very few uncalled bases (denoted by “N” in the read sequence). For (b), we expect the mean base quality across reads to be a monotonically increasing function of read base position. Similar behavior is expected for the quality score standard deviation. 4.4.2. Complexity and quality filtering Reads with low trinucleotide complexity, uncalled bases, and poor quality scores are removed before alignment to a scaffold. The complexity score is defined as the sum of the squares of the trinucleotide frequencies in each read. This metric indicates low complexity (e.g., AAAAAA) with a large complexity score. An upper bound cutoff is determined by manual inspection of the distribution of complexity scores. Reads whose complexity score exceed this cutoff are removed from further processing. 4.4.3. Alignment to scaffold The reads are mapped to a reference scaffolds using BFAST (Homer et al., 2009a,b). BFAST offers substantial control over mismatch detection through its indexing structure. We generated a set of 10 indexes and applied them to our reference scaffolds in the style of what Homer et al. refer to as “accurate” (Homer et al., 2009a). These indexes are predicted to capture 100% of exact and 1 mismatch reads and up to 96% of two mismatch reads. In our case, as a primary reference scaffold, we used the previously described Lake Washington combined metagenomic dataset (JGI Taxon ID 2006543005). As a negative control, we chose the termite hind-gut metagenome ( JGI Taxon ID 2004080001, Warnecke et al., 2007). Below, we refer to “read hits” as the count of all mappings, including instances where a read mapped with equal quality to multiple loci, as in the cases where multiple strains with significant overlap are present in the metagenome. The count of distinct reads mapped is referred to as unique reads and ignores that an individual read may be mapped more than once. 4.4.4. Postprocessing of alignment outputs The resulting BFAST alignments are subsequently filtered to remove overly generous alignments (i.e., those with very many mismatches, insertions, and deletions) by parsing the Concise Idiosyncratic Gapped Alignment Report (CIGAR) strings reported in the output. We considered two filtering schemes where we required the entire read be included in the alignment or 90% of the read to be aligned to the scaffold. The selection of a 90% cutoff for read mapping is based on our understanding of the level of diversity of M. mobilis, a common member of the enriched samples of Lake Washington sediment (up to 20% divergence at the DNA level). Finally, SAMtools (Li et al., 2009) is used to generate “pileup” formatted files that contain the number of reads mapped at each open reading frame (ORF).

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4.4.5. Database integration The scaffold metadata are loaded into a MySQL RDBMS database. These data include unique ORF identifiers assigned by scaffold providers (i.e., JGI; http://img.jgi.doe.gov/cgi-bin/m/main.cgi) and annotations, including the predicted product, predicted transmembrane helices, functional assignments, etc. The “pileup” files from the alignments are read into a specific database and joined to the scaffold annotation data. Finally, the “reads mapped per base” data are rolled up to a single number for each ORF, that is, reads mapped per ORF. 4.4.6. Statistical analysis and visualization We found R to be a convenient environment for computing statistics, producing visualizations/plots, and clustering expression values. R has a rich ecosystem of contributed analyses for bioinformatics, including Bioconductor (Gentleman et al., 2004) and is capable of accessing the data in our RDBMS directly. For visualizing the reads mapped to ORFs, we found Integrative Genomics Viewer (v1.4.2, http://www.broadinstitute.org/igv) suitable, although it required us to view one contig at a time rather than the entire metatranscriptome mapping.

4.5. Metatranscriptome coverage and specificity The number of reads obtained from each enrichment as well as total number of reads are shown in Table 6.3 as are the statistics on how many reads were filtered out. Across all samples, we obtained 66.02 million reads (5 Gb of sequence). The complexity and quality-filtering process on the dataset as a whole removed about 18% of the reads, leaving 54.44 million reads (or 4 Gb) to be aligned to the metagenome. Table 6.4 shows the number of reads mapped and the counts of those mapped uniquely to the reference metagenome of the enriched Lake Washington sediment community (matching scaffold) and to the metagenome of termite hind-gut (negative control scaffold). The conservative Table 6.3 Summary of metatranscriptomic sequencing Reads (millions)

Bases (Gb)

Enrichment

Raw

After filtering (%)

Raw

After filtering

Methane Methanol Methylamine Formaldehyde Total

13.39 16.97 17.44 18.22 66.02

10.05 (75) 14.50 (85) 14.11 (81) 15.49 (85) 54.14 (82)

1 1.27 1.31 1.37 4.95

0.75 1.08 1.06 1.16 4.06

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Table 6.4 Summary of metatranscriptomic reads mapped 90% identity (%)

100% identity (%)

Reads mapped (millions) to Lake Washington reference metagenome Read hits 5.17 (9.5)a 1.82 (3.4)a a Unique reads mapped 4.12 (7.6) 1.31 (2.4)a b Unique reads mapped to CDS 2.49 (60) 0.71 (55)b b Unique reads mapped to rRNA 1.63 (40) 0.60 (45)b 4 b Unique reads mapped to tRNA 6.01  10 (0) 2.54  10 4 (0)b Reads mapped (millions) to negative control metagenome (Termite hind-gut) Read hits 0.80 (1.5)a 0.19 (0.35)a a Unique reads mapped 0.52 (0.96) 0.12 (0.22)a b Unique reads mapped to CDS 0.17 (32) 0.02 (14)b b Unique reads mapped to rRNA 0.36 (68) 0.11 (86)b 4 b Unique reads mapped to tRNA 1.12  10 (0) 8  10 6 (0)b a b

Reads mapped out of total reads. Reads mapped to gene types out of unique reads mapped.

alignment strategy that required 100% of the read to match the metagenome scaffold mapped 3.4% of the reads to the specific scaffold, whereas the more liberal 90% cutoff mapped 9.5% of total reads. In contrast, less than 1% of reads were mapped to a random scaffold (negative control), demonstrating that the availability of a matching metagenome enhances gene identification in a metatranscriptome. It is noteworthy that most of the transcripts with matches in the specific metagenome mapped to protein coding regions (55–60% dependent on the cutoff), while most of the transcripts with matches to the negative control metagenome mapped to rRNA coding genes (68–86%). Table 6.5 shown the number of genes in the specific metagenome and in a negative control metagenome matched to transcripts. A total of 122,984 protein coding genes in the specific metagenome were matched (38% of the protein coding regions identified in the metagenome) while only 5845 (or 7% of the protein coding regions) in the negative control were matched. Transcripts were matched separately to the M. mobilis composite genome that represents the most well covered portion of the Lake Washington metagenome (Kalyuzhnaya et al., 2008). In this case, a significant majority of the genes (up to 91%) were matched with transcripts, at a 90% cutoff. Some of the highly transcribed genes in the M. mobilis composite genome were the known (hexulosephosphate synthase, formaldehyde activating enzyme) or inferred (XoxF) methylotrophy genes and nitrogen metabolism (glutamate synthase, glutamine synthetase) genes, in agreement with prior proteomic analysis (Bosch et al., 2009).

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Table 6.5 Summary of genes matched to the metatranscriptome 90% identity (% of total)

100% identity (% of total)

Genes mapped in Lake Washington metagenome Protein coding 122,984 (38) 79,439 (25) tRNA 365 (18) 175 (9) rRNA 281 (71) 273 (69) Total 123,630 (38) 79,887 (25) Genes mapped in Termite hind-gut metagenome Protein coding 5845 (7) 1227 (1) tRNA 79 (17) 8 (2) rRNA 81 (74) 81 (74) Total 6005 (7) 1316 (2) Genes mapped in Methylotenera mobilis composite genome Protein coding 13,957 (91) 2465 (84) tRNA 68 (47) 36 (25) rRNA 8 (57) 8 (57) Total 14,033 (91) 12,935 (84)

5. Conclusions and Future Perspectives We demonstrate that a functional metagenomics approach involving a specific enrichment step such as SIP can enable detailed analysis of the genomes of environmentally relevant microbes, even if the species in question comprise a minor fraction in a highly complex microbial community. A detailed analysis of the genome of a novel methylotroph, M. mobilis was made possible by this approach (Kalyuzhnaya et al., 2008). A genome of an uncultivated M. tundripaludum was also analyzed in detail, expanding the current genomic knowledge of methane utilizers. We are currently generating additional metagenomic datasets, after methane enrichments, in which the genome of this important organism will be highly covered allowing for assembly and detailed analysis (unpublished data). A metatranscriptomics approach using ultrashort sequence reads that are matched to a specific scaffold originating from the same environment, while still in development, shows promise. With a sufficient sequencing effort, this approach should provide information on major expressed pathways, specific responses to specific stimuli and substrate-specific shifts in gene expression patterns. Ultimately, the success of this approach depends on the quality and the completeness of the available metagenomic scaffolds. The recent advances in sequencing technologies offer a significantly higher throughput and significantly reduced cost of sequencing, setting a stage for much larger, Gb-scale metagenomics projects. The increased

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sequencing effort should result in better sequence coverage and thus in increased resolution of sequence analysis as well as of the derived biological knowledge. However, while scaling up the sequence production using the new technologies is straightforward, new challenges have arisen such as the diminished quality of sequence data and shorter read length, necessitating increasingly complex computation as well as special means for data handling, transfer, and storage and requiring new and advanced computer infrastructures. In the near future, Gb-scale or even Tb-scale metagenomics will become a reality. However, approaches employing function-specific enrichments will remain an important step for connecting a specific function in the environment to specific sequence signatures. The functional metagenomics approach described herein has the potential to be used in a wide variety of ecosystems with a wide variety of labeled substrates, as well as with other types of enrichment, in combination with next-generation sequencing technologies.

ACKNOWLEDGMENTS The authors acknowledge support by the National Science Foundation as part of the Microbial Observatories program (MCB-0604269). The sequencing was provided through the US Department of Energy (DOE) Community Sequencing Program, and the work was performed, in part, under the auspices of the DOE Office of Science, Biological and Environmental Research Program, University of California, Lawrence Livermore National Laboratory, and Los Alamos National Laboratory.

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Genetic Systems for Moderately Halo (alkali)philic Bacteria of the Genus Methylomicrobium David S. Ojala,* David A. C. Beck,*,† and Marina G. Kalyuzhnaya‡ Contents 1. Introduction 2. Methods 2.1. Part 1. Validation and application of broad-host-range vectors in Methylomicrobium spp. 2.2. Part 2. Construction of small promoter-probe and expression vectors for use in Methylomicrobium strains 2.3. Part 3. Gene expression studies 3. Outlook Acknowledgments References

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Abstract Biotechnologies for effective conversion of atmospheric greenhouse gases (CO2 and CH4) into valuable compounds, such as chemical and petrochemical feedstocks or alternative fuels, offer promising new strategies for stabilization of global warming. A novel approach in this field involves the use of methanotrophic bacteria as catalysts for CH4 conversion. In recent years, extremophilic methanotrophic species related to the genus Methylomicrobium have become favorable systems for bioprocess engineering, due to their high growth rates and tolerance of a wide range of environmental conditions and perturbations. While the cultures hold the potential of producing a broader range of chemicals from methane, the biotechnologies are still limited by the lack of reliable genetic approaches for system-level studies and strain engineering. In this chapter, we describe a set of molecular tools for genetic investigation and alteration of the Methylomicrobium spp.

* Department of Chemical Engineering, University of Washington, Seattle, Washington, USA eScience Institute, University of Washington, Seattle, Washington, USA Department of Microbiology, University of Washington, Seattle, Washington, USA

{ {

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00007-3

#

2011 Elsevier Inc. All rights reserved.

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1. Introduction It is well recognized that aerobic methanotrophic bacteria have a great potential in commercial production of fine chemicals from methane. However, the lack of a reliable, well-studied microbial system and a limited set of tools for genetic alteration restrict the application of methanotrophs for biotechnology. Recent efforts in culturing novel methanotrophic species have resulted in isolation and characterization of a variety of novel strains. For example, the long-standing notion that methane utilizers are obligate methylotrophs has been reversed by the identification of facultative methanotrophs, Methylocella spp. (Dedysh et al., 2005; Theisen et al., 2005), and species with unique morphology, such as filamentous Crenotrix/Clonothrix (Stoecker et al., 2006; Vigliotta et al., 2007). In addition, methanotrophy was demonstrated for representatives of the phylum Verrucomicrobia (Dunfield et al., 2007; Hou et al., 2008; Islam et al., 2008; Pol et al., 2007). More recently, it has been confirmed that methane oxidation can be linked to denitrification in the absence of oxygen (Ettwig et al., 2009). A number of extremely thermophilic, psychrophilic, acidophilic, alkaliphilic, and halophilic methanotrophs have also been isolated, thus expanding the physiological range of aerobic methanotrophy (Kalyuzhnaya et al., 2008; Kaluzhnaya et al., 2001; Trotsenko and Murrell, 2008). These microbes with a quite unusual physiology continue to challenge our understanding of biochemistry, biology, and the evolution of methanotrophy as one of the core microbial functions. At the same time, they provide a multitude of potential applications for green technologies ( Jiang et al., 2010; Trotsenko and Khmelenina, 2008). (Halo)alkalotolerant obligate methanotrophic bacteria Methylomicrobium alcaliphilum 20Z and Methylomicrobium buryatense 5G and 5B were isolated from saline soda lakes characterized by dynamic seasonal changes (Kalyuzhnaya et al., 2008; Kaluzhnaya et al., 2001; Khmelenina et al., 1997). These cultures stay active over a wide range of physicochemical parameters (pH, temperature, salinity) and quickly adapt to environmental perturbations. Thus, it is not surprising that Methylomicrobium spp. are becoming the most attractive microbial systems for environmental bioprocess design (Trotsenko and Khmelenina, 2008). In this chapter, we describe a set of molecular tools that were adapted or newly constructed for the investigation and targeted genetic alterations of moderately halo (alkali)philic bacteria of the genus Methylomicrobium spp.

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2. Methods 2.1. Part 1. Validation and application of broad-host-range vectors in Methylomicrobium spp. 2.1.1. Overview of the haloalkali(philic) strains Strains described in this chapter are representatives of the genus Methylomicrobium, belonging to two different species—M. alcaliphilum (20Z) and M. buryatense (5G, 5B) (Kalyuzhnaya et al., 2008; Kaluzhnaya et al., 2001; Khmelenina et al., 1997). These bacteria are typical Type I methanotrophs: they are short rods with Type I ICM, use methane or methanol as carbon and energy sources, but not other carbon compounds tested, and assimilate methane carbon via the RuMP pathway. They use nitrate and ammonium as nitrogen sources. All cultures stay active at a wide range of physicochemical parameters such as pH (7–11), temperature (4–48
C), and salinity (0.1–9%) and quickly adapt to environmental perturbations. The strains are resistant to heat treatment (80
C, 25 min) and desiccation. In addition, all strains contain pMmO (Kapp ¼ 0.9–2 mM). The strain 5G possesses mmoX gene and high activity of sMmO was detected in cells grown in a copper depleted medium (Kalyuzhnaya et al., 2008; Kaluzhnaya et al., 2001). Both strains are capable of producing valuable compounds, such as glutamate, proline, and ectoine (Trotsenko and Khmelenina, 2008). The bacterial strains and growth conditions are presented in Table 7.1. As optimal growth of the culture was observed at high pH, the following adjustments were made to the standard NMS medium (Whittenbury et al., 1970): Ca2þ concentration was lowered to 0.02 g/l to prevent precipitation, pH of the phosphate solution was increased (to 7.2–7.8), and carbonate buffer (0.1 M, pH 9.2–9.5) and NaCl (0.1–9%) were added (Table 7.1). 2.1.2. Routine genetic manipulations DNA from Methylomicrobium strains could be isolated by a standard phenol: chloroform method (Sambrook et al., 1989; Griffiths et al., 2000, summarized in Table 7.2). Commonly used DNA extraction kits, such DNeasy Plant Mini Kit (Qiagen) or UltraCleanÒ Microbial DNA Isolation Kit (MoBio, USA), could be applied for a quick extraction, but they usually yield relatively low amounts of DNA. Most of the genes amplified could be cloned into any commonly used plasmid such as pCR2.1 (Invitrogen) or pDrive (Qiagen). However, these high copy number vectors are not optimal for cloning of methylotrophic genes such as full-size pmoABC, fae, mdh, etc., most likely due to high toxicity of the product to Escherichia coli strains (Semrau et al., 1995). The problem of cloning and subcloning of toxic or unstable genes has been overcome by using low copy number vectors with

Table 7.1 Bacterial strains and growth conditions Growth medium

Source

Reference

NCIMB14124 VKM B-2133

Khmelenina et al., (1997), Kalyuzhnaya et al. (2008)

Strain

Genotype/Description

Methylomicrobium alcaliphilum sp. 20Z Methylomicrobium buryatense spp.

Wild type, ChmR, NalR, GenS, StmS, AmpS, TetS, KanS

NMS

Wild type, ChmR, GenR, StmR, NalS, AmpS, TetS, KanS

NMS

E. coli JM109

recA1 D(lac–proAB) endA1 gyrA96 thi-1 hsdR17 relA1 supE44 F0 [traD36 proAB lacIqZDM15] recA pro hsdR RP4-2-Tc::Mu-Km:: LB (Difco) Lab culture collection Tn7 TmpR, SpcR, StrR LB (Difco) Invitrogen USA F-mcrA D(mrr-hsdRMS-mcrBC)f 80lacZDM15 DlacX74 recA1 araD139 D(ara-leu)7697 galU galK rpsL StrRendA1nupG

E. coli S17-1 One Shot TOP 10

NMS Medium



MgSO4 7H2O CaCl2 6H2O KNO3 NaCl Agar (if added) Nutrient Broth (Difco) Trace element solution



VKM B-2245 (strain 5B) Lab culture collection (strain 5G) LB (Difco) Promega USA

Kaluzhnaya et al. (2001)

Yanish-Perron et al. (1985)

Simon et al. (1984)

Growth Mating Selective Comment

1.00 g 0.02 g 1.00 g 15 g 15 g –

1.00 g 0.02 g 1.00 g 2g 15 g 1.2 g

1.00 g 0.02 g 1.00 g 30 g 15 g 0

2 ml

2 ml

2 ml

Dissolve the ingredients in about 700 ml of distilled water. Add ddH2O to 1 l and sterilize by autoclaving. Selective medium should be supplemented with chloramphenicol (15 mg/ml) and a selective agent (100 mg/ml Amp, 100 mg/ml Kan, and 15 mg/ml Tet). Grow cultures under an atmosphere of methane and air (50:50 by volume). Methanol (0.1%) could be used as an alternative source of carbon/energy. **Sterile phosphate buffer and carbonate buffer solutions should be added to media cooled to room temperature.

dH2O 1l 1l 1l Phosphate buffer 20 ml 50 ml 20 ml solution** 50 ml 5 ml 50 ml Carbonate buffer solution** Phosphate solution (g/l) Dissolve salts in about 800 ml of water, adjust pH to 7.2. Add dH2O to 1 l. KH2PO4 Autoclave the solution. Add 2 ml per 100 ml of NMS. 5.44 Na2HPO4 12H2O 14.34 1 M Carbonate solutions (g/l) Make 1 M solution of NaHCO3 and 1 M solution Na2CO3. Sterilize by filtration. Add 4.5 ml of 1 M NaHCO3 and 0.5 ml of 1 M Na2CO3 per 100 ml NMS. NaHCO3 84.0 Na2CO3 106 Trace element solution (g/l) 0.5, Na2-EDTA; 1, FeSO4 7H2O; 0.75, Fe-EDTA; 0.8, ZnSO4 7H2O; 0.005, MnCl2 4H2O; 0.03, H3BO3; 0.05, CoCl2 6H2O; 0.4, Cu-EDTA; 0.6 CuCl2  2 H2O; 0.002, NiCl2 6H2O; 0.05, Na2MoO4 2H2O















Table 7.2 Protocols for DNA, RNA purifications, and mRNA enrichment from Methylomicrobium spp.

Preparation of genomic DNA

Preparation of RNA samples

 Resuspend cell pellet (2 g) in 5 ml of  Collect sample by centrifugation at

 









10 mM NaCl, 20 mM Tris–HCl (pH 8.0), 1 mM EDTA, 100 mg/ml proteinase K, 50 mg/ml RNase A, and 2% (w/v) SDS. Mix gently and incubate 6 h or overnight at 50
C. Extract the DNA by gentle inversion with an equal volume of phenol: chloroform:isoamyl alcohol pH 8 (25:24:1) for 10 min at room temperature. Centrifuge the DNA at 4000 rpm at 4
C for 15 min and transfer the upper aqueous layer with a widebore tip. Add an equal volume of chloroform: isoamyl alcohol (24:1). Mix gently by inversion for 10 min at room temperature. Centrifuge the DNA at 4000 rpm at 4
C for 15 min and transfer the upper aqueous layer with a widebore tip. Repeat steps 6–7.





  

    

Preparation of enriched mRNA samples for RNA-seq

RNA preparations purified as described in the left column should be concentrated to 5000 rpm for 10–20 min at 4
C. 0.8–1 ug/ml of RNA. Resuspend cell pellet in 0.75 ml of extraction buffer (5% CTAB in 0.8 M  Enrich mRNA in accordance with NaCl and 0.1 M phosphate buffer MicrobExpressTM Bacterial mRNA pH 7.2). purification Kit (Ambion, cat# 1905) with the following modification: the Oligo Mix Transfer samples into a 2 ml screw-cap capturing add 4 ml of biotinylated at 30 tube containing 0.5 g of 0.1 mm zirconia-silica beads (Biospec probes specific to Methylomicrobium products), 75 ml of 10% SDS, 75 ml of rRNAs: (16S-1, CCACTCGTCAGC 10% sodium lauryl sarkosine, and GCCCGA; 16S-2, AATCGC 0.75 ml of phenol:chloroform: TAGTAATCGCGAATC; 23S-1 TAC isoamylic alcohol (25:24:1). TTAGATGTTTCAGTT; 23S-2, CAC TAACTGGGGCTGGACTT; and 5S-1, Homogenize in a mini bead beater for TGGGACACGCTCGCTAT). 1 min. Centrifuge at 14,000 rpm for 5 min at  Resulting RNA samples should be treated 4
C. with the Terminator 5-Phosphate dependent exonuclease (EpiBio, cat# Take the upper aqueous phase and add TER51020). Check the quantity and equal volume of chloroform:isoamylic quality of RNA preparation by a alcohol (24:1). NanoDrop and a Bioanalyzer 2100 Centrifuge at 14,000 rpm for 5 min at (Agilent). Typical results of mRNA 4
C. enrichments are illustrated below: Prepare the tubes for the next steps. Take the upper and add. 0.1 volume of 3 M sodium acetate. 0.8 volume of isopropanol.

 Add 0.1 volume of 0.3 M sodium     

acetate (pH 5.5) and 2 volumes of ethanol on top of the DNA solution. Collect DNA by using a sterile tip, transfer into a new clean tube. Wash the DNA with 70% (v/v) ethanol. Dry sample at RT for 15–20 min. Dissolve the DNA in 5 ml of water. Check the quality of the genomic DNA by electrophoresis through a 0.3% (w/v) agarose gel. Use lambda DNA (50 kb) as size standard. Goodquality DNA runs above or with lambda DNA.

May be used directly for all routine applications such as PCR amplification, digestion, cloning, genome sequencing.

 Incubate 4–6 h or overnight at

Total

After

After

Ladder  80
C. RNA step 1 step 2  Centrifuge at 14,000 rpm for 35–40 min at 4
C.  Wash with 70% (v/v) ethanol.  Dry sample at room temperature for 10 min.  Resuspend in 100 ml of 1 DNase I buffer containing 10 U/ml DNase I (Ambion).  Incubate at 37
C for 30 min.  Purify sample using RNease kit (Invitrogen, cat# 74104) in accordance with RNA Cleanup protocol with oncolumn DNase digestion. RNA before and after enrichment with the  Check the quantity and quality of MicrobExpress kit (step 1) and terminator RNA preparation by a NanoDrop and exonuclease treatment (step 2). a Bioanalyzer 2100 (Agilent), using Agilent RNA 6000 nano kit as suggested by manufacturer. May be used directly for RNA labeling Enriched mRNA sample may be sequenced just as any RNA sample using Illumina for microarray experiments, real-time sequencing. RT-PCR reactions.

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strong terminators flanking the cloning site for elimination of insert transcription, such as pSMART (Lucigen). The system was successfully applied for cloning pmoCAB cluster and individual pmo genes from M. alcaliphilum 20Z (Table 7.3). 2.1.3. Antibiotic resistance markers for genetic manipulations of Methylomicrobium spp. To identify genetic markers suitable for genetic manipulation of Methylomicrobium spp., an antibiotic resistance screening was performed (Table 7.1). Both cultures showed high levels of resistance to chloramphenicol (Chl, 15 mg/ml) and were sensitive to tetracycline (Tet, 15 mg/ml), kanamycin (Kan, 100 mg/ml), and ampicillin (Amp, 100 mg/ml). M. alcaliphilum 20Z was resistant to nalidixic acid (Nal, 30 mg/ml) but sensitive to streptomycin (Stm, 10 mg/ml). Reverse pattern was observed for M. buriatenses strains (NalS, StmR). We did not observe appearance of spontaneous mutants resistant to Tet, Kan, or Amp in any strain tested. Thus, the genetic systems carrying the markers could be applied for genetic manipulations of the Methylomicrobium spp. Chloramphenicol and nalidixic acid (in case of strain 20Z) may serve as selective signatures of the methanotrophic bacteria and could be added to a cultivation medium to eliminate contamination during routine growth in the laboratory. 2.1.4. Efficient conjugal transformation of Methylomicrobium strains A variety of versatile broad-host-range (bhr) and promoter-probe vectors have been previously developed for use in different groups of methylotrophic bacteria (Ali and Murrell, 2009; Chistoserdov et al., 1994; Marx and Lidstrom, 2001, 2002). Some of these tools were successfully applied for genetic manipulation of methanotrophic cultures (Ali and Murrell, 2009; Berson and Lidstrom, 1997; Stolyar et al., 1999). To validate the applicability of the vectors as genetic tools for Methylomicrobium spp., the previously developed bhr vectors, such as an allelic exchange vector pCM184 (Marx and Lidstrom, 2002) and cloning vectors pCM130, pCM66, and pCM62 (Marx and Lidstrom, 2001), were tested for a genetic manipulation of M. alcaliphilum 20Z and M. buryatense 5G. So far, we were not able to develop an efficient electroporation or chemical transformation protocol for the strains. However, we found that plasmid DNA can be introduced into Methylomicrobium spp. via conjugation. An extremophilic lifestyle of the moderately halophilic/alkaliphilic bacteria required a few adjustments for the mating protocol. The optimization of the conjugation was performed by using pCM66-based constructs (Table 7.2). Vectors were introduced into a donor strain E. coli S17-1 via standard transformation procedure (Sambrook et al., 1989). The donor strain grown on LB-agar medium

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Table 7.3 List of plasmids used in this study Plasmids

pDrive pmCherry

Description

Source/Reference r

r

Commercial cloning vector (Amp , Kan ) Prokaryotic expression vector that encodes red fluorescent protein, mCherry mCherry: Excmax ¼ 587 nm; Emmax ¼ 610 nm pEGFP-N1 Gene fusion and expression vector (Neor, Kanr) GFP-N1: Excmax ¼ 488 nm; Emmax ¼ 507 nm pSMART- Low copy (LC) versions of the pSMART LCKan transcription-free cloning vectors (KanR) pTSGex pCM139 with Plac–gfpuv (TetR) pRK2013 pCM130

pCM62 pCM66

pEBPR01 pEBP1 pMOC1 pMO1 pDO1 pDO2 pDO3

pDO4

Helper plasmid expressing IncP tra functions (Kanr) pCM76 with trrnB of E. coli, lowbackground bhr xylE promoter-probe vector (Tetr) Hybrid of pUC19 and pCM51, improved bhr cloning vector (Tetr) Kanamycin cassette inserted into tetA of pCM62, improved bhr cloning vector (Kanr) pCM184 with ectR1 upstream and downstream flanks (KanR, TetR) pTSGex with ectAp1p2–gfpuv (TetR). Gfp: Excmax ¼ 405 nm; Emmax ¼ 509 nm pSMART-LCKan vector harboring pmoCAB cluster pCM184 with upstream and downstream pmoCAB flanks pCM66 with pmo–mCherry fusion cloned between XbaI and EcoRI sites (Kanr) pCM66 with pect–gfp fusion cloned between KpnI and XbaI sites (Kanr) Self-ligation of pDrive with fragment between PvuII sites including lacZ alpha-peptide removed (Ampr, Kanr) pDO3 with csf1 fragment cloned between PvuII sites (Ampr, Kanr)

Qiagen Clontech (USA)

Clontech (USA)

Lucigen Corporation (USA) Strovas and Lidstrom, 2009 Figurski and Helinski (1979) Marx and Lidstrom (2001) Marx and Lidstrom (2001) Marx and Lidstrom (2001) Mustakhimov et al. (2010) Mustakhimov et al. (2010) This study This study This study This study This study

This study (continued)

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Table 7.3 (continued) Plasmids

Description

Source/Reference

pDO5

pDO4 with pmoC cloned between XbaI and KpnI sites (Aapr, Kanr) pDO4 with pect cloned into EcoR1 site (Ampr, Kanr) pDO4 with pect–gfp fusion cloned between XbaI and KpnI sites (Apr, Knr) pDO4 with gfp cloned between KpnI and EcoRI sites (Ampr, Kanr) pDO4 with mCherry cloned between KpnI and EcoRI sites (Ampr, Kanr)

This study

pDO6 pDO7 pDO8 pDO9

This study This study This study This study

supplemented with appropriate antibiotic and the recipient Methylomicrobium strain grown on NMS-agar medium were mixed in a donor:recipient ratio of 1:1, 1:2, 1:5, and 1:10 and plated on the optimized mating medium (Table 7.1). Plates were incubated at 30
C under methane:air atmosphere (25:75) for 24, 48, 72, and 120 h, and cells were transferred from a mating medium onto selective plates. Chloramphenicol, high pH, and 3% salinity were applied for counter-selection against the donor cells. Kanamycinresistant clones were generated, demonstrating that the cloning and promoter-probe vector could be transferred into Methylomicrobium spp. cells. The optimal donor:recipient ratio was found to be 1:2, and the optimal conjugation time was 48 h. A triparental mating, with a donor, recipient, and helper strain (like pRK2013), could facilitate the plasmid transfer into the methanotrophic strains. 2.1.5. Construction of M. alcaliphilum 20Z mutant strains Empty pCM184, an allelic exchange vector, was also tested for genetic manipulations in M. alcaliphilum. We used pCM184 and pMO1 (a pCM184based vector carrying upstream and downstream flanks of the pmo gene cluster). The plasmids were introduced into wild type of M. alcaliphilum 20Z via conjugation procedure described above. As expected, no recombinants were obtained for the empty suicide vector; however, KanR-clones were generated for the pMO1 system. The recombinants were further tested for resistance to tetracycline, and Tet-sensitive (TetS) mutants were chosen as possible doublecrossover recombinants. The identity of the double-crossover mutants was verified by diagnostic PCR with primers specific to the insertion sites. The pmoCAB-lacking strains have impaired growth on methanol (five times slower than wild type) and as expected were not able to utilize methane. A similar

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strategy was used to generate M. alcaliphilum mutant strain lacking the ectR gene, a regulator of ectoine biosynthesis (Mustakhimov et al., 2010). These data confirmed the utility of the allelic exchange vector for targeted gene manipulation in Methylomicrobium species. 2.1.6. Validation of the pCM66 plasmid-based promoter-probe vector pTSGex, a pCM66-based Gfp promoter-probe vector (Strovas and Lidstrom, 2009), was used for investigation of in vivo transcriptional regulation of the ectoine biosynthesis pathway (Mustakhimov et al., 2010). The 352 bp fragment containing ectR-operon promoter region (ectAp1p2) was amplified by PCR and cloned into the pCR2.1 vector. The fragments were subsequently excised by PstI and BamHI and cloned into the pTSGex vector, resulting in the pEBP1 that contains the respective DNA fragments upstream of the promoterless reporter gene gfp. The resulting construct was transformed into E. coli S17-1 and then transferred into M. alcaliphilum 20Z using the conjugation procedure described above. Tetracycline-resistant transconjugants were grown at different salinity (1%, 3%, and 6%) and assayed for gfp expression. Cells grown at high salinity yielded higher level of fluorescence as a result of the ectAp1p2 promoter activation (Fig. 7.1; Mustakhimov et al., 2010). The data demonstrate that the Gfp probe is a useful tool for assessing promoter activity in Methylomicrobium spp. RFU 0

50

100

150

200

6% NaCl

3% NaCl

1% NaCl

Figure 7.1 Activities of the ectAp1p2–gfp promoter fusion in cells of M. alcaliphilum 20Z grown under different salinity conditions. RFU: relative fluorescence unit. Fluorescence measurements were carried out with a Shimadzu RF-5301PC fluorimeter. GFPuv excitation was conducted at 405 nm and emissions were monitored at 509 nm. Promoter activities were calculated by plotting fluorescence versus OD600.

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2.2. Part 2. Construction of small promoter-probe and expression vectors for use in Methylomicrobium strains 2.2.1. Construction of pDO4-based promoter-probe vector Most of the currently available bhr vectors for use in methylotrophic bacteria are relatively large in size (8–12 kb). It is well recognized that plasmid maintenance may influence cell growth and productivity. To characterize gene expression profiles across environmental perturbations that significantly differ in energy requirements, for example, growth at low salinity versus high salinity, the use of an integrative promoter probe or small plasmids seems to be more biologically relevant. This way, less cellular energy would be spent supporting plasmid replication. Integrative promoter probes are ideal for in vivo assay of transcriptional activities; however, the system must be specific for a given host. To develop a probe that could be used in different species of Methylomicrobium, we used the pDrive (Qiagen, USA), a small 3.8 kb pUCbased cloning vector. We found that the vector could be introduced into different Methylomicrobium strains via conjugation. To generate a small, low-background, versatile promoter-probe vector, the commercial cloning vector was cut with PvuII to remove a fragment containing the lacZ alpha-peptide and Plac and T7 RNA polymerase promoters. The resulting vector was then self-ligated to produce pDO3 (Table 7.3). A synthetic multiple cloning site segment (CSF1), containing two terminal PvuII restriction sites and HindII/SacII/SacI/EcoRI/KpnI/XhoI/XbaI/NcoI/NdeI cloning sites, was ligated into PvuII site of pDO3 to produce pDO4 (Fig. 7.2). The resulting plasmid was sequenced and used as a base for developing cloning vectors described below. Two low-background promoter-probe vectors were constructed with either green (Gfp-N1) or red (mCherry) fluorescent proteins as reporter genes. We used modified Gfp protein, as it has been optimized for brighter fluorescence (Cormack et al., 1996). To generate gfp-based promoter-probe construct, the 0.72 kb gfp gene was amplified from pEGFP-N1 vector (Clontech) and cloned into KpnI/EcoRI sites of pDO4 to produce pDO8 (Fig. 7.2). The second reporter fusion system is based on a red fluorescent protein, mCherry. The 0.71 kb mCherry fragment cut with KpnI–EcoRI was cloned into pDO4 cut with KpnI and EcoRI to produce pDO9 (Table 7.2, Fig. 7.2). Both constructs could be cloned and maintained in Methylomicrobium spp. 2.2.2. Construction of pDO4-based expression vectors for use in M. alcaliphilum 20Z Two complementary strategies were used for construction of the gene expression vectors for use in M. alcaliphilum 20Z: (1) design a vector that could grant a high constitutive expression of the target, and (2) design a vector that could provide a tunable expression of the targeted genes. It is

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HindIII (243) SacII (252) pUC ori

SacI (259) EcoRI (261) KpnI (271) XhoI (273) XbaI (279)

pDO4 KAN

NcoI (285)

3451 bp

NdeI (292)

Beta-lactamase

pUC ori

HindIII (243)

pUC ori

SacII (252)

ect promoter

SacI (259)

KAN

KpnI (659) XhoI (661) XbaI (667) NcoI (673)

pDO6 3839 bp

EcoRI (261)

KAN

pDO8

Gfp

4365 bp

Beta-lactamase Beta-lactamase pUC ori

SacII (252) SacI (259) KpnI (271) PstI (284)

pUC ori

mCherry

BamHI (300) KAN

KAN

pDO5 4348 bp

Beta-lactamase

pmo promoter

Expression vectors

pDO9 4187 bp

Beta-lactamase

KpnI (1007) XhoI (1009) NdeI (1028)

Reporters

Figure 7.2 Plasmid maps depicting the relevant features of the low-background promoter-probe vectors (pDO8 and pDO9) and expression vectors (pDO5 and pDO6).

well known that pmo genes are highly expressed genes in methanotrophic cultures (Ali and Murrell, 2009). According to transcriptomic data presented below, the pmoCAB genes are highly expressed in M. alcaliphilum 20Z. The 0.8 kb fragment flanking 818 to 10 (relative to the putative translational start) upstream region of pmoC gene was amplified, cloned into pDrive vector, and subcloned into XbaI–KpnI sites of pDO4 to produce expression vector pDO5 (Fig. 7.2).

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We used the ectAp1p2 promoter regions to a generate system for tunable expression of the targeted genes. The promoters are similar to the s70dependent promoters of E. coli. The ectP1P2 are well-characterized regulatory elements that facilitate the transcription of osmoprotector biosynthesis genes in M. alcaliphilum 20Z in response to osmotic stress (Fig. 7.1; Mustakhimov et al., 2010). The 0.28 kb ectAp1p2 was cloned into the EcoRI site of pDO4 to produce the inducible expression vector pDO6 (Fig. 7.2). Both pDO5 and pDO6 vectors contain the high copy number pUC origin of replication and produce a relatively high yield of DNA. These systems are efficient in M. alcaliphilum strain 20Z only (data not shown).

2.3. Part 3. Gene expression studies Genome sequencing of M. alcaliphilum 20Z has been carried out by Genoscope (https://www.genoscope.cns.fr/agc/mage). This genomic information could be used to gain insights into the regulatory and metabolic networks of the microbe and to interpret the cellular behavior under a range of growth conditions. Global comparative gene expression analysis is one of the most efficient ways of refining the cellular functions in a high throughput mode. In this part of the chapter, we focus on microarray-based characterization of M. alcaliphilum 20Z transcriptome. However, instead of hybridization on a microarray, the enriched RNA samples could be sequenced. A protocol for efficient RNA preparation from Methylomicrobium strains is shown in Table 7.2. Enriched mRNA samples could be sequenced just as any RNA sample using Illumina/Solexa, ABI/SOLiD, or Roche 454 sequencing platforms. The bioinformatics tools described for metatranscriptomics (this book, by Kalyuzhnaya et al., 2011) could be adapted and applied to Methylomicrobium spp. The single organism RNAseq experiment is simpler than the metatranscriptomic variant in that the reference scaffold consists only of one genome, that is, 20Z or 5G. In addition, as the sequenced genome is from the same direct source as the transcriptome, only 100% identical alignments of the sequencer reads to the reference scaffold need be considered. 2.3.1. Microarray design The draft genome for M. alcaliphilum 20Z (http://www.genoscope.cns.fr) was used to construct the microarray. All ORFs annotated as protein coding were targeted for detection on the array. Probe oligos were selected using Agilent’s eArray Web-based tool (http://www.genomics.agilent.com). Probe selection criteria were based on optimizing the Tm for 80
C while selecting three or four of the best sense strand probes for every predicted ORF. For 10 ORFs, it was not possible to generate probes of sufficient uniqueness or quality. These ORFs were annotated as multiple fragments of transposase and integrase catalytic region, and three hypothetical proteins of

Genetic Tools for Methylomicrobium spp.

113

unknown function. Through this process, we obtained 12,463 probes. To fill out the array’s features, 65 ORFs with relevant annotations were more deeply sampled by selecting additional probes with four copies of each probe placed over the array. As a result, 2560 probes were replicated across the array. Agilent internal controls made up 536 additional features. The final feature layout was generated randomly. The microarray design identifier assigned by Agilent was 027442. 2.3.2. RNA isolation and labeling Cultures were grown at optimal conditions (3% NaCl, pH 9.2) and at low (0.1% NaCl) or high salinity (9% NaCl). Two biological replicates per growth variants were set up. When the cultures reached OD ¼ 0.3–0.4, a stop solution (5% water saturated phenol in ethanol) was added up to 1/10 of the culture volume and cells were harvested by centrifugation. The optimized RNA extraction protocol is shown in Table 7.2. RNAse kit (Qiagen) could be used alone; however, it led to drop in RNA yield up to 10-fold. In this chapter, the labeling of purified RNA samples, hybridization, scanning of the microarrays, feature extraction and a basic analysis were performed by the MoGen, LC (http://www.mogene.com/). However, all these steps could be run in a laboratory. Labeled cDNA samples could be prepared by using the SuperScriptTM Plus Indirect cDNA Labeling System (Invitrogen). The resulting samples were hybridized to the Agilent 027442 oligonucleotide microarrays and scanned with an Agilent Scanner. The scanner images were processed by using Agilent Feature Extraction software (Agilent, USA) or GenPix software (Molecular Devices, USA). All these steps and the data processing strategies were explicitly described (Kalyuzhnaya et al., 2008; Knudsen, 2004; Okubo et al., 2007). 2.3.3. Validation of microarray platform To validate the custom oligonucleotide microarray, a set of experiments was designed: two technical replicates and two biological replicates (to assess reproducibility), and dye-swap experiment (to assess signal correlation between Cy5 and Cy3 dyes). Figures 7.3 and 7.4 show examples of scatter plots obtained for dye-swap experiment (self–self hybridization) and two independent microarray experiments, respectively. The correlation of log ratios in all control experiments was good, indicating that the labeling procedure, dye normalization, and reproducibility of arrays are satisfactory. Some results of comparative expressional analyses of cells grown at low salinity versus high salinity are listed in Table 7.4. During growth at an elevated salinity, cells of M. alcaliphilum significantly altered expression of approximately 340 genes (200 upregulated, 140 downregulated). A significant fraction of these genes were hypothetical. However, a few correlations for well-studied functions could be made. As expected, genes essential for operation of ectoine biosynthesis pathway were upregulated (up to three- to

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18 16

Log2 (Cy5)

14 12 10 8 6 4 2 0

0

5

10 Log2 (Cy3)

15

20

Figure 7.3 Visualization of normalized dye-swap microarray data. Log2 ratios from Cy3 versus Cy5. 1.2 1 0.8 0.6 0.4 0.2 -1

-0.5

0 -0.2

0

0.5

1

1.5

-0.4 -0.6 -0.8

Figure 7.4 Comparison of Log2 ratio for two biological replicates. Log ratios from methane versus methanol growth data set were plotted against each other.

fivefold) in cells grown at 9% NaCl compared to growth at 3% or 0.1% NaCl. These data correlate well with the promoter-probe-based analyses of the ect genes transcription. It was known that cells of M. alcaliphilum grown at high salinity have higher methane oxidation potential (Khmelenina et al., 1999). Indeed, primary methane oxidation systems (pmo genes) were significantly upregulated in cells grown at high salinity. Overall, the results presented herein validate the design and implementation of the first genome-wide microarray platform for a methanotrophic culture. Further experiments will focus on elucidation of specific cellular functions of newly identified genes.

Table 7.4

A subset of differentially expressed genes during growth at high salinity (9%) versus low salinity (0.1%)

Gene ID

Functional annotation

MEALZv2_350065 MEALZv2_560038 MEALZv2_560040 MEALZv2_90002 MEALZv2_640009 MEALZv2_640011 MEALZv2_640010 MEALZv2_150013 MEALZv2_900047 MEALZv2_1120112 MEALZv2_1120108 MEALZv2_1120109 MEALZv2_920013 MEALZv2_60014 MEALZv2_1030034 MEALZv2_430028 MEALZv2_240004

Conserved membrane protein of unknown function pmoB, methane monooxygenase pmoC, methane monooxygenase xoxF, putative dehydrogenase fadA, acetyl-CoA acyltransferase fadE, acyl coenzyme A dehydrogenase phbB, acetoacetyl-CoA reductase feoB, ferrous iron transport protein B Naþ/Picotransporter ectA, L-2,4-diaminobutyric acid acetyltransferase ectD, ectoine hydroxylase ask, Aspartokinase Transcriptional regulator, TetR family maxF, methanol dehydrogenase subunit corA, copper-repressible polypeptide nirB, nitrite reductase, large subunit, NAD(P)H-binding glnK, nitrogen regulatory protein

Fold change

18.93 5.78 3.03 7.04 8.99 8.61 5.99 5.11 5.27 5.03 4.05 3.13  25.12  5.43  3.47  3.08  3.00

P value

6.74 3.87 9.72 1.98 1.81 3.29 2.49 6.29 7.24 1.27 4.53 6.05 8.67 1.02 3.70 2.35 3.07

                

10 21 10 33 10 23 10 17 10 18 10 16 10 16 10 16 10 15 10 33 10 35 10 24 10 22 10 15 10 28 10 23 10 10

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3. Outlook Methane-oxidizing bacteria are systems selected by nature for attenuation of methane emission. The biotechnological potential of this group of bacteria has been extensively discussed, but, for years, the actual application of methane oxidizers for industrial needs (such as methanol production and biosynthesis from methane) or bioremediation (TCE degradation) was limited, as no robust, stable, and predictable system has been developed. Most representatives of the genus Methylomicrobium are methane-oxidizing bacteria isolated from saline environments. Physiological properties of these strains make them unique targets for the development of green technologies based on methane as a feedstock, with a focus on efficient utilization of bio-methane from waste sources. However, to establish a robust microbial system for industrial production of chemicals from renewable sources of methane, a multipronged approach is needed. In this chapter, we described a variety of genetic tools that could be applied for investigation of Methylomicrobium spp. Most of our methods were designed and optimized for use in M. alcaliphilum 20Z, as a draft genome of the culture is available. Similar approaches could be elaborated for genetic characterization of other representatives of the genus. Genomes of two representatives of family Methylomicrobium: M. album BG8 and M. buryatense 5G are currently in the process of being sequenced at the DOE-JGI facilities (USA). These additional genomes, once available, will open new avenues for comparative analyses and gene mining.

ACKNOWLEDGMENTS This work was supported by grants CRDF Rub1-2946-PU-09 and NSF Grant MCB0604269.

REFERENCES Ali, H., and Murrell, J. C. (2009). Development and validation of promoter-probe vectors for the study of methane monooxygenase gene expression in Methylococcus capsulatus Bath. Microbiology 155, 761–771. Berson, O., and Lidstrom, M. E. (1997). Cloning and characterization of corA, a gene encoding a copper-repressible polypeptide in the type I methanotroph, Methylomicrobium albus BG8. FEMS Microbiol. Lett. 148, 169–174. Chistoserdov, A. Y., Chistoserdova, L. V., McIntire, W. S., and Lidstrom, M. E. (1994). Genetic organization of the mau gene cluster in Methylobacterium extorquens AM1: Complete nucleotide sequence and generation and characteristics of mau mutants. J. Bacteriol. 176, 4052–4065.

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Cormack, B., Valdivia, R., and Falkow, S. (1996). FACS-optimized mutants of the green fluorescent protein (GFP). Gene 173, 33–38. Dedysh, S. N., Knief, C., and Dunfield, P. F. (2005). Methylocella species are facultatively methanotrophic. J. Bacteriol. 187, 4665–4670. Dunfield, P. F., Yuryev, A., Senin, P., Smirnova, A. V., Stott, M. B., Hou, S., Ly, B., Saw, J. H., Zhou, Z., Ren, Y., Wang, J., Mountain, B. W., et al. (2007). Methane oxidation by an extremely acidophilic bacterium of the phylum Verrucomicrobia. Nature 450, 879–882. Ettwig, K. F., van Alen, T., van de Pas-Schoonen, K. T., Jetten, M. S. M., and Strous, M. (2009). Enrichment and molecular detection of denitrifying methanotrophic bacteria of the NC10 Phylum. Appl. Environ. Microbiol. 75, 3656–3662. Figurski, D. H., and Helinski, D. R. (1979). Replication of an origin-containing derivative of plasmid RK2 dependent on a plasmid function provided in trans. Proc. Natl. Acad. Sci. USA 76, 1648–1652. Griffiths, R. I., Whiteley, A. S., O’Donnell, A. G., and Bailey, M. J. (2000). Rapid method for coextraction of DNA and RNA from natural environments for analysis of ribosomal DNA- and rRNA-based microbial community composition. Appl. Environ. Microb. 66, 5488–5491. Hou, S., Makarova, K. S., Saw, J. H., Senin, P., Ly, B. V., Zhou, Z., Ren, Y., Wang, J., Galperin, M. Y., Omelchenko, M. V., Wolf, Y. I., Yutin, N., et al. (2008). Complete genome sequence of the extremely acidophilic methanotroph isolate V4, Methylacidiphilum infernorum, a representative of the bacterial phylum Verrucomicrobia. Biol. Direct 3, 2610.1186/1745-6150-3-26. Islam, T., Jensen, S., Reigstad, L. J., Larsen, O., and Birkeland, N. K. (2008). Methane oxidation at 55 degrees C and pH 2 by a thermoacidophilic bacterium belonging to the Verrucomicrobia phylum. Proc. Natl Acad. Sci. USA 105, 300–304. Jiang, H., Chen, Y., Jiang, P., Zhang, C., Smith, T. J., Murrell, J. C., and Xing, X. H. (2010). Methanotrophs: Multifunctional bacteria with promising applications in environmental bioengineering. Biochem. Eng. J. 49, 277–288. Kaluzhnaya, M., Khmelenina, V., Eshinimaev, B., Suzina, N., Nikitin, D., Solonin, A., Lin, J.-L., McDonald, I., Murrell, C., and Trotsenko, Y. (2001). Taxonomic characterization of new alkaliphilic and alkalitolerant methanotrophs from soda lakes of the Southeastern Transbaikal region and description of Methylomicrobium buryatense sp. nov. Syst. Appl. Microbiol. 24, 166–176. Kalyuzhnaya, M. G., Khmelenina, V., Eshinimaev, B. T., Sorokin, D. Yu., Fuse, H., Lidstrom, M. E., and Trotsenko, Y. A. (2008). Reclassification and emended description of halo(alkali)philic and halo(alkali)tolerant methanotrophs of the genera Methylomicrobium and Methylobacter. Int. J. Syst. Evol. Microbiol. 58, 591–596. Kalyuzhnaya, M. G., Beck, D. A. C., and Chistoserdova, L. (2011). Functional metagenomics of methylotrophs. Methods Enzymol. 495, 81–98. Khmelenina, V. N., Kalyuzhnaya, M. G., Starostina, N. G., Suzina, N. E., and Trotsenko, Yu.A. (1997). Isolation and characterization of halotolerant alkaliphilic methanotrophic bacteria from Tuva soda lakes. Curr. Microbiol. 35, 257–261. Khmelenina, V. N., Kalyuzhnaya, M. G., Sakharovsky, V. G., Suzina, N. E., Trotsenko, Yu.A., and Gottschalk, G. (1999). Osmoadaptation in halophilic and alcaliphilic methanotrophs. Arch. Microbiol. 172, 321–329. Knudsen, S. (2004). Guide to Analysis of DNA Microarray Data. 2nd edn. John Wiley & Sons, Inc., Hoboken, New Jersey. Marx, C. J., and Lidstrom, M. E. (2001). Development of improved versatile broad-hostrange vectors for use in methylotrophs and other Gram-negative bacteria. Microbiology 147, 2065–2075.

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Marx, C. J., and Lidstrom, M. E. (2002). Broad-host-range cre-lox system for antibiotic marker recycling in gram-negative bacteria. Biotechniques 33, 1062–1067. Mustakhimov, I. I., Reshetnikov, A. S., Glukhov, A. S., Khmelenina, V. N., Kalyuzhnaya, M. G., and Trotsenko, Y. A. (2010). Identification and characterization of EctR, a new transcriptional regulator of the ectoine biosynthesis genes in the halotolerant methanotroph Methylomicrobium alcaliphilum 20Z. J. Bacteriol. 192, 410–417. Okubo, Y., Skovran, E., Guo, X., Sivam, D., and Lidstrom, M. E. (2007). Implementation of microarrays for Methylobacterium extorquens AM1. OMICS 11, 325–340. Pol, A., Heijmans, K., Harhangi, H. R., Tedesco, D., Jetten, M. S., and Op den Camp, H. J. (2007). Methanotrophy below pH 1 by a new Verrucomicrobia species. Nature 450, 874–878. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual. 2nd edn. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Semrau, J. D., Chistoserdov, A., Lebron, J., Costello, A., Davagnino, J., Kenna, E., Holmes, A. J., Finch, R. J., Murrell, C., and Lidstrom, M. E. (1995). Particulate methane monooxygenase genes in methanotrophs. J. Bacteriol. 177, 3071–3079. Simon, R., Priefer, U., and Pu¨hler, A. (1984). A broad host range mobilization system for in vivo genetic engineering: Transposon mutagenesis in Gram-negative bacteria. Biotechnology 1, 784–791. Stoecker, K., Bendinger, B., Scho¨ning, B., Nielsen, P. H., Nielsen, J. L., Baranyi, C., Toenshoff, E. R., Daims, H., and Wagner, M. (2006). Cohn’s Crenothrix is a filamentous methane oxidizer with an unusual methane monooxygenase. PNAS 103, 2363–2367. Stolyar, S., Costello, A. M., Peeples, T. L., and Lidstrom, M. E. (1999). Role of multiple gene copies in particulate methane monooxygenase activity in the methane-oxidizing bacterium Methylococcus capsulatus Bath. Microbiology 145, 1235–1244. Strovas, T. J., and Lidstrom, M. E. (2009). Population heterogeneity in Methylobacterium extorquens AM1. Microbiology 155, 2040–2048. Theisen, A. R., Ali, M. H., Radajewski, S., Dumont, M. G., Dunfield, P. F., McDonald, I. R., Dedysh, S. N., Miguez, C. B., and Murrell, J. C. (2005). Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol. Microbiol. 58, 682–692. Trotsenko, Y. A., and Khmelenina, V. N. (2008). Extremophilic Methanotrophs. ONTI PSC RAS, pp. 206. Pushchino, Russia. Trotsenko, Y. A., and Murrell, J. C. (2008). Metabolic aspects of aerobic obligate methanotrophy. Adv. Appl. Microbiol. 63, 183–229. Vigliotta, G., Nutricati, E., Carata, E., Tredici, S. M., De Stefano, M., Pontieri, P., Massardo, D. R., Prati, M. V., De Bellis, L., and Alifano, P. (2007). Clonothrix fusca Roze 1896, a filamentous, sheathed, methanotrophic g-Proteobacterium. Appl. Environ. Microbiol. 73, 3556–3565. Whittenbury, R., Phillips, K. C., and Wilkinson, J. F. (1970). Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61, 205–218. Yanish-Perron, C., Vieira, J., and Messing, J. (1985). Improved M13 phage cloning vectors and host strains: Nucleotide sequences of the M13 mp19 and pUC19 vectors. Gene 33, 103–119.

C H A P T E R

E I G H T

Development of a System for Genetic Manipulation of the Facultative Methanotroph Methylocella silvestris BL2 Andrew Crombie and J. Colin Murrell Contents 1. 2. 3. 4. 5.

Introduction Growth of M. silvestris Introduction of Plasmid DNA by Conjugation and Electroporation Gene Deletion by Electroporation of Linear DNA Preparation of Competent Cells for Electroporation of M. silvestris 6. Construction of Linear DNA for Gene Deletion by Homologous Recombination 7. Electroporation 8. Efficiency of Gene Deletion 9. Case Study: Deletion of Isocitrate Lyase 10. Complementation 11. Phenotype of the Isocitrate Lyase Deletion Mutant 12. Conclusions References

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Abstract An understanding of the metabolism and metabolic regulation of the facultative methanotroph Methylocella silvestris BL2 is required to understand its role in methane oxidation in the environment, and methods for genetics manipulation are essential tools in these investigations. In addition, the ability to engineer the metabolic capabilities of M. silvestris may well have useful biotechnological applications. We describe a simple and effective method of genetic manipulation for this organism which relies on the electroporation of a linear DNA fragment to introduce chromosomal gene deletions. In a two-step procedure, the gene of interest is first replaced with an antibiotic-resistance cassette which School of Life Sciences, University of Warwick, Coventry, United Kingdom Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00008-5

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2011 Elsevier Inc. All rights reserved.

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is subsequently removed, resulting in an unmarked gene deletion. This method is illustrated by the deletion of isocitrate lyase, which abolished growth on onecarbon and severely disabled growth on two-carbon compounds. Subsequent complementation with the wild-type gene and promoter restored growth, demonstrating stable transcription from the broad-host-range plasmid employed.

1. Introduction Methylocella silvestris BL2 is a moderately acidophilic methanotroph originally isolated from a forest soil in Germany (Dunfield et al., 2003). Although phylogenetically related to other Alphaproteobacterial methanotrophs of the genera Methylocystis and Methylosinus, on the basis of 16S rRNA gene sequences, it is most closely affiliated to representatives of the Beijerinckia. As a methanotroph, it plays its part in methane cycling, and an understanding of its occurrence and behavior is thus important from the perspective of global warming and climate change. Previously, methanotrophy was considered an obligate trait (reviewed by Theisen and Murrell, 2005); however, recently it was shown that M. silvestris is also capable of growth on a variety of multicarbon compounds including, for example, acetate, ethanol, propionate, and succinate (Dedysh et al., 2005). Over the past few years, other methanotrophs capable of growth on acetate and ethanol have been described (Belova et al., 2010; Dunfield et al., 2010; Im et al., 2010) but M. silvestris remains the only characterized methanotroph capable of robust growth on a comparatively wide range of multicarbon compounds. M. silvestris is also unusual in not possessing a membranebound form of the methane monooxygenase (MMO), the initial enzyme of the methane oxidation pathway. Whereas all other known methanotrophs possess this particulate form (pMMO), and a few also possess a cytoplasmic soluble form (sMMO), M. silvestris is unique in using only the sMMO. Since this enzyme has a wide substrate specificity and is able, for example, to co-oxidize environmental pollutants including aromatics and halogenated hydrocarbons (Colby et al., 1977), it has attracted considerable interest for its potential use in bioremediation. Whereas obligate methanotrophs are by definition unable to obtain energy and biomass heterotrophically, in appropriate circumstances, it might be possible for M. silvestris to oxidize recalcitrant organic chemicals in soils while using an alternative carbon source for growth. The genome sequence of M. silvestris has recently become available (Chen et al., 2010). This confirmed the absence of the pMMO, but unexpectedly revealed a second soluble diiron monooxygenase in addition to the sMMO, which was provisionally identified as a propane

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monooxygenase. Serine cycle genes for one-carbon assimilation were found, confirming earlier predictions (Dunfield et al., 2003). The presence of glyoxylate bypass genes isocitrate lyase and malate synthase (although not located close to each other on the chromosome) suggested their involvement in anaplerosis during growth on two-carbon compounds. The absence of genes with high similarity to those of the alternative pathway of glyoxylate regeneration recently described by Erb et al. (2009) suggested that isocitrate lyase may also be required for glyoxylate regeneration during one-carbon growth using the serine cycle. These data, and the ongoing debate regarding the reasons for obligate methanotrophy (reviewed by Wood et al., 2004), suggested numerous metabolic and regulatory hypotheses which required confirmation in this organism using biochemical and genetic methods. In order to use the methods of reverse genetics, we attempted to develop an efficient method of targeted mutagenesis, which would also be a prerequisite for any attempt to engineer a strain with the aim of maximizing this organism’s capacity for effective bioremediation or biotransformation. In the laboratory, it is possible to grow M. silvestris on methane in batch culture with a generation time of approximately 35 h (Theisen et al., 2005), or somewhat more quickly using methanol or ethanol as growth substrates. On agar plates, colonies appear after 2–4 weeks. Growth in liquid and on plates is often accompanied by the production of a large amount of polysaccharide slime, in common with other members of the Beijerinckiaceae (Becking, 2006), which hinders transfers and manipulations. This, and the comparatively slow growth, necessitates care to maintain contaminationfree cultures. The most common method of introducing DNA into methanotrophs and methylotrophs has historically been by conjugation (Murrell, 1992), although electroporation has also sometimes been used (Baani and Liesack, 2008; Kim and Wood, 1998; Toyama et al., 1998). For marker-exchange mutagenesis using homologous recombination, two recombination events are necessary, upstream and downstream of the gene of interest, to replace it with a selectable marker, for example, an antibiotic cassette. When introducing DNA on a circular plasmid, the detection of the comparatively rare second recombination event, and consequent loss of the vector backbone, typically requires screening for double-crossover colonies using sensitivity to an antibiotic, resistance to which is encoded on the vector backbone, as indicator. Although the loss of the plasmid backbone may be forced by incorporation of a counter-selectable marker, for example, the sacB gene from Bacillus subtilis (Scha¨fer et al., 1994), a (possibly large) proportion of single crossovers will recombine to wild type unless selective pressure exists. If the intention is to construct a mutant with a deletion of two or more genes, it is convenient if the gene is deleted without incorporation of an antibiotic selectable marker. This is also desirable if the organism is destined

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for release into the environment, in order to prevent the possible horizontal transfer of antibiotic-resistance genes (Davison, 2005). Most of the genetic methods in common use with methanotrophs therefore require timeconsuming screening and numerous transfers of colonies on plates. The introduction of linear DNA, comprising an antibiotic-resistance cassette flanked by regions homologous to sequences upstream and downstream of the gene of interest, requires two simultaneous recombination events if the organism is to gain resistance to the selective antibiotic. Thus, marker-exchange gene replacement is achieved in one operation. This approach, although common in yeast (Rothstein et al., 1983), has less commonly been used in bacteria, for example, in Bordetella pertussis (Zealey et al., 1990), Escherichia coli (El Karoui et al., 1999; Jasin and Schimmel, 1984), Haemophilus ducreyi (Hansen et al., 1992), Methylobacterium extorquens (Toyama et al., 1998), Rickettsia prowazekii (Driskell et al., 2009), and Streptomyces coelicolor (Oh and Chater, 1997), but in most cases, the frequency of gene replacements is low (unless one of a number of methods has been used to modify the host cells to be more receptive to incoming DNA (see, e.g., Murphy, 1998), a strategy usually only available with organisms already engineered for that purpose). Incorporation of specific DNA sequences adjacent to the antibiotic-resistance cassette allows subsequent removal of the cassette by site-specific recombination, using, for example, the Flp-FRT or Cre-loxP recombinase systems (Ayres et al., 1993; Hoang et al., 1998), resulting in unmarked gene deletions. Here we describe an efficient method of gene deletion for M. silvestris and outline, as an example, the removal from the chromosome of the gene encoding the glyoxylate bypass enzyme isocitrate lyase.

2. Growth of M. silvestris M. silvestris was grown as previously described (Theisen et al., 2005), except that the growth medium (dilute nitrate mineral salts, DNMS) contained (in mg l
1) MgSO47H2O (108), CaCl22H2O (26), KNO3 (100), FeEDTA (3.8), ZnCl2 (0.035), MnCl24H2O (0.05), H3BO4 (0.003), CoCl26H2O (0.05), CuCl22H2O (0.001), NiCl26H2O (0.012), Na2MoO42H2O (0.05), FeCl24H2O (0.75), and phosphate buffer pH 5.5 (2 mM) in addition to carbon source. For growth on agar plates, nitrate was replaced with ammonium chloride (1 mM) (DAMS medium). Nitrogen fixation is possible under reduced aeration (Dunfield et al., 2003), in which case fixed nitrogen was omitted from medium. Carbon sources (except where indicated) were supplied at 5 mM (succinate) or 0.1% (v/v) (methanol) or, on agar plates, by incubation in sealed containers in a methanol-rich atmosphere. Growth is comparatively slow in both

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liquid and solid media. Four weeks may be necessary for colony formation on plates, and 2–4 weeks required after a colony is transferred into liquid. Liquid cultures are subcultured, when the cell density (OD540) reaches approximately 0.4, at 1/10 or 1/20. The sensitivity of M. silvestris to several commonly used antibiotics was determined in liquid culture with methanol (0.1% v/v) as growth substrate. The minimum inhibitory concentrations (MIC; mg ml
1) are kanamycin, <0.25; chloramphenicol, > 50; streptomycin, 2; gentamicin, 1; spectinomycin, 5; tetracycline, 0.5; neomycin, <2; aprimycin, <2; nalidixic acid, >50. We have found that kanamycin and gentamicin are suitable antibiotics for use with this organism. Tetracycline is satisfactory for transient expression of, for example, the Cre recombinase as described later, but is not sufficiently stable for long incubation periods. It should also be noted that due to the slime producing ability of M. silvestris, growth on plates continues at considerably higher antibiotic concentrations. These antibiotics are used at 20 mg ml
1 (kanamycin), 5 mg ml
1 (gentamicin), and 10 mg ml
1 (tetracycline) in both liquid and solid media.

3. Introduction of Plasmid DNA by Conjugation and Electroporation It was previously shown that the IncP broad-host-range promoter probe vector pMHA203 can be introduced into M. silvestris by conjugation (Ali and Murrell, 2009; Theisen et al., 2005). This RK2-derived vector incorporates the s54 promoter of the sMMO fused to a gene encoding green fluorescent protein (GFP). Analysis of cells containing this plasmid during growth on various substrates demonstrated that only during growth on methane was GFP expressed. As proof of principle, this vector was reintroduced into M. silvestris by both conjugation and electroporation. Roughly equivalent numbers of transformants per microgram of DNA were obtained by both methods, demonstrating the potential of electroporation as a method of DNA delivery.

4. Gene Deletion by Electroporation of Linear DNA Electroporation proved to offer benefits over conjugation in terms of reducing the danger of contamination in the slow-growing and facultative organism M. silvestris. As mentioned, electroporation can be used to introduce circular double stranded plasmid DNA. However, for targeted mutagenesis, this method of

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DNA introduction also permitted the use of linear DNA to force gene deletion by two simultaneous recombination events, as described above, thus further minimizing transfers and screening. While not effective in many organisms, this method proved relatively efficient in M. silvestris.

5. Preparation of Competent Cells for Electroporation of M. silvestris Electroporation is feasible as a vehicle for the introduction of exogenous DNA into M. silvestris using cells grown in batch culture in both flasks and fermenter vessels. Cells were grown in a 4 l fermenter in fed-batch mode using an appropriate carbon source, for example methanol (0.1% v/v), under nitrogen-fixing conditions (i.e. with nitrogen-free medium under oxygen limitation). Cells (400 ml) at an OD540 of 2.0 were cooled on ice and harvested by centrifugation (6000g, 15 min, 4  C), washed twice in ice-cold sterile water, and resuspended in 10 ml 10% (v/v) cold glycerol. Cells can either be used immediately for electroporation, or snap frozen in liquid nitrogen and stored for several weeks at
80  C in aliquots.

6. Construction of Linear DNA for Gene Deletion by Homologous Recombination There are few restrictions on the choice of vector for construction of the linear DNA fragment. Allelic exchange vector pCM184 (Marx and Lidstrom, 2002) is convenient to use for the assembly of this construct, although this vector contains features (such as oriT) which are not used in this approach. This vector includes multiple cloning sites either side of a kanamycin-resistance cassette, itself flanked by loxP recombination sites, as shown in Fig. 8.1. Approximately 500 bp of chromosomal DNA were amplified by PCR, from regions upstream and downstream of the desired gene deletion, using M. silvestris genomic DNA as template and primers as shown in Table 8.1. In most cases, primers were designed to incorporate restriction sites. These fragments were cloned separately into pCR2.1 Topo (Invitrogen) following the manufacturer’s instructions. The upstream and downstream fragments were excised from pCR2.1 by digestion and ligated separately into pCM184 cut with the same enzymes. Digestion of the resultant molecule with the enzymes at the extremities of the homologous regions resulted in a linear DNA fragment that was purified from an agarose gel.

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A

EcoRI

B

Msil3157

(A)

Mlul

Kpnl loxP

kan

Sacl

loxP

bla oriT

pCM184

ColE1 ori tet

EcoRI

3157Tf

(B)

Sacl

kan

Msil3157 3157Tr

(C)

kan Cre

(D)

loxP

Figure 8.1 The gene deletion method is illustrated using the deletion of Msil3157 (isocitrate lyase) as an example. Primers 3157Af/3157Ar and 3157Bf/3157Br were used to amplify sequences A and B (depicted as open rectangles) located upstream and downstream of the coding sequence, as shown at (A). These fragments were then cloned into the EcoRI/KpnI and MluI/SacI sites of pCM184. The EcoRI/SacI fragment containing the kanamycin cassette was excised and purified, before electroporation into wild-type M. silvestris cells, as shown at (B). Acquisition of kanamycin resistance required double homologous recombination, in which the targeted gene was replaced by the kanamycin cassette (C). Following electroporation of this strain with pCM157, and expression of Cre recombinase, site-specific recombination between loxP sites resulted in the loss of the kanamycin cassette and the production of an unmarked mutant (D). Primers 3157Tf/3157Tr (B–D), located outside the manipulated region, were used to monitor gene replacement and deletion.

Table 8.1 Primer sequences used in this work Name

Description

Sequence 50 –30

3157Af 3157Ar 3157Bf 3157Br 3157Tf 3157Tr 3157Cf 3157Cr 3157f 3157r 1641Af 1641Ar 1641Bf 1641Br 1641Tf 1641Tr Kanf Kanr

Upstream homology fragment Upstream homology fragment Downstream homology fragment Downstream homology fragment Detection of gene deletion/substitution Detection of gene deletion/substitution Complementation Complementation Coding region Coding region Upstream homology fragment Upstream homology fragment Downstream homology fragment Downstream homology fragment Detection of gene substitution/deletion Detection of gene substitution/deletion Kanamycin cassette forward Kanamycin cassette reverse

TCACTGTGCGGCGACTATG TATCGGTACCCGTTGAGGACCGCCTCAAG TATCACGCGTTGCGTCTGCCTTGTTCAGTC TATCGAGCTCCCAGCGCCAGCTGTTCTTC AAGTCTCGGCTTCATGCTAGCG CGTCGATCTCGTCCGACATTTC ATCAGGTACCGAGGCTCCGCGCTGTTTC ATCACCCGGGATCTGCCGGCGTTCTTTG GATCATGCGCAAAGACATGG TTTCTTGGCGAGGAGATACG ATCAGAGCTCAAAGCACGGCCGCTATCG ATCAACGCGTGCGCTTTCGCCCTGATAACC ATCAGGTACCCGTCATTGGGCAACGATAAG GAAACCGCCAATGCATCTC GCCGATTGGAGCTAAACTTC GGCGAGATTCTTCTTCGTTC GCGATAATGTCGGGCAATCAG AAACTCACCGAGGCAGTTCC

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7. Electroporation For electroporation, 100 ml competent cell suspension was gently mixed with 100–500 ng linearized DNA and transferred to a 1 mm cooled electroporation cuvette (BTX Harvard Apparatus, Holliston, USA). Electroporation settings were 2.2 kV, 400 O, 25 mF, using a Bio-Rad Gene Pulser (Bio-Rad, Hercules, USA), resulting in a time constant of approximately 8 ms. Cells were immediately washed from the cuvette with 500 ml DAMS medium (containing a suitable carbon source), and allowed to recover overnight at 25  C with shaking at 150 rpm. Cells were pelleted by centrifugation (1500g, 8 min, room temperature), washed twice in 1 ml, and resuspended in 500 ml DAMS medium, before plating on DAMS agar plates containing kanamycin. After 3–4 weeks, a single colony was picked into 10 ml DNMS medium with kanamycin and subcultured into 20 ml of the same medium. The turbid culture was then serially diluted (10
4 to 10
7) and 100 ml plated on DAMS/kanamycin plates. A single colony was transferred to liquid medium without selection. A 50 ml culture was used to generate competent cells as described above, which were electroporated with plasmid pCM157 (300 ng) (Marx and Lidstrom, 2002) for expression of the Cre recombinase. Following recovery, cells were plated on DAMS/tetracycline plates. A single colony was picked into liquid culture without selection. In the absence of selection, the plasmid was lost after a few transfers, as determined by sensitivity to tetracycline and PCR using primers internal to the tetracycline gene. Substitution of the gene of interest with the kanamycin cassette, and subsequent removal of the marker following Cre recombination, was confirmed by PCR using a forward primer upstream and reverse primer downstream of the cloned homology regions used for recombination, followed by sequencing.

8. Efficiency of Gene Deletion The efficiency of gene deletion was determined using a construct designed for the deletion of open reading frame Msil1641, annotated as encoding gluconate dehydrogenase. This gene product is unnecessary for growth on methanol, which allows the use of this substrate for growth. Deletion is unlikely to have any polar effect, since the predicted downstream open reading frame is transcribed in the reverse orientation. The procedure as described above was employed, using primers 1641Af/1641Ar and 1641Bf/1641Br, using methanol (0.1% v/v) as growth substrate. As shown in Fig. 8.2, gene replacement events were approximately proportional to DNA mass added to the electroporation cuvette. All colonies

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Gene replacements (ng DNA)-1

5 4 3 2 1 0

0

25

50 DNA mass (ng)

75

100

Figure 8.2 Frequency of replacement of Msil1641 with the kanamycin cassette as a function of mass of linear DNA incorporated in the electroporation reaction.

tested (20/20) contained the desired gene deletion, as determined by diagnostic PCR using primers 1641Tf/1641Tr.

9. Case Study: Deletion of Isocitrate Lyase To determine the requirement for isocitrate lyase during growth on both one- and two-carbon compounds, the putative open reading frame encoding this enzyme, Msil3157, was deleted from the M. silvestris chromosome. This gene encodes a monocistronic mRNA molecule, and it is flanked by predicted open reading frames transcribed in the opposite direction. A predicted s70 promoter is located adjacent to a transcription start site 88 bp upstream of the putative start codon. The method of gene deletion (Fig. 8.1) was as described above, using primers 3157Af/3157Ar and 3157Bf/3157Br. Since this deletion may affect growth on one- and two-carbon compounds, succinate (5 mM) was used as growth substrate. Genetic manipulations were monitored by PCR using primers 3157Tf/3157Tr and Kanf/Kanr (see Fig. 8.3).

10. Complementation The isocitrate lyase deletion strain of M. silvestris was complemented with the wild-type gene and promoter expressed from a broad-host-range plasmid. Vector pCM132 (Marx and Lidstrom, 2001) was digested with SphI, blunted using T4 polymerase, and digested with KpnI. The M. silvestris

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A

B

kb

kb

4 3 2

1 0.75 0.5

1

2

3

4

1

2

3

4

5

Figure 8.3 Agarose gels confirming the replacement of Msil3157 by kan and its subsequent removal. (A) PCR using primers 3157Tf/3157Tr. Lane1: wild-type; lane 2: D3157::kan double-crossover; lane 3: DIcl mutant following Cre-mediated removal of the kanamycin cassette; lane 4: no template control (NTC). (B) KanF/KanR primers. Lane 1: wild-type; lane 2: D3157::kan double-crossover; lane 3: DIcl mutant; lane 4: positive control (pCM184 vector DNA); lane 5: NTC.

isocitrate lyase coding sequence (Msil3157) and 182 bp upstream and 185 bp downstream were amplified by PCR using primers 3157Cf/3157Cr, cloned into pCR2.1 Topo, cut with KpnI and SmaI, and ligated with the major fragment of the digested vector, resulting in pAC105, as shown in Fig. 8.4A. This vector was introduced into the isocitrate lyase deletion M. silvestris strain by electroporation. The genotype of the complemented mutant was verified by PCR. Transcription of isocitrate lyase was investigated by RTPCR using cDNA generated from mRNA extracted from wild-type, mutant, and complemented mutant cells (using standard methods; Sambrook et al., 2001). These data confirmed the presence of the isocitrate lyase gene located on the vector in the complemented mutant, and that in this strain, as in the wild type, it was transcribed during growth on both succinate and methanol, as shown in Fig. 8.4B.

11. Phenotype of the Isocitrate Lyase Deletion Mutant Deletion of isocitrate lyase from M. silvestris had little effect on growth on succinate, but abolished growth on methanol and severely restricted growth on acetate (Fig. 8.5), confirming the essential role of this gene

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oriV

oriT

A

B

traJ⬘ ColE1 ori

trfA

pAC105 kan

Msil3157

3157f

3157r trrnB

1 2 3 4 5 6 7 8 9

Figure 8.4 (A) Map of complementation plasmid pAC105. (B) RNA extracted from cells was used to synthesize cDNA which was used in RT-PCR reactions. Lane 1: wildtype, succinate-grown; lane 2: mutant, succinate-grown; lane 3: complemented mutant, succinate-grown; lane 4: complemented mutant, methanol-grown; lane 5: genomic DNA template; lanes 6–9: negative controls.

product in regeneration of glyoxylate as part of the serine cycle during onecarbon growth, and the importance of the glyoxylate bypass during growth on two-carbon compounds. Complementation largely restored wild-type growth on both substrates.

12. Conclusions The development of an effective method of genetic manipulation was an essential tool for unraveling the metabolic pathways employed by M. silvestris during growth on different substrates. Using the methods described here, we have constructed several mutant strains in our laboratory. This procedure has significant advantages over the conjugation methods which we have previously employed with other methanotrophs (e.g., Ali et al., 2006), due to the growth characteristics of M. silvestris. The methods are now in place to dissect the metabolic pathways of this organism, and explore the regulatory mechanisms operational during growth on different substrates. Additionally, the potential now exists to engineer strains with desirable properties for biotransformation and bioremediation, for example, a strain expressing the sMMO constitutively and thus potentially able to degrade environmental pollutants while growing on an alternative carbon source, for example, acetate or ethanol.

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B

0.7

0.7

0.6

0.6

0.5

0.5

0.4

0.4

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A

0.3

0.3

0.2

0.2

0.1

0.1

0

0

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0

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0

50

Time (h) C

100

150

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Time (h)

0.5

OD540

0.4 0.3 0.2 0.1 0

0

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Figure 8.5 Growth curves of M. silvestris wild type (solid lines), isocitrate lyase mutant (dotted lines), and complemented mutant (dashed lines) during growth on (A) succinate, (B) methanol, and (C) acetate. Error bars show the standard deviation of three replicates.

REFERENCES Ali, H., and Murrell, J. C. (2009). Development and validation of promoter-probe vectors for the study of methane monooxygenase gene expression in Methylococcus capsulatus Bath. Microbiology 155, 761–771. Ali, H., et al. (2006). Duplication of the mmoX gene in Methylosinus sporium: Cloning, sequencing and mutational analysis. Microbiology 152, 2931–2942. Ayres, E. K., et al. (1993). Precise deletions in large bacterial genomes by vector-mediated excision (VEX): The trfA gene of promiscuous plasmid RK2 is essential for replication in several Gram-negative hosts. J. Mol. Biol. 230, 174–185. Baani, M., and Liesack, W. (2008). Two isozymes of particulate methane monooxygenase with different methane oxidation kinetics are found in Methylocystis sp. strain SC2. Proc. Natl. Acad. Sci. USA 105, 10203–10208.

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Becking, J. (2006). The genus Beijerinckia. In “The Prokaryotes,” (M. Dworkin, et al., eds.) Vol. 5, pp. 151–162. Springer, New York. Belova, S. E., et al. (2010). Acetate utilization as a survival strategy of peat-inhabiting Methylocystis spp. Environ. Microbiol. Rep. 10.1111/j.1758-2229.2010.00180.x (in press). Chen, Y., et al. (2010). Complete genome sequence of the aerobic facultative methanotroph Methylocella silvestris BL2. J. Bacteriol. 192, 3840–3841. Colby, J., et al. (1977). The soluble methane mono-oxygenase of Methylococcus capsulatus (Bath). Its ability to oxygenate n-alkanes, n-alkenes, ethers, and alicyclic, aromatic and heterocyclic compounds. Biochem. J. 165, 395–402. Davison, J. (2005). Risk mitigation of genetically modified bacteria and plants designed for bioremediation. J. Ind. Microbiol. Biotechnol. 32, 639–650. Dedysh, S. N., et al. (2005). Methylocella species are facultatively methanotrophic. J. Bacteriol. 187, 4665–4670. Driskell, L. O., et al. (2009). Directed mutagenesis of the Rickettsia prowazekii pld gene encoding phospholipase D. Infect. Immun. 77, 3244–3248. Dunfield, P. F., Belova, S. E., Vorobe´v, A. V., Cornish, S. L., and Dedysh, S. N. (2010). Methylocapsa aurea sp. nov., a facultative methanotroph possessing a particulate methane monooxygenase, and emended description of the genus Methylocapsa. Int. J. Syst. Evol. Microbiol. 60, 2659–2664. Dunfield, P. F., et al. (2003). Methylocella silvestris sp. nov., a novel methanotroph isolated from an acidic forest cambisol. Int. J. Syst. Evol. Microbiol. 53, 1231–1239. El Karoui, M., et al. (1999). Gene replacement with linear DNA in electroporated wild-type Escherichia coli. Nucleic Acids Res. 27, 1296–1299. Erb, T. J., et al. (2009). (2S)-Methylsuccinyl-CoA dehydrogenase closes the ethylmalonylCoA pathway for acetyl-CoA assimilation. Mol. Microbiol. 73, 992–1008. Hansen, E. J., et al. (1992). Use of electroporation to construct isogenic mutants of Haemophilus ducreyi. J. Bacteriol. 174, 5442–5449. Hoang, T. T., et al. (1998). A broad-host-range Flp-FRT recombination system for sitespecific excision of chromosomally-located DNA sequences: Application for isolation of unmarked Pseudomonas aeruginosa mutants. Gene 212, 77–86. Im, J., et al. (2010). Characterization of a novel facultative Methylocystis species capable of growth on methane, acetate and ethanol. Environ. Microbiol. Rep. 10.1111/j.17582229.2010.00204.x (in press). Jasin, M., and Schimmel, P. (1984). Deletion of an essential gene in Escherichia coli by site-specific recombination with linear DNA fragments. J. Bacteriol. 159, 783–786. Kim, C., and Wood, T. K. (1998). Electroporation of pink-pigmented methylotrophic bacteria. Appl. Biochem. Biotechnol. 73, 81–88. Marx, C. J., and Lidstrom, M. E. (2001). Development of improved versatile broad-hostrange vectors for use in methylotrophs and other Gram-negative bacteria. Microbiology 147, 2065–2075. Marx, C. J., and Lidstrom, M. E. (2002). Broad-host-range cre-lox system for antibiotic marker recycling in gram-negative bacteria. Biotechniques 33, 1062–1067. Murphy, K. C. (1998). Use of bacteriophage lambda recombination functions to promote gene replacement in Escherichia coli. J. Bacteriol. 180, 2063–2071. Murrell, J. C. (1992). Genetics and molecular biology of methanotrophs. FEMS Microbiol. Lett. 88, 233–248. Oh, S. H., and Chater, K. F. (1997). Denaturation of circular or linear DNA facilitates targeted integrative transformation of Streptomyces coelicolor A3(2): Possible relevance to other organisms. J. Bacteriol. 179, 122–127. Rothstein, R. J. (1983). One-step gene disruption in yeast. Methods Enzymol. 101, 202–211. Sambrook, J., et al. (2001). Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, New York.

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Scha¨fer, A., et al. (1994). Small mobilizable multi-purpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19: Selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145, 69–73. Theisen, A. R., and Murrell, J. C. (2005). Facultative methanotrophs revisited. J. Bacteriol. 187, 4303–4305. Theisen, A. R., et al. (2005). Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol. Microbiol. 58, 682–692. Toyama, H., et al. (1998). Construction of insertion and deletion mxa mutants of Methylobacterium extorquens AM1 by electroporation. FEMS Microbiol. Lett. 166, 1–7. Wood, A. P., et al. (2004). A challenge for 21st century molecular biology and biochemistry: What are the causes of obligate autotrophy and methanotrophy? FEMS Microbiol. Rev. 28, 335–352. Zealey, G. R., et al. (1990). Gene replacement in Bordetella pertussis by transformation with linear DNA. Biotechnology 8, 1025–1029.

C H A P T E R

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Mutagenesis of Soluble Methane Monooxygenase Thomas J. Smith* and J. Colin Murrell† Contents 136 137 139 139 140 141 141 144 144 145

1. Introduction 2. Bacteriological Growth Media and Antibiotics 3. Mutagenesis and Subcloning 3.1. Design of mutants 3.2. Mutagenesis 3.3. Subcloning of mutants 4. Expression Hosts and Conjugation 5. Analysis of Mutants 5.1. Confirmation of the mutant genotype 5.2. Initial characterization of mutant enzymes 5.3. Large-scale production of cultures expressing mutant enzymes Acknowledgments References

145 146 146

Abstract The hydroxylase component of soluble methane monooxygenase (sMMO), which is the site of oxidation of methane and many adventitious substrates of commercial and environmental interest, has proved challenging to manipulate genetically because of difficulties with obtaining functional expression in Escherichia coli. Here, we describe methods that allow site-directed mutagenesis of the hydroxylase-encoding genes and subsequent production of mutant proteins in a modified strain of a methane-oxidizing bacterium, using methane as the carbon and energy source. Mutagenesis and other genetic manipulations are performed in E. coli via standard methods and then, a shuttle plasmid is used to transfer the mutant genes via conjugation to a strain of Methylosinus trichosporium in which the chromosomal copy of the sMMO operon has been partially deleted. Expression is directed by the natural sMMO promoter at high cell density under appropriate culture conditions. The system is not restricted * Biomedical Research Centre, Sheffield Hallam University, Sheffield, United Kingdom School of Life Sciences, University of Warwick, Coventry, United Kingdom

{

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00009-7

#

2011 Elsevier Inc. All rights reserved.

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to active mutants of sMMO because Ms. trichosporium can grow on methane using the membrane-associated particulate methane monooxygenase (pMMO) even when it has no active sMMO.

1. Introduction Soluble methane monooxygenase (sMMO) is a three-component nonheme iron oxygenase that is one of the two enzyme systems that methane-oxidizing bacteria possess to catalyze the chemically demanding initial step of the methane oxidation pathway, conversion of methane to methanol. sMMO has many potential applications in synthetic organic chemistry and bioremediation because it can cooxidize a very wide range of adventitious substrates, including alkanes, alkenes, alcohols, ethers, alicyclics, aromatics (Colby et al., 1977; Stirling and Dalton, 1979), and chlorinated organic compounds such as the pollutant trichloroethene (Fox et al., 1990). Most well-characterized organisms that contain sMMO can produce the alternative methane-oxidizing enzyme particulate methane monooxygenase (pMMO) under conditions of sufficient copper availability (Hakemian and Rosenzweig, 2007; Hanson and Hanson, 1996; Semrau et al., 2010; Smith and Murrell, 2009; Stanley et al., 1983; Trotsenko and Murrell, 2008). However, facultative methanotrophs belonging to the genus Methylocella that have been characterized to date possess sMMO but not pMMO (Theisen et al., 2005). The expression system that we have developed for sMMO is based upon Methylosinus trichosporium OB3b, a methanotroph that can produce either sMMO or pMMO, dependent upon the copper-to-biomass ratio of the culture. Problems that have prevented expression of the terminal oxygenase (hydroxylase) component of the enzyme in Escherichia coli (West et al., 1992) are overcome by using a homologous expression system, that is, by expressing the recombinant enzymes in Ms. trichosporium. sMMO comprises a multisubunit hydroxylase that is the site of methane oxidation, an NAD(P)H-dependent reductase with FAD and Fe2S2 prosthetic groups, and a third component that is also necessary for full activity, termed protein B. Hence, genetic manipulation of sMMO involves dealing with a multigene operon, mmoXYBZorfYmmoC (Cardy et al., 1991), which encodes the full enzyme complex. The diiron active site at which substrate oxygenation occurs resides in the a-subunit (MmoX) of the hydroxylase component, which has an a2b2g2 structure. The b and g subunits are encoded by mmoY and mmoZ, respectively. The reductase component, which provides electrons from NADH (or NADPH), is encoded by mmoC. Many detailed studies have indicated multiple and complex roles

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for protein B (MmoB), one of which is to favor coupling of NAD(P)H oxidation by the reductase and oxygenation of the substrate at the hydroxylase (Fox et al., 1991; Lipscomb, 1994; Murrell et al., 2000). OrfY (also known as MmoD) is not required for activity of the final complex but may have a role in complex assembly (Merkx and Lippard, 2002). The reductase and protein B components are active when expressed individually in E. coli using standard expression vectors and so, mutagenesis of these components is relatively straightforward. Details of the mutagenesis and expression systems for these components from the two model sMMOexpressing methanotrophs, Ms. trichosporium OB3b and Methylococcus capsulatus (Bath), can be found in the literature (Blazyk and Lippard, 2004; Brandstetter et al., 1999; Brazeau et al., 2001; West et al., 1992). The system described here for manipulation of the hydroxylase component comprises a broad host-range vector that allows construction of mutants in E. coli and sMMO-minus derivatives of Ms. trichosporium into which mutant sMMO-encoding genes can be transferred by means of conjugation, for subsequent expression, analysis, and protein purification. Ms. trichosporium grows on methane or methanol. The chromosomal copies of the pMMO remain intact, and so the sMMO-minus host strains or strains expressing inactive mutants of sMMO can be grown on methane via the pMMO. The natural promoter system is used to activate (mutant) sMMO expression in response to low copper-to-biomass ratio in the culture. Although this regulation system may sound cumbersome, in practice, it works very well as long as appropriate growth conditions are chosen, because cultures begin growing on the pMMO at low density and spontaneously switch to expressing the recombinant sMMO as the density of the culture increases. sMMO naturally constitutes around 10% of cellular protein in sMMO-expressing Ms. trichosporium cells, and so, the yield of protein from this system is very workable. Basic methods for subcloning and generation of mutants in E. coli are widely used and are described in outline only. Methods specific to sMMO and methanotrophs, which we have developed in our laboratories, are described in detail.

2. Bacteriological Growth Media and Antibiotics Ms. trichosporium strains are grown in NMS medium based upon that described in the literature (Dalton and Whittenbury, 1976) containing appropriate antibiotics. NMS medium is made from five stock solutions:

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10
NMS salts KNO3 MgSO47H2O CaCl22H2O Stored at 4  C

10 g L 1 10 g L 1 2 g L 1

Na molybdate solution NaMoO42H2O Stored at 4  C

0.5 g L 1

FeEDTA solution FeEDTA Stored in the dark at 4  C

38 g L 1

NMS trace elements CuSO45H2O FeSO47H2O ZnSO47H2O Orthoboric acid CoCl36H2O Na2EDTA MnCl24H2O NiCl26H2O Stored at in the dark at 4  C

100 mg L 1 500 mg L 1 400 mg L 1 15 mg L 1 50 mg L 1 250 mg L 1 20 mg L 1 10 mg L 1

NMS phosphate buffer solution Na2HPO412H2O (or Na2HPO4) KH2PO4

107.4 g L 1 49.7 g L 1 39 g L 1

Liquid NMS medium is prepared by mixing: 10
NMS salts Na molybdate solution NMS trace elements Fe EDTA solution Water

100 mL 1 mL 1 mL 0.1 mL to 1 L

pH should be 6.8 without adjustment. Sterilized by autoclaving in 100-mL aliquots and stored at room temperature.

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This mixture is sterilized by autoclaving and may be stored at room temperature. For NMS agar plates, Difco or Oxoid bacteriological agar is added to 15 g L 1 before autoclaving. The liquid medium may develop a small amount of gray/green precipitate, which does not interfere with its use. Immediately before use, when the medium is fully cooled (or, in the case of agar medium, can be comfortably held in the bare hands), 10 mL of sterile NMS phosphate buffer solution is added per liter of medium. All methods described in this chapter utilize NMS with the copper content described above, that is, 0.1 mg of copper sulfate pentahydrate per liter of final medium. E. coli is propagated in LB medium (tryptone, 10 g L 1; yeast extract, 5 g L 1; NaCl, 5 g L 1) or on plates of LB medium containing agar (1.5% w/v). All plasmids that we have constructed for use during expression and mutagenesis of sMMO possess beta-lactamase genes and are selected in E. coli using ampicillin (100 mg mL 1 routinely or, when selecting transformants immediately after ligation, 50 mg mL 1). Antibiotics used for selection of plasmid- and chromosome-encoded resistance markers in strains of Ms. trichosporium are kanamycin (10–50 mg mL 1), gentamicin (5 mg mL 1), streptomycin (20 mg mL 1), and spectinomycin (20 mg mL 1). Nalidixic acid (20 mg mL 1), to which Ms. trichosporium is intrinsically resistant, is used to eliminate the E. coli donor strain after conjugation.

3. Mutagenesis and Subcloning 3.1. Design of mutants In designing mutations, we have wherever possible avoided introducing codons that are rare in Ms. trichosporium. The codon usage table based on the sMMO operon of Ms. trichosporium (Murrell, 1993) has served well in this regard. Since Ms. trichosporium is a GC-rich organism, selection of alternatives with C or G at the third position is generally sufficient. Construction of mutants requires a number of genetic manipulations in E. coli and Ms. trichosporium, followed in many cases by large-scale cultivation of the mutant-expressing strain, which may run over a number of weeks or months. It is advantageous to be able easily to tell whether the strain has the desired mutant, which can be done by PCR amplification of a region spanning the mutation, followed by sequencing. With some mutations, we have designed the mutagenic oligonucleotides to introduce changes in restriction pattern that can be used to identify mutants by restriction digestion after PCR. In some instances, the change that introduces the desired change in encoded amino acid is sufficient; in other cases, a nearby

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Table 9.1 Examples of oligonucleotides used to mutate mmoX Oligonucleotide (50 –30 )a

Mutation/primer

Mutagenic primers C151E

Novel StuI site C ACG CAT CAA GAG GCC TTC ATC AAT C Glu 151 Novel RsaI site

C151Y

C ACG CAT CAG TAC GCC TTC ATC AAT C Tyr 151 T213A

PvuII site removed GCT CGT CGG CGA AGC CTG CTT CGC GAA TCC GCT CAT C Ala 213 PvuII site removed CGG ATT CGC GAA GCA GGC TTC GCC GAC GAG CTG CAG A

External primers Upper

Ala 213

CAG GAA ACA GCT ATG AC

Lower

NdeI CC GTT CGC CAT ATG ACG CAG CTC GTC

a

Mutations are shown in bold type. Codons encoding altered amino acids are underlined; changes to the restriction pattern associated with each mutation are shown by a line above the sequence. The pairs of mutagenic primers used to create each mutation at amino acid 151 were complementary and so, only the forward primer is shown. The pairs of mutagenic primers used to make changes at amino acid 213 spanned slightly different regions of the gene; both are shown for each mutation, the forward primer first (Smith et al., 2002).

change has been made that is silent in terms of the encoded protein. Examples are given in Table 9.1.

3.2. Mutagenesis Mutants of the sMMO-encoding genes are constructed in a general cloning strain of E. coli (such as DH5a or INVaF0 [Invitrogen]) by means of the four-primer overlap extension PCR method described by Ho et al. (1989). Use of a proof-reading polymerase for PCR (we regularly use Pfu TURBO from Stratagene) prevents the unacceptable frequency of PCR-derived mutations that we have observed with Taq polymerase. The template for the mutagenic PCRs is pNPB101, a subclone of part of the 50 portion of mmoX (1.0 kb, including the start of the gene and coding sequence as far as a natural NdeI site, i.e., allowing mutagenesis within the region from the N-terminus up to and including Arg 245 out of 526 amino acyl residues). The external primers used in the mutagenic and combinatorial PCRs are given in Table 9.1.

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3.3. Subcloning of mutants The mutagenized 1.0-kb gene fragment is cloned back into the same vector (pUC19) with BamHI and NdeI. After sequencing the cloned insert and confirming the absence of unwanted mutations, the mutant insert is subcloned with BamHI and NdeI into the rest of the sMMO operon in pTJS176 (Fig. 9.1). The 10-kb insert of the mutant pTJS176, containing the mutagenized sMMO operon, is then transferred to the E. coli–Ms. trichosporium shuttle vector pTJS140 using KpnI. pTJS140, which has an RK2 replicon and is based upon pJB3Km1 (Blatny et al., 1997), gives low transformation frequencies compared to most commercial vectors and so, we routinely dephosphorylate the vector and use commercially prepared supercompetent cells (Stratagene Solopak Gold) when cloning into pTJS140. pTJS140 permits blue–white selection of recombinant clones based on the inactivation of lacZ0 by the insert, but does not, in this application, allow directed cloning. A restriction digestion of the progeny (BamHI is convenient) is needed to select a clone with the insert in the correct orientation, with the sMMO operon and the lac promoter running in the same direction. The result of this cloning is a mutant of pTJS175, which contains a (modified) 10-kb fragment of the Ms. trichosporium chromosome that includes mmoR and mmoG (encoding putative sN-dependent transcriptional activator and a GroEL homolog, both of which are needed for production of functional sMMO; Stafford et al., 2003) as well as the mutagenized sMMO structural genes (Fig. 9.1). Currently, we are manipulating the restriction sites of the expression plasmid pTJS175 to enable cloning of the 1.0-kb BamHI–NdeI fragments, carrying the mutated region of mmoX, directly into the expression plasmid as a single cloning step in E. coli. The plasmid containing the mutant or recombinant sMMO operon is transformed by means of electroporation into the conjugation donor strain E. coli S17-1, and transformants are selected on the basis of resistance to ampicillin (50 mg mL 1). A small-scale DNA preparation is made and analyzed to confirm the presence of the desired plasmid.

4. Expression Hosts and Conjugation We have constructed two sMMO-minus derivatives of Ms. trichosporium OB3b by means of marker exchange mutagenesis in order to allow expression of mutants of the hydroxylase component of sMMO. In Ms. trichosporium mutant F, a 1.2-kb fragment internal to the chromosomal copy of mmoX is replaced by a kanamycin-resistance cassette (Martin and Murrell, 1995). Since the deleted portion of mmoX spans the codons for amino acids Val 112-Thr 508 within the MmoX protein, we have used this strain only for expression of

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mmoX⬘ pNPB101 (vector pUC18), for mutagenesis of mmoX Mutagenesis via four-primer PCR method and cloning into same vector with BamHI and NdeI

* mmoX⬘ pNPB101 (vector pUC18), with desired mutation in mmoX Cloning of mutant mmoX’ fragment into rest of sMMO operon in pTJS176, using BamHI and NdeI

* mmoR

G

mmoX

Y

B

Z

orfY mmoC

Mutant version of pTJS176 (vector pMTL24), without additional NdeI and BamHI sites, that allowed inward cloning of BamHI–NdeI mmoX fragments Cloning of whole sMMO operon into shuttle vector pTJS140 using KpnI. Orientation of insert confirmed via restriction analysis with BamHI

* mmoR Sm/SpR

G

mmoX

Y

B

Z

orfY mmoC

Direction of interrupted lacZ⬘ oriT

ApR

ori-RK2

Mutant pTJS175 (vector pTJS140) for transfer by conjugation and expression of sMMOin Ms. trichosporium mutant F or SMDM

Figure 9.1 Cloning strategy for generation of mutant sMMO operons. The asterisk represents the mutation in mmoX. Additional details are given in the text.

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recombinant wild-type sMMO and derivatives where the mutation lies within the deleted region. Use of mutant F for expression of mutants outside this region is not recommended, because it is possible that the mutation will be repaired by recombination with the chromosome of the host. The more recently constructed sMMO-minus strain Ms. trichosporium SMDM (for soluble MMO-deleted mutant) is deleted in the whole of the sMMO-encoding operon, except for the 30 portion of mmoC, and has a gentamicin-resistance cassette in place of the deleted genes. We have used Ms. trichosporium SMDM successfully for expression of variants with mutations throughout MmoX (Borodina et al., 2007). In principle, it should be suitable for expression of mutants anywhere in the sMMO complex, except for the reductase (MmoC). Ms. trichosporium mutant F and strain SMDM are propagated on NMS medium containing the relevant antibiotic to select for the chromosomal inactivation (kanamycin and gentamicin, respectively). The strains have proved stable in the absence of antibiotic selection, and addition of an antibiotic to select for the gene inactivations should be avoided with cultures to be used for expressing sMMO to minimize antibiotic stress on the cells. Conjugation of the mutant pTJS175 plasmid into Ms. trichosporium mutant F or SMDM is effected as follows: Several colonies of the recipient strain are inoculated into 50 mL of NMS medium (containing kanamycin if Ms. trichosporium mutant F is used or gentamicin for strain SMDM) in a 250-mL Quickfit conical flask and sealed with a rubber Subaseal (Fisher Scientific). Fifty milliliters of headspace gas is removed using a 50-mL disposable hypodermic syringe and needle and replaced with 60 mL of methane. The culture is incubated at 30  C with shaking (180 rpm) until the OD540 is approximately 0.2 (which generally takes between 24 h and several days). The methanotroph cells are harvested by centrifugation (7000
g, 5 min, room temperature), washed with 50 mL of fresh NMS medium, and resuspended in 10 mL of the same medium. The donor strain is grown overnight from a single colony in LB medium (plus 100 mg mL 1 of ampicillin to select for the plasmid), centrifuged as above, washed with 50 mL of fresh NMS medium, and then resuspended in 10 mL of the same medium. The suspensions of washed donor and recipient cells are mixed and the cells are collected onto a 47-mm diameter sterile filter (0.2 mm pore size), by using a sterile reusable plastic filter unit connected to a suction pump. The filter unit is then dismantled. A pair of stainless steel tweezers are sterilized by dipping in absolute ethanol and flaming to remove the ethanol and are then used to transfer the filter, bacteria-side-up onto a plate of NMS agar plus 0.02% (w/v) Proteose Peptone (BD catalog no. 212010) and incubated for 24 h at 30  C in an air/methane atmosphere (50:50 by volume, in a sealed container such as used for cultivation of anaerobes). By using a 1-mL micropipette, a pair of sterile tweezers, and a sterile plastic Petri dish, the bacteria are then washed from the filter into

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approximately 10 mL of fresh sterile NMS medium. The bacterial suspension is concentrated by centrifugation (7000
g, 5 min, room temperature) and the cells are resuspended in 1 mL of NMS medium, before plating of 200 mL aliquots on NMS plates containing 20 mg mL 1 of streptomycin to select for the transformants. A negative control plate is inoculated with the same density of recipient cells as added to the experimental plate to show the rate of spontaneous streptomycin-resistant mutants. The exconjugant and control plates are incubated at 30  C in an air/methane atmosphere until colonies appear on the experimental plates. This typically takes up to 2 weeks. Conjugation frequencies vary greatly, typically from one to 100 colonies per plate. Colonies are then picked and streaked onto fresh NMS plates containing streptomycin, spectinomycin, and nalidixic acid and incubated at 30  C in an air/methane atmosphere. Gentamicin or kanamycin may also be included to provide selection for the chromosomal rearrangement in the Ms. trichosporium hosts SMDM and mutant F, respectively. Nalidixic acid is present to kill E. coli donor cells that persist as scavengers on the methanotrophs. Use of both streptomycin and spectinomycin selects for the plasmid encoding the sMMO genes and use of both of these antibiotics minimizes spontaneous resistance mutants. Single colonies of the purified exconjugants are usually visible within 2 weeks. We have found that this two-stage selection procedure is necessary. Our attempts to shorten the selection procedure by plating the exconjugants initially on plates containing more than one antibiotic have not yielded exconjugants.

5. Analysis of Mutants 5.1. Confirmation of the mutant genotype Confirmation of the genotype of the mutant is performed by PCR amplification of a region (typically, 1–2 kb) around the mutation site and then performing restriction digestion or sequencing to confirm that the mmoX gene is present and has the desired mutation. The recipient methanotroph strain is used as a negative control in the PCR and where restriction analysis is used, the same recipient strain containing the recombinant wild-type sMMO genes is used as a comparison to observe the change in restriction pattern. It is sometimes difficult to obtain PCR products when using whole cells as the template for the PCRs; when such problems arise, we usually purify the DNA before PCR. Purification of total DNA can be done from 0.5 L of culture by the neutral lysis/CsCl method (Oakley and Murrell, 1988) or from smaller liquid cultures using the Qiagen DNEasy Blood and Tissue (50) disposable columns and reagents, according to the manufacturer’s instructions for Gram-negative bacteria.

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5.2. Initial characterization of mutant enzymes The easiest test of the activity of new mutants is the naphthalene oxidation test. This can be done as soon as the exconjugants have been recultured on NMS plates with antibiotics and gives the first indication of whether a new mutant has oxygenase activity or not. It works well either on plates that have been streaked to get single colonies, or if colonies are patched with sterile toothpicks at up to about 50 patches per agar plate. If standard NMS agar containing 0.1 mg L 1 of copper sulfate pentahydrate is used, the colonies are reliably naphthalene-positive if the sMMO is active, presumably because the high cell density of a colony produces a locally low copper-to-biomass ratio and thus induces sMMO. Use of medium with less copper than this does not give reliable results because less biomass is produced. The standard naphthalene oxidation test is performed as follows: A few crystals of naphthalene (50 mg) are scattered across the lid of the Petri dish and the plate is inverted over it, bacteria side down. The plate is sealed in a plastic sandwich box and placed in an incubator at 30  C for 1–4 h or at 45  C for 40 min. The optimum temperature for wild-type Ms. trichosporium sMMO is 30  C; the use of a higher incubator temperature is effective because the local high temperature on the base of the box helps to vaporize some of the naphthalene, while the plate remains sufficiently cool for the enzyme not to be inactivated. The plates are removed from the incubator and a freshly prepared solution of tetrazotized o-dianisidine (fast blue B salt, Sigma product no. D9805; 5 mg mL 1) is added dropwise onto the colonies. A purple or pink color that develops instantly or within a few minutes indicates sMMO activity. Positive (recombinant wild-type sMMO-expressing Ms. trichosporium) and negative (sMMO-minus Ms. trichosporium mutant F or SMDM) controls should be performed in parallel. Naphthalene is toxic to Ms. trichosporium so stock plates of new strains should not be subjected to the naphthalene oxidation tests, and sandwich boxes that have been used with naphthalene must not be used to store or cultivate plates of live bacteria.

5.3. Large-scale production of cultures expressing mutant enzymes Production of large amounts of biomass for detailed analysis of recombinant sMMO enzymes is best done in a bioreactor fitted with an additional rotameter to allow gassing with methane. Such large-scale cultivation of methanotrophs using methane as the growth substrate is described in a previous Methods Enzymol. chapter (Pilkington and Dalton, 1990). The medium used at the beginning of the growth should be NMS containing 0.1 mg L 1 of copper sulfate pentahydrate. The bioreactor can be operated batchwise or switched to continuous operation (with a dilution rate of 0.01 h 1) once the culture has switched to the production of sMMO

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(as judged by a positive naphthalene oxidation test). Antibiotics are used to prepare the starter culture for the bioreactor but are not added to the bioreactor itself to maximize production of the recombinant enzyme. We always confirm the genotype of the cells in the bioreactor by PCR amplification of a fragment surrounding the mutation site and confirmation of the presence of the mutation by restriction digestion or sequencing. Our experience with this expression system indicates that pMMO is generally still active even when sMMO is being expressed. Since the substrate range of sMMO is much wider than that of pMMO (reviewed by Smith and Dalton, 2004), many sMMO-specific whole cell screens and enzyme assays can be done directly on cells expressing both. Where it is necessary to separate sMMO from pMMO, this can easily be done by ultracentrifugation after cell breakage.

ACKNOWLEDGMENTS Both authors gratefully acknowledge funding from the Biotechnology and Biological Sciences Research Council (BBSRC).

REFERENCES Blatny, J. M., Brautaset, T., Winter-Larsen, H. C., Haugen, K., and Valla, S. (1997). Construction and use of a versatile set of broad-host-range cloning and expression vectors based on the RK2 replicon. Appl. Environ. Microbiol. 63, 370–379. Blazyk, J. L., and Lippard, S. J. (2004). Domain engineering of the reductase component of soluble methane monooxygenase from Methylococcus capsulatus (Bath). J. Biol. Chem. 279, 5630–5640. Borodina, E., Nichol, T., Dumont, M. G., Smith, T. J., and Murrell, J. C. (2007). Mutagenesis of the “leucine gate” to explore the basis of catalytic versatility in soluble methane monooxygenase. Appl. Environ. Microbiol. 73, 6460–6467. Brandstetter, H., Whittington, D. A., Lippard, S. J., and Frederick, C. A. (1999). Mutational and structural analyses of the regulatory protein B of soluble methane monooxygenase from Methylococcus capsulatus (Bath). Chem. Biol. 6, 441–449. Brazeau, B. J., Wallar, B. J., and Lipscomb, J. D. (2001). Unmasking of deuterium kinetic isotope effects on the methane monooxygenase compound Q reaction by site-directed mutagenesis of component B. J. Am. Chem. Soc. 123, 10421–10422. Cardy, D. L. N., Laidler, V., Salmond, G. P. C., and Murrell, J. C. (1991). Molecular analysis of the methane monooxygenase (MMO) gene-cluster of Methylosinus trichosporium OB3b. Mol. Microbiol. 5, 335–342. Colby, J., Stirling, D. I., and Dalton, H. (1977). The soluble methane monooxygenase of Methylococcus capsulatus (Bath): Its ability to oxygenate n-alkanes, n-alkenes, ethers, and alicyclic, aromatic and heterocyclic compounds. Biochem. J. 165, 395–402. Dalton, H., and Whittenbury, R. (1976). The acetylene reduction technique as an assay for nitrogenase activity in the methane oxidizing bacterium Methylococcus capsulatus (Bath). Arch. Microbiol. 109, 147–151. Fox, B. G., Bourneman, J., Wackett, L., and Lipscomb, J. D. (1990). Haloalkene oxidation by the soluble methane monooxygenase from Methylosinus trichosporium OB3b: Mechanistic and environmental implications. Biochemistry 29, 6419–6427.

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Fox, B. G., Liu, Y., Dege, J. E., and Lipscomb, J. D. (1991). Complex formation between the protein components of methane monooxygenase from Methylosinus trichosporium OB3b. J. Biol. Chem. 266, 540–550. Hakemian, A. S., and Rosenzweig, A. C. (2007). The biochemistry of methane oxidation. Annu. Rev. Biochem. 76, 223–241. Hanson, R. S., and Hanson, T. E. (1996). Methanotrophic bacteria. Microbiol. Rev. 60, 439–471. Ho, S. N., Hunt, H. D., Horton, R. M., Pullen, J. K., and Pease, R. L. (1989). Site-directed mutagenesis by overlap extension using the polymerase chainreaction. Gene 77, 51–59. Lipscomb, J. D. (1994). Biochemistry of the soluble methane monooxygenase. Annu. Rev. Microbiol. 48, 371–399. Martin, H., and Murrell, J. C. (1995). Methane monooxygenase mutants of Methylosinus trichosporium constructed by marker-exchange mutagenesis. FEMS Microbiol. Lett. 127, 243–248. Merkx, M., and Lippard, S. J. (2002). Why OrfY? Characterization of MmoD, a long overlooked component of the soluble methane monooxygenase from Methylococcus capsulatus (Bath). J. Biol. Chem. 277, 5858–5865. Murrell, J. C. (1993). Molecular biology of methane oxidation. In “Microbial Growth on C1 Compounds,” (J. C. Murrell and D. P. Kelly, eds.), pp. 109–120. Intercept Press, Andover, UK. Murrell, J. C., Gilbert, B., and McDonald, I. R. (2000). Molecular biology and regulation of methane monooxygenase. Arch. Microbiol. 173, 325–332. Oakley, C. J., and Murrell, J. C. (1988). nifH genes in the obligate methane oxidising bacteria. FEMS Microbiol. Lett. 49, 53–57. Pilkington, S. J., and Dalton, H. (1990). Soluble methane monooxygenase from Methylococcus capsulatus Bath. Methods Enzymol. 188, 181–190. Semrau, J. D., DiSpirito, A. A., and Yoon, S. (2010). Methanotrophs and copper. FEMS Microbiol. Rev. 34, 496–531. Smith, T. J., and Dalton, H. (2004). Biocatalysis by methane monooxygenase and its implications for the petroleum industry. In “Petroleum Biotechnology, Developments and Perspectives,” (R. Vazquez-Duhalt and R. Quintero-Ramirez, eds.), pp. 177–192. Elsevier, Amsterdam. Smith, T. J., and Murrell, J. C. (2009). Methanotrophy/methane oxidation. In “Encyclopedia of Microbiology,” (M. Schaechter, ed.)vol. 3, , pp. 293–298. Elsevier, Amsterdam. Smith, T. J., Slade, S. E., Burton, N. P., Murrell, J. C., and Dalton, H. (2002). Improved system for protein engineering of the hydroxylase component of soluble methane monooxygenase. Appl. Environ. Microbiol. 68, 5265–5273. Stafford, G. P., Scanlan, J., McDonald, I. R., and Murrell, J. C. (2003). rpoN, mmoR and mmoG, genes involved in regulating the expression of soluble methane monooxygenase in Methylosinus trichosporium OB3b. Microbiology 149, 1771–1784. Stanley, S. H., Prior, S. D., Leak, D. J., and Dalton, H. (1983). Copper stress underlies the fundamental change in intracellular location of methane monooxygenase in methaneoxidizing organisms—Studies in batch and continuous cultures. Biotechnol. Lett. 5, 487–492. Stirling, D. I., and Dalton, H. (1979). Purification of the methane monooxygenase from extracts of Methylosinus trichosporium OB3b and evidence for its similarity to the enzyme from Methylococcus capsulatus (Bath). Eur. J. Biochem. 96, 205–212. Theisen, A. R., Ali, M. H., Radajewski, S., Dumont, M. G., Dunfield, P. F., McDonald, I. R., Dedysh, S. N., Miguez, C. B., and Murrell, J. C. (2005). Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol. Microbiol. 58, 682–692. Trotsenko, Y. A., and Murrell, J. C. (2008). Metabolic aspects of aerobic obligate methanotrophy. Adv. Appl. Microbiol. 63, 183–229. West, C. A., Salmond, G. P. C., Dalton, H., and Murrell, J. C. (1992). Functional expression in Escherichia coli of protein B and protein C from soluble methane monooxygenase of Methylococcus capsulatus (Bath). J. Gen. Microbiol. 138, 1301–1307.

C H A P T E R

T E N

Single Cell Methods for Methane Oxidation Analysis Michael C. Konopka,* Sarah McQuaide,† David S. Ojala,* Marina G. Kalyuzhnaya,‡ and Mary E. Lidstrom*,‡ Contents 1. Introduction 2. Methods 2.1. Flow cytometry-based redox sensing of actively metabolizing microbes 2.2. Respiration response imaging 2.3. Respiration detection system: Microobservation chamber Acknowledgment References

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Abstract Respiration is widely used for evaluation of the metabolic capabilities or physiological state of the microbial culture. This chapter describes novel approaches for characterization of respiration at a single cell level: (1) flow cytometry-based redox sensing (FCRS) of actively metabolizing microbes; (2) respiration response imaging (RRI) for real-time detection of substrate stimulated redox responses of individual cells; (3) respiration detection system: microobservation chamber (RDS: MC), a single cell analysis system for carrying out the physiological and genomic profiling of cells capable of respiring C1 compounds. The techniques are suitable for description of physiological heterogeneity among cells in a single microbial population and could be used to characterize distribution of methylotrophic ability among microbial cells in the natural environmental samples.

* Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Department of Electrical Engineering, University of Washington, Seattle, Washington, USA Department of Microbiology, University of Washington, Seattle, Washington, USA

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Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00010-3

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2011 Elsevier Inc. All rights reserved.

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1. Introduction Each ecological niche is characterized by a set of specific biogeochemical processes that are mediated largely by microbes. One of the key objectives in environmental microbiology is to understand which bacteria are involved in which biogeochemical processes. Rapid advances in modern molecular methods such as the whole genome community sequencing approach open new approaches to studying microbial ecology (Allen and Banfield, 2005; Handelsman, 2004; Hugenholtz and Tyson 2008). However, in most cases, matching a function of interest (via a known gene) to a specific organism or population (via a phylogenetic marker) remains a formidable task. To truly understand the role of microbes in the environment, it is important to correlate observed environmental perturbation with the structure of the microbial population that is active at the time when the change is observed, and then reconsider genomic sequences in the context of physiological data. Methods that would allow for detection of active members of the microbial community, as well as methods that would monitor fluctuations in specific microbial activities in response to environmental perturbations in real time are important tools for environmental microbiology. Here, we describe three approaches for characterization of microbial functions at “close-to-in situ” settings: (1) flow cytometry-based redox sensing (FCRS) of actively metabolizing microbes; (2) respiration response imaging (RRI) for real-time detection of methylotrophic abilities at a single cell level; (3) respiration detection system: microobservation chamber (RDS: MC), a single cell analysis system for carrying out the physiological and genomic profiling of cells capable of respiring C1 compounds. An overview of our approaches is presented in Fig. 10.1. l. Flow cytometry redox sensing (FCRS) of actively metabolizing microbes

Separate active microbial population using FC sorting Environmental sample

Enrichment Genotyping//sequencing lll. Respiration detection system: microobservation chamber Single-cell physiological profiling on glass chip containing arrays of microwells

Cell transfer

Cultivation MDA//genotyping Whole genome sequencing

ll. Respiration response imaging Use fluorescence imaging microscopy to monitor cell respiration in incubation chamber before and after the addition of C1 stimuli

Figure 10.1 Overview of the single cell methods described in this chapter.

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Approaches (1) and (2) utilize fluorescence redox probes, and the third approach allows targeted detection of O2-dependant respiratory abilities of microbes at a single cell level.

2. Methods 2.1. Flow cytometry-based redox sensing of actively metabolizing microbes 2.1.1. Overview Detection of respiratory activity of cells has been a prominent technique for estimating the productivity and substrate utilization potential of microbial populations for some time (Bernard et al., 2000, del Giorgio et al., 1997). A number of redox probes have been tested in direct visualization of actively respiring microbes, the most widely used being 2-(p-iodophenyl)3-(p-nitrophenyl)-5-phenyl tetrazolium chloride (INT), 5-cyano-2,3ditolyl tetrazolium chloride (CTC), and more recently, RedoxSensor Green (RSG, Molecular Probes, Invitrogen). The principle is that these compounds are reduced to their respective colored (red crystalline INT– formazan; Zimmermann et al., 1978) or fluorescent (CTC–formazan, RSG; Gray et al., 2005; Rodriguez et al., 1992) compounds, as a result of modification by an active electron transport system (ETS) of respiring organisms. One of the major disadvantages of the IMT- or CTC-based assays is the suppression of bacterial metabolism by the end product, formazan (Ullrich et al., 1996). However, RSG does not appear to affect core cell functions (Gray et al., 2005, Kalyuzhnaya et al., 2008), and thus presents a step forward in this field. Recently, we described an approach based on the use of RSG, as a dynamic fluorescence indicator of bacterial reductase activity in combination with substrate stimulation and fluorescence activated flow cytometry (FC)/cell sorting (Kalyuzhnaya et al., 2008). This approach, named here FCRS, could be used for a quick evaluation of the physiological state of cells, such as active metabolism versus substrate limitation/starvation, as well as for functional profiling of natural microbial populations. 2.1.2. Evaluation of RedoxSensor Green dye performance The RSG staining protocol has been optimized on eight pure cultures of methylotrophic bacteria: (1) Type I methanotrophs Methylococcus capsulatus, and Methylomonas sp. LW13; (2) Type II methanotrophs Methylosinus trichosporium OB3b; (3) alpha-proteobacterial methylotroph Methylobacterium extorquens; (4) beta-proteobaterial methylotrophs Methylobacillus flagellatus, Methylotenera mobilis, Methyloversatilis universalis. For all cultures, 1 ml of RSG stock solution (1 mM solution in DMSO, Component A of the BacLightTM RedoxSensorTM Green Vitality Kit, B34954, Invitrogen) per

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1 ml sample of exponentially grown cells (105–106 cells/ml) was found to be an appropriate staining concentration. Ten minutes were sufficient to stain all tested cell samples, and only minor changes in the total cell count (< 0.01%) could be observed when extended to 20–30 min incubations. 2.1.3. Evaluation of the physiological state of cells by flow cytometry RSG staining is suitable for a quick evaluation of the physiological state of microbial culture, such as active growth, starvation, or stress by FC. Unstained and single-color stained control samples are included for each strain and the instrument settings are adjusted with respect to sample properties (Table 10.1). A significant portion (75–94%) of cells from an exponentially grown culture display a high fluorescence signal after staining with RSG. As a rule, starved cultures display a significant drop in the count of RSG-positive cells (< 20%). A steady increase in fluorescence could be observed after addition of electron donors (Kalyuzhnaya et al., 2008). The intensity of fluorescence differs between different cultures. For example, methylotrophic beta-proteobacteria, such as of M. flagellatus and M. mobilis, produce a very bright signal. A subpopulation of actively grown cultures (10–20%) did not move into a fluorescent state even after prolonged incubation with the carbon/energy source. In order to further describe this subset and distinguish between live, dormant, and dead cells, an additional staining with propidium iodide (PI; Component B of the BacLightTM RedoxSensorTM Green Vitality Kit, B34954, Invitrogen) was performed. The majority of the cells that were negative for RSG staining and demonstrate negative staining with PI were described as dormant. In addition to these physiological commonalities, we found that actively grown methanotrophic cultures have very low redox buffering abilities and are not able to switch to endogenous sources of electrons when external electron donors are eliminated. Samples of exponentially grown cultures of Methylomonas LW13 (Type I culture) or Methylosinus LW4 (Type II culture) stained with RSG in the presence of a C1 substrate showed a high count of RSG-positive (respiring) cells. However, a significant drop in the count of RSG-positive cells is observed if the carbon substrate is eliminated (Fig. 10.2). Cell respiration could be restored almost immediately by addition of methane or methanol. The physiological state of these cells can be attributed neither to the active state, as they are not respiring, nor to the dormant, since most of them display high respiration potential almost immediately after addition of electron donors. We named this part of the population as “cell-in-transition.” Since methane has low solubility and it may quickly escape from the medium when cells are transferred from the cultivation vial to the tube, this switching metabolic characteristic of methanotrophic cultures should be categorized and counted during respiration-based physiological evaluations.

Table 10.1 Summary of the FCRS method Dye characteristics RedoxSensor Green

Targeted function Redox activity (active cells) Excitation (max) 488 nm* argon ion laser Emission (max) 530* Dye stock 1 mM solution in solutions DMSO, Component A of the BacLightTM RedoxSensorTM Green Vitality Kit (Invitrogen, B34954) Detection FL1 (green) channel***

Propidium iodide*

FM1-43

Membrane integrity (dead cells) 488*

Membranes (all types of cells) 488 nm argon ion laser

610* 598 20 mM solution in 1.6 mM in water, Store at DMSO, Component
20  C, FM1-43Ò TM B of the BacLight Lipophilic Styryl Dye (Invitrogen, F35356) RedoxSensorTM Green Vitality Kit (Invitrogen, B34954) FL3 (red) FL2 (yellow)

Comments

* While double staining could provide an overview of the physiological state of the culture/population, PI treatment should not be used if cells are intended to be sorted for cultivation attempts due to high toxicity of the dye. ** Recommended by manufacturer for flow cytometry. *** Should be verified for each application specific instrument settings.

Sample preparation

Pure cultures

1. Cell extraction step is not needed. 2. Add 1 ml of the dye stock solution per 1 ml of cell sample containing 105–107 cell/ml. Water samples 1. Cell extraction step is not needed. 2. Add 1 ml of the dye stock solution per 1 ml of cell sample containing 105–107 cell/ml. Soil or Sediment 1. Add 15 ml of 0.2 PBS to 5 g of sample. Homogenize sample by using a samples PRO200 115 V (PROScientific) or similar homogenizer. Dilute the sample to 50 times with 0.2 PBS* and centrifuge for 1 min at 750g. To eliminate sediment particles, the blended material could be filtered through 5-mm NY20 filters (Millipore). The sample should contain 105–107 cell/ml. 2. Add 1 ml of the dye stock solution per 1 ml of sample. Flow cytometer settings

FSC/SSC settings

Different sets of 1–5 mm calibration beads could be used to adjust FSC and SSC settings, such as flow cytometry size calibration kit (Invitrogen, F13838); Calibration Beads 3 mm (Partec, 05-4018); SpheroRainbow calibration particles (Spherotech Inc RCP-205). The last set could be also used for adjustments of green-, orange-, or red-fluorescence channels. Set the amplification of the signals from forward and side scatters so that the 1–3 mm beads are in the center of the plot. Verify and adjust the setting with a pure culture of methanotrophic bacteria.

* Addition of organic extractants (Triton X100, Tween 20 or sucrose) may improve the efficiency of cell extraction. However, the extractants may also stimulate heterotrophic populations, and should not be used if samples are intended for functional profiling of the sample.

* Use an actively grown culture of Fluorescence methanotrophic bacteria as a control to adjust settings of the green (for RSG) and amplification yellow (FL1-43)-fluorescence channels* so that the signals from the stained cells settings appear in the center of the plot (Fig. 10.1). Use unstained culture to set sorting gates. Use killed cells** of your control culture. Set the amplification of the orange or redfluorescence channel so that the signals from the killed PI-stained cells appear in ** the center of the plot. Use unstained culture as a control.

Due to some overlap in emission spectra of RSG and FL1-43, in addition to dual staining, perform two control experiments with each dye individually (single stain). Do not use formaldehyde for cell fixation, as it could result in nonspecific increase of RSG-staining.

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A 1 mm beads

Methylomonas LW13

Methylosinus LW4

Methylomonas sp. LW13

B -RSG

+RSG

+RSG +methanol

Methylosinus sp. LW4

C -RSG

+RSG

+RSG +methanol

Figure 10.2 Flow cytometric analyses of Methylomonas sp. LW13 and Methylosinus sp. LW4 cells. Panel (A) shows FSC versus SSC density plots of 1 mm beads (left), Methylomonas sp. LW13 cells (center), Methylosinus sp. LW4 cells (right). Panel (B) shows FL1 versus SSC density plot of Methylomonas sp. LW13 cells before staining (left), cells stained with RSG (center), and cells incubated with methanol for 20 min and stained with RSG (right). Panel (C) shows FL1 versus SSC density plot of Methylosinus sp. LW4 cells before staining (left), cells stained with RSG (center), and cells incubated with methanol for 20 min and stained with RSG (right). SSC, side scatter; FSC forward scatter, FL1-green fluorescence.

2.1.4. Application of the FCRS for detection of actively metabolizing microbes in natural samples The FCRS has been applied for detection of active microbial populations from environmental samples and characterization of responses to specific environmentally relevant compounds/perturbations (Kalyuzhnaya et al., 2008).

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We found that most environmental water samples (marine or freshwater) could be stained directly; however, soil or sediment samples require an additional cell extraction step. A simple cell extraction method is described in Table 10.1 (see also Kalyuzhnaya et al., 2006). Examples of FC plots of the RSG-stained sediment samples are presented in Fig. 10.2. Each environmental sample was divided into two subsamples: experiment, a sample incubated for 2–12 h with a stimulus (methane, methanol, etc.); and control, a sample incubated without substrate. Concentrations of the substrates were determined empirically for each organism/ecosystem. We found that for freshwater sediment samples, the response of the microbial community was more pronounced in the lower range of C1 substrate concentrations (25–100 mM), with the initial response detected approximately 30 min after the addition of the substrate and the maximum response reached in approximately 2 h. After this, the number of fluorescing cells usually declines, eventually reaching the initial background level. After incubation, cell samples were stained with RSG, FM1-43, and PI. Stained and unstained samples were analyzed by using a CyFlow space flow cytometer. Examples of density plots of stimulated and control samples are shown in Fig. 10.3. We use FM1-43 staining to obtain complementary information about the total cell count (Fig. 10.3B and C). We found that

SSC

C

SSC

B

SSC

A

FL2

FSC

D

FL2

E

F F

R1

SSC

R1 SSC

SSC

R1

+CH4 -RSG

+RSG FL1

+RSG FL1

Figure 10.3 Flow cytometric analyses of cell population extracted from Lake Washington sediment. (A) FSC versus SSC density plot, (B) FL2 versus SSC density plot of untreated sample, (C) FL2 versus SSC density plot of sample stained with FM1-43, (D) FL1 versus SSC density plot of untreated sample, (E) FL1 versus SSC density plot of control sample stained with RSG, (F) FL1 versus SSC density plot of sample preincubated with methanol for 12 h and stained with RSG.

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other commonly used cell dyes, such as DAPI or SYBR Green, could be applied with a caveat for the enumeration of the total microbial population. Such dyes may inhibit cellular functions and thus should not be applied if cell samples are intended for sorting and cultivation. Unstained sample was used to determine background and select a sorting gate (Fig. 10.3D). Cells with green fluorescence above background were sorted (gate 1; Fig. 10.3E and F). So far, only the one-step substrate stimulation/fluorescence activated FC cell sorting approach was described (Kalyuzhnaya et al., 2008). The major limitation of this approach is that the sorted sample contains cells that were already active without substrate combined with cells that were stimulated by a substrate of interest. In order to deduce what microbial population responded to addition of stimuli, active cell fractions from both “stimulated” and “unstimulated” samples should be characterized side by side. Microbial species that were more pronounced in the stimulated samples, but were not detected or detected at a low level in the “unstimulated” samples, could be described as species activated by the substrate. 2.1.5. Future directions A new generation of high-speed cell sorters, such as the FACSAria or BD Influx, opens new possibilities for functional profiling of natural microbial communities. In theory, if a high-speed sorter is available for environmental studies, a two-step FCRS could be applied. In this case, the first round of cell sorting targets the “inactive fraction” of cells. These collected cells could then be used for stimulation experiments. Cells that became active after addition of stimuli could be collected in a second round of cell sorting. Such an approach may provide more specific enrichment of targeted population from complex samples. So far, the cell fraction selected via FCRS has been used for cultivation attempts, DGGE profiling, and PCR amplification of genes of interest. Other potential downstream applications for cell populations sorted based on activity stain include metagenomic sequencing and metaproteomic analysis.

2.2. Respiration response imaging 2.2.1. Overview RRI uses fluorescence microscopy and RSG to monitor cells over time to determine whether individual cells respond to a specific substrate. Since RSG is an indicator of bacterial reductase activity, an increase in fluorescence signal following stimulation would indicate an increase in respiration and therefore utilization of the substrate. As opposed to the FC method described in Section 2.1, this allows for specific single cells to be targeted for further physiological profiling (Fig. 10.1), although some throughput in the total number of cells will be sacrificed. While it is possible to make RRI

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measurements directly from environmental samples, to ensure proper settings and interpretation of the fluorescence profile, it is suggested that experimental conditions for a specific substrate first be optimized with a pure cell culture (Konopka et al., 2010). RRI can be performed on any fluorescence microscope system (epifluorescence or confocal) that has the ability to collect multiple images over time and has the proper excitation and emission filters for RSG. While it is possible to perform a manual analysis of the images, an analysis program will greatly increase the number of cells studied by this method. In the following paragraphs, a general protocol for RRI is described. Specific details about the procedure for data presented in Section 2.2.5 are presented elsewhere (Konopka et al., 2011). 2.2.2. Observation chamber preparations An observation chamber with a volume of at least 3 ml produces the best results for RRI measurements, as it provides sufficient volume to ensure that there are no effects from evaporation when it is covered. Cells are grown or collected using standard methods for the sample. To improve the adhesion of cells to the surface, the coverslip or slide in the cell observation chamber is pretreated with Poly-L-Lysine (Sigma-Aldrich, P8920) per the vendor’s protocols. One milliliter of sample is added directly to the coverslip in the chamber and allowed to stand for at least 10 min for the cells to adhere. The liquid is then removed from the chamber and replaced with 3 ml of the same medium or sample liquid (henceforth referred to as medium) that has been passed through a 0.22-mm filter (Millipore). For some environmental samples, especially sediment, it may be necessary to rinse the coverslip with two 1 ml aliquots of filtered media prior to the 3-ml addition to clear the samples that are not adhered to the coverslip. Alternatively, if cells are not readily adhering to the coverslip, it may be placed in a 24-well plate and centrifuged at 2250g for 15 min. The coverslip is removed from the centrifuge, rinsed if necessary, and placed in the observation chamber already containing the 3 ml of filtered media. An OD600 of 0.1–0.3 for pure cultures and direct deposition of environmental samples (lake sediment) produced an optimal seeding density based on past results (Konopka et al., 2011), although this may vary based on the sample. 2.2.3. RRI protocol RSG is part of the BacLightTM RedoxSensorTM Green Vitality kit (Invitrogen). It is rather viscous as it comes in the kit (1 mM), so diluting it by at least an order of magnitude into a filtered medium before adding it to the observation chamber helps with mixing and ensures that there is no image shift when it is added to the chamber while it is on the microscope. RSG is readily excited by a 488-nm laser line (or a corresponding excitation filter).

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The emission is collected with a filter covering all or part of the 500–550 nm range. Either an epifluorescence or confocal microscope may be used, but using an objective with at least 60 magnification ensures good detection of the cells. After placing the observation chamber on the microscope, the location (or locations) on the coverslip to be measured are selected and their focus determined. The experiment should last at least 5 h, so measuring multiple locations on a chip significantly improves the throughput. Using a marked or gridded chip will assist in finding the same locations for downstream analysis. Also, due to the experiment length, running an autofocus routine will help ensure that no significant drift in the z-axis occurs during the measurement. Especially with environmental samples, the RSG fluorescence signal may vary greatly from cell to cell. To ensure detection of the dimmest cells while also monitoring the change in fluorescence intensity of the brightest cells, it is necessary to take multiple images at several different detector gain settings for each time point. Otherwise, the brightest cells may have a measured intensity value that is at the maximum for the detector even before stimulation, making it impossible to detect an increase in RSG intensity in response to the substrate addition. Taking several sets of images with different gain (or sensitivity) settings for each time point ensures that both the brightest and dimmest cells can be measured and assessed. A transmission image is also helpful for assessing whether cells move or whether they arrive and leave the field of view. Time points should be spaced 5–10 min apart. The general protocol for taking fluorescence images for RRI after finding the locations and setting up the time course is as follows: 1. Take 2–3 data points without RSG or the substrate added. This is a control for autofluorescence. 2. Add the diluted RSG to a final concentration of 1 mM and continue taking data points for at least 30 min. Because of the high RSG viscosity, even when diluted, it is important to be careful during this step to ensure that you do not shift the placement of the coverslip in the field of view or alter the focus. Alternatively, it is possible to perform Step 1 separately and instead add the RSG off the microscope to ensure that no shift in focus occurs. Take at least 4–5 data points following RSG addition so it is possible to determine a stable prestimulation signal. 3. Add the substrate for stimulation. Exact concentrations of the substrate will depend on the sample type. Continue to take images for at least 4 h following the addition of the substrate. 4. When the RRI measurements are complete, downstream analysis may be performed. These could include PI for dead cell determination, FISH, or cell selection for other single cell analysis.

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2.2.4. Image processing A good image processing program will significantly shorten the time needed for RRI analysis. It should be able to find the location of a cell, track it frame by frame (including if there is movement or drift in the x–y plane), and calculate the cell intensity. This could be a commercial program, or freeware, such as ImageJ. For RRI analysis, we have also used a program written in Interactive Data Language (Research Systems Inc.) which is available from the lab Web site (https://depts.washington.edu/mllab/). It takes into account that multiple images are taken at different gains and matches up cells accordingly, as well as providing an interactive display of intensity/pixel versus time plots to allow immediate assessment of a cell’s response to stimulation. Before determining a detected cell’s response to stimulation, it is important to establish that the cell has been successfully tracked and that changes in the fluorescence data are not due to the cell landing in the focal plane after stimulation, cell movement, or cells intersecting one another. This can be accomplished by examining the transmission and fluorescence data. Cells that are acceptable can then be classified with regard to their response. 2.2.5. Sample classification data Cells that respond to a substrate have an intensity/pixel versus time plot that increases in fluorescence following stimulation. Other behaviors can be observed, including a constant fluorescence, slowly increasing or decreasing fluorescence, or combinations of behaviors. All of these are described in Table 10.2 with examples shown in Fig. 10.4. These results were from uncultured cells from Lake Washington sediment that were stimulated with methanol. Cells showing an increase in signal can be classified as cells of interest and then used in further testing.

2.3. Respiration detection system: Microobservation chamber 2.3.1. Overview Recently, a system was developed for measuring single-cell respiration rates for eukaryotic cells on a microscope (Molter et al., 2009). It features an array of microwells into which single cells are seeded, each well containing a platinum porphyrin that is used to measure the oxygen concentration by phosphorescence lifetime determinations. The wells are diffusionally sealed with a lid under load, and the consumption of oxygen is measured over time (Fig. 10.5). Adapting the wells and procedures to account for the smaller size and respiration rates of bacteria provides another method to utilize respiration in the analysis of single cells. The actual consumption of oxygen by a single cell is directly measured, as opposed to an indirect measurement

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Table 10.2 Classifying RRI outputs

Classification

Constant

Intensity/pixel fluorescence data

Intensity/pixel remains steady before and after stimulation for the experiment length. Increase Intensity/pixel is steady after RSG is added and then increases after substrate is added. If the experiment is sufficiently long, the intensity plateaus and remains constant. Slow increase/ The intensity/pixel shows a slow decrease slight increase or decrease after the substrate is added, but it is not large enough to assess whether significantly different from constant. Often never plateaus. Unclassifiable Erratic behavior of the data versus time, such as increasing and decreasing or random behavior.

Interpretation

Cell does not utilize the substrate or is in a dormant state. Cell responds to the substrate with an increase in respiration.

Not classified as stimulated. May be due to secondary effects or constant respiration.

Unknown behavior or the cell moved before and after stimulation. Not classified as stimulated.

in bulk cultures. Many of the experimental details have been described for the eukaryotic system, so here we highlight a few of the main differences for the system used for bacteria, as well as results from cells grown in pure culture. 2.3.2. Microwell array chips Microwell arrays and oxygen sensors are fabricated as previously described (Molter et al., 2009), with the exceptions that the first lithography process creates a 4  4 array of raised platforms 290 mm square, and the second lithography process creates a 4  4 array of microwells (2 pl) in volume on top of the raised platforms. To add the phosphorescent oxygen sensors to each microwell, the chips are cleaned in fuming H2SO4 for 30 min, rinsed, and dried. Chips are then cleaned in oxygen plasma for 10 min. A pipette tip is used without the pipette itself to dip into an Eppendorf tube containing 40 nm FluoSpheresÒ Platinum carboxylate-modified microspheres (Invitrogen) that had been sonicated for 10 min prior to use. The pipette tip is brought into contact with one platform of microwells, leaving a tiny droplet

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Figure 10.4 Examples of average intensity per pixel plotted versus time from RRI. Lake Washington sediment sample stimulated with methanol (stimulation time indicated by þ MeOH arrow). RedoxSensor Green (þRSG) was added after several frames as indicated by its labeled arrow. The Constant (upper left) is not utilizing the new substrate as there is no change in fluorescence after the methanol was added. Cells that do respond to the substrate can show differing times to reach maximum fluorescence, as Increase, 8 min (upper right) and Increase, 65 min (lower left) illustrate. The Slow Increase does not reach a plateau.

over the platform. This process is repeated with each platform of wells. The chip is left to dry for a few minutes and finally, the chips are put on a 170  C hot plate for 30 min to slightly melt the beads to ensure adherence. 2.3.3. Observation chamber preparation The microwell array chip with sensor is treated with Poly-L-Lysine (SigmaAldrich, P8920) as directed by the supplier prior to loading with cells. It is loaded with single cells by first placing it in a well on a 24-well plate and covering it with 2 ml of filtered medium (0.22 mm filter—Millipore). The cell concentration used to achieve the maximum loading of single cells in microwells varies with the culture, although in general, adding 60 ml of a 0.017 OD600 cell suspension produces a reasonable distribution of cells. The 24-well plate is centrifuged for 15 min at 2250g at 20  C to promote adhesion of the cells to the chip. The load chip is shaken in 1 ml of 0.22-mm filtered medium to dislodge cells that are not adhered. The chip is moved to an observation chamber cassette, covered with 2 ml of filtered medium, and placed in the microscope in preparation for measurements. Cells are stained with the

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Figure 10.5 Microobservation chamber for respiration detection. (A) The Microobservation chamber insert is part of a stage plate that can sit in any platen designed for microwell plates. A lid attached to a piston will lower to seal the wells on the chip with pressure. (B and C) The stage plate (1) has a window (6) that allows for observation with a microscope objective. The microobservation chamber (2) is centered above the window. A holder (5) helps to keep the chip (4) in place when the lid (3) comes down to seal the array of microwells. (D) Each chip has 16 arrays in a 4  4 arrangement. Each array is a 4  4 arrangement of microwells on a plateau raised above the rest of the chip to help in sealing the microwells. Each microwell is approximately 2 pL in volume and contains the platinum porphyrin that acts as the oxygen sensor. (E) Example of Methylomonas sp. LW13 cells labeled with FM1-43 to show wells containing single cells.

membrane dye FM1-43 (Table 10.1) to assist in counting the number of cells in a microwell and ensure only single cells are being measured. 2.3.4. Respiration measurements For a given array, it is possible to make multiple oxygen consumption rate measurements. First, the lid actuator is sealed with seven pounds of pressure and the same oxygen consumption measurement is made as outlined by Molter et al. (2009), except that a Zeiss LD Plan-Neofluar 63/0.75 NA Corr objective is used. The measurement lasts at least 20 min, with data points spaced 3 min apart to determine a good linear fit to the oxygen consumption. A z-stack of the microwells is taken with the confocal microscope (excitation wavelength of 488 nm and emission filter of 510–550 nm) to count the number of cells per well using the FM1-43 signal. A z-stack is required since cells often are located on several different planes of focus, or may be moving within the well. After a measurement is completed, the lid is raised and the sample is allowed to reoxygenate for a couple of minutes. The procedure could be repeated to perform another oxygen consumption measurement on this array or another array on the chip.

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Figure 10.6 Oxygen consumption plot, for example, Methylomonas sp. LW13 cells 1 h after stimulation with 250 mM methanol. Filled symbols are wells containing a single cell with respiration rates as follows: 68 amol/min cell (square), 37 amol/min cell (circle), and 19 amol/min cell (triangle). The open diamond is the plot from a well with no cells.

2.3.5. Sample respiration data Examples of this method being applied to Methylomonas sp. LW13 are presented below. These cultures were grown in NMS1 media under 50/50 methane/air at 30  C and stimulated with 250 mM methanol 1 h prior to oxygen consumption measurements. Figure 10.6 illustrates typical single-cell oxygen consumption measurements for several single cells (filled symbols), as well as an empty well (open symbols). Respiration rates are determined for a single bacterial cell in attomol (amol) O2 per cell per minute based on the slope of the oxygen levels in the well versus time and the volume of the sealed well. The measurements are calibrated to known oxygen levels. Respiration rates can vary from cell to cell, as in the example provided, which shows a threefold difference in single-cell respiration rate between the highest and the lowest respiring cell.

ACKNOWLEDGMENT This work was supported by the U. S. Department of Energy (DE-PS02-07ER07-14).

REFERENCES Allen, E. E., and Banfield, J. F. (2005). Community genomics in microbial ecology and evolution. Nat. Rev. Microbiol. 3, 489–498. Bernard, L., Courties, C., Servais, P., Troussellier, M., Petit, M., and Lebaron, P. (2000). Relationships among bacterial cell size, productivity, and genetic diversity in aquatic environments using cell sorting and flow cytometry. Microb. Ecol. 40, 148–158.

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Del Giorgio, P. A., Cole, J. J., and Cimbleris, A. (1997). Respiration rates in bacteria exceed phytoplankton production in unproductive aquatic systems. Nature 385, 148–151. Gray, D. R., Yue, S., Chueng, C. Y., and Godfrey, W. (2005). Bacterial vitality detected by a novel fluorogenic redox dye using flow cytometry. Presented at the American Society of Microbiology Meeting, Atlanta, GA. Handelsman, J. (2004). Metagenomics: Application of genomics to uncultured microorganisms. Microbiol. Mol. Biol. Rev. 68, 669–685. Hugenholtz, P., and Tyson, G. W. (2008). Microbiology: Metagenomics. Nature 455, 481–483. Kalyuzhnaya, M. G., Zabinsky, R., Bowerman, S., Baker, D. R., Lidstrom, M. E., and Chistoserdova, L. (2006). Fluorescence in situ hybridization-flow cytometry-cell sortingbased method for separation and enrichment of Type I and Type II methanotroph populations. Appl. Environ. Microbiol. 72, 4293–4301. Kalyuzhnaya, M. G., Lidstrom, M. E., and Chistoserdova, L. (2008). Real-time detection of actively metabolizing microbes by redox sensing as applied to methylotroph populations in Lake Washington. ISME J. 2, 696–706. Konopka, M. C., Strovas, T. J., Ojala, D. S., Chistoserdova, L., Lidstrom, M. E., and Kalyuzhnaya, M. G. (2011). Respiration response imaging for real-time detection of microbial function at the single-cell level. Appl. Environ. Microbiol. 77, 67–72. Molter, T. W., McQuaide, S. C., Suchorolski, M. T., Strovas, T. J., Burgess, L. W., Meldrum, D. R., and Lidstrom, M. E. (2009). A microwell array device capable of measuring single-cell oxygen consumption rates. Sens. Actuators B Chem. 135, 678–686. Rodriguez, G. G., Phipps, D., Ishiguro, K., and Ridgway, H. F. (1992). Use of a fluorescent redox probe for direct visualization of actively respiring bacteria. Appl. Environ. Microbiol. 58, 1801–1808. Ullrich, S., Karrasch, B., Hoppe, H., Jeskulke, K., and Mehrens, M. (1996). Toxic effects on bacterial metabolism of the redox dye 5-cyano-2, 3-ditolyl tetrazolium chloride. Appl. Environ. Microbiol. 62, 4587–4593. Zimmermann, R., Iturriaga, R., and Becker-Birck, J. (1978). Simultaneous determination of the total number of aquatic bacteria and the number thereof involved in respiration. Appl. Environ. Microbiol. 36, 926–935.

C H A P T E R

E L E V E N

Methanotroph Outer Membrane Preparation Odd A. Karlsen, Frode S. Berven,1 Harald B. Jensen, and Anne Fjellbirkeland2 Contents 168 170 170 170 172 172 172 172 173 173 175

1. Introduction 2. Isolation of OMs 2.1. M. capsulatus (Bath) culture conditions 2.2. Isolation of OM and associated proteins 3. Enrichment of Integral and Tightly Associated OMPs 3.1. Releasing proteins loosely associated to the OM 4. Extraction of Surface-Associated Proteins 4.1. Isolation of loosely associated cell-surface proteins 5. Biotin Labeling of Surface-Exposed OMPs 5.1. Biotin labeling of intact cells References

Abstract All presently known methanotrophs are gram-negative bacteria suggesting that they are surrounded by a two-layered membrane: an inner or cytoplasmic membrane and an outer membrane. In the methanotroph Methylococcus capsulatus (Bath), separation of the two membranes has allowed studies on protein and lipid composition of the outer membrane. Its outer membrane can be isolated from purified cell envelopes by selective solubilization of the inner membranes with the detergent Triton X-100. The proteins associated with the outer membrane can further be fractionated into integral and tightly associated proteins and peripheral loosely associated proteins. We present here protocols for this fractionation and show how the proteins associated with the outer leaflet of the outer membrane can be isolated and identified by whole-cell biotin surface labeling. Department of Molecular Biology, University of Bergen, Bergen, Norway Present address: Department of Biology/Proteomic Unit (PROBE), Department of Biomedicine, University of Bergen, Bergen, Norway 2 Present address: Department of Biology, Centre for Geobiology, University of Bergen, Bergen, Norway 1

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00011-5

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2011 Elsevier Inc. All rights reserved.

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1. Introduction The gamma-proteobacterium Methylococcus capsulatus (Bath) is one of the most extensively studied methanotrophs. It is ubiquitous in nature and has frequently been used in commercial pilot studies involving methanotrophs (Larsen and Joergensen, 1996; Murrell, 2010). Electron micrographs show that its cell envelope contains a three-layered structure (Fjellbirkeland et al., 1997). This is a characteristic feature of the gram-negative cell envelope (Silhavy et al., 2010), and, starting from the cell exterior, the three layers comprise the outer membrane (OM), the peptidoglycan layer, and the inner membrane (IM). In methanotrophs, extensive intracytoplasmic membranes can also be seen by electron microscopy. These are produced when cells are grown at high copper-to-biomass ratios (Prior and Dalton, 1985). They are believed to be formed by invaginations of the IM and thus can be considered as a part of the methanotroph cell envelope. The OM of M. capsulatus can be separated from the IM both by the classical sucrose density gradient centrifugation technique and by methods involving treatment of cell envelopes with the detergent Triton X-100 (Fjellbirkeland et al., 1997; Jahnke et al., 1992). Both methods have been successfully applied on a range of gram-negative bacteria to separate OM and IM. The first method generally leaves the IM intact and is the method of choice when lipid distributions are to be studied. Jahnke et al. (1992) used this method to demonstrate that the OM of M. capsulatus is enriched in hopanoids and sterols. To date, this lipid combination is not known in any other biological membrane, including membranes of closely related methanotrophs (Cvejic et al., 2000). The second method for separating the OM and IM of gram-negative bacteria involves treating the cellular cell envelope with detergent. This method was first used by Schnaitman (1971a,b) to isolate the OM from Escherichia coli (Schnaitman (1971a,b)). The detergent (e.g., Triton X-100) selectively solubilizes the IM while leaving the OM intact. This method is frequently the choice when proteins of the OM are to be studied, as it relatively rapidly generates large volumes of membranes. Proteins of the OM are part of the interface between the bacterium and its environment and are essential for cells in colonization and interaction with other molecules and cells. These proteins are diverse in functions, including protection against environmental challenges, uptake of growth factors, bacterial interaction, and pathogenicity. The OM proteins (OMPs) can mainly be divided into two classes, lipoproteins and b-barrel proteins. While the b-barrel proteins are embedded in the gram-negative OM, the lipoproteins are attached to the OM through lipid anchors (Silhavy et al., 2010). In addition to these two types of OMPs, proteins may also be

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associated to the outer or inner leaflet of the OM as peripheral proteins, exposed to either the cellular surface or the periplasm, respectively. Subcellular fractionation allows elucidation of cellular localization of proteins; it reduces sample complexity, which also increases the probability of observing low abundance proteins. Subcellular fractionation is often combined with proteomics, and recently we described both the OM subproteome and the surfaceome of M. capsulatus (Berven et al., 2006; Karlsen et al., 2008). In this chapter, we describe how to isolate the cell envelope from M. capsulatus, and subsequently the OMs. Further, we show how to separate the integral OMPs from the loosely associated OMPs, that is, peripheral proteins associated through noncovalent ionic/electrostatic interactions. We also show how to release loosely associated proteins from the cellular surface and give a protocol for labeling surface-exposed OMPs in general. The fractionation scheme used to obtain the protein samples described in this chapter is depicted in Fig. 11.1.

M. capsulatus cells in NMS (Section 2.1) Centrifuge

Surface-protein labeling (Section 5.1) Intact cells

Resuspend

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Intact cells NaCl, centrifuge

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Cell wall

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Enrichment of integral and tightly associated OMPs (Section 3.1)

Outer membrane (OM) Na2CO3, centrifuge

Integral outer membrane proteins

Figure 11.1 Fractionation scheme. Schematic overview of the strategies for subfractionation and protein labeling described in this chapter. M. capsulatus cells are grown in nitrate mineral salts (NMS) medium as described in Section 2.1.

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2. Isolation of OMs 2.1. M. capsulatus (Bath) culture conditions M. capsulatus (Bath) NCIMB 11132 is generally grown in a chemostat with nitrate mineral salts medium (Whittenbury et al., 1970) under an atmosphere of air:methane (5:1). The growth temperature is maintained at 45  C, and the pH of the growth medium is continuously adjusted to 6.7 by automatic addition of 0.25 M HCl. The cultures are grown with constant agitation at 600 rpm and with a dilution rate of 0.05 h 1. The optical density of the cultures is generally sustained at approximately 8.0.

2.2. Isolation of OM and associated proteins The Triton X-100-based separation of the M. capsulatus cell envelope has proven to have similar effects as first observed for E. coli (Fjellbirkeland et al., 1997; Schnaitman, 1971a,b). Visual inspection of whole cells and total membrane fractions with electron microscopy revealed structures reminiscent of intracytoplasmic membranes (Fig. 11.2A and B). As the internal membranes most likely are the results of invaginations of the cytoplasmic membrane, they should be solubilized by Triton X-100. Accordingly, large sheets of membranes dominated the Triton X-100-insoluble material, and only trace amounts of stacked membrane structures were observed (Fig. 11.2C). Further, the Triton X-100-insoluble fraction prepared from M. capsulatus is depleted of the membrane-bound methane monooxygenase, contains all the labeled proteins from extrinsically labeled whole cells, and is A

B

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Figure 11.2 Electron microscopic appearance of M. capsulatus whole cells (A), total membranes (B), and Triton X-100-insoluble membranes (C). The total membranes were obtained by centrifugation of disrupted cells. The Triton X-100-insoluble membranes were recovered by centrifugation after Triton X-100 extraction of total membranes. The figure was adapted from Fjellbirkeland et al. (1997) with kind permission from Springer-Verlag.

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enriched in b-OH palmitic fatty acids of the lipopolysaccharide. Isolation of the OM is essentially performed as described in Fjellbirkeland et al. (1997) with a few modifications. This protocol has also been successfully applied on the Type I methanotroph Methylomicrobium album BG8 (unpublished results). 1. Cells are harvested by centrifugation at 10,000g for 30 min. 2. 1–2 g (wet weight) of cells are resuspended in 20 ml 50 mM Tris–HCl, pH 7.5, and opened by three passages through a French pressure cell at 1000 psi. 3. Whole cells and debris are subsequently removed by centrifuging three times at 5000g. 4. The total membranes are collected from the resulting supernatant by centrifugation at 100,000g for 1 h at 4  C. The soluble fraction will be enriched in cytoplasmic and periplasmic proteins. 5. The pellet is resuspended in 10 ml 50 mM Tris–HCl, pH 7.5, 5 mM MgCl2, and 2% Triton X-100 for solubilization of the IMs. The suspension is stirred at room temperature for 1 h. 6. The Triton X-100-insoluble material is harvested by centrifugation at 100,000g at 4  C for 1 h. 7. The resulting pellet is resuspended in 4 ml of 50 mM Tris–HCl, pH 7.5 and 5 mM MgCl2 and corresponds to the enriched OM fraction. The different fractions obtained during the procedure should be assessed by SDS-PAGE to evaluate the effect of separation (Fig. 11.3). kDa

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Figure 11.3 SDS-PAGE of proteins obtained during the fractionation of M. capsulatus. Samples of each step during the fractionation procedure were collected and comparable samples (10 ml of each 1 ml fraction) were analyzed. Lane 1, whole cells (W); lane 2, soluble fraction (S); lane 3, total membrane fraction (TM); lane 4, Triton X-100soluble membranes (enriched inner membrane fraction, IM); lane 5, Triton X-100insoluble membranes (enriched outer membrane fraction). Molecular mass markers are indicated to the left. The figure was reproduced from Karlsen et al. (2005) with kind permission from John Wiley & Sons, Inc.

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3. Enrichment of Integral and Tightly Associated OMPs 3.1. Releasing proteins loosely associated to the OM Purification of OM as described above will result in a fraction containing both integral proteins and proteins associated with the inner or outer surface of the membrane. The loosely associated proteins are attached through noncovalent ionic/electrostatic interactions to other components of the cell membrane, and they can generally be released by introducing high ionic strength to the buffers (Feldman et al., 1983). We use a protocol which involves treating the Triton X-100-insoluble membranes with 0.1 M Na2CO3, pH 11, to release the proteins loosely associated to the OM (Berven et al., 2006). This result in a pellet further enriched in integral and tightly associated OMPs. 1. Fractionate the bacterium to obtain the OMP pellet as described in Section 2.2 (see also Fig. 11.1). 2. Resuspend the OM pellet in ice-cold 0.1 M Na2CO3 (pH 11) by vortexing or water bath-based sonication until the OM pellet has fully solubilized. 3. Keep the fraction at 4 ºC for 1 h with agitation. 4. Centrifuge the fraction at 100,000g for 1 h at 4 ºC. 5. The pellet contains the carbonate insoluble OMP fraction enriched in integral and tightly associated OMPs.

4. Extraction of Surface-Associated Proteins 4.1. Isolation of loosely associated cell-surface proteins Introducing high ionic strength to whole, intact cells can be used to selectively isolate the proteins loosely associated with the OM on the cellular surface. Treating cells with Na2CO3 (Section 3.1) will result in cellular lysis, and we generally use high concentration of NaCl (0.5 or 1 M) to release the surface-exposed proteins from the cellular surface (Karlsen et al., 2005, 2008). 1. Cells are harvested by centrifugation at 5000g for 10 min and carefully resuspended in a small volume of 20 mM Tris–HCl, pH 7.4. 2. Centrifuge the cell suspension as described above, and collect the supernatant as a reference of a low ionic cell wash. 3. Carefully resuspend the pelleted cells in 20 mM Tris–HCl, 0.5 M NaCl, and incubate with rotation for 1–2 h at 4  C. Subsequently, collect the cells by centrifugation (5000g).

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kDa

1

2

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115.5 82.2 64.2

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Figure 11.4 SDS-PAGE analysis of protein fractions obtained from NaCl extraction of M. capsulatus surface proteins. Lane 1, whole cells prior to NaCl treatment; lane 2, NaCl extract; lane 3, NaCl-treated whole cells. The NaCl extract was concentrated 10 times using selective centrifugation (Amicon, 10 kDa molecular weight cutoff) prior to application on the gel. MopE, the large subunit of the methanol dehydrogenase (MeDH) and molecular mass markers are indicated. The figure was reproduced from Karlsen et al. (2008) with kind permission from John Wiley & Sons, Inc.

4. The resulting supernatant contains the 0.5 M NaCl-extracted proteins. Treated cells can be resuspended in 20 mM Tris–HCl and kept for comparison. Samples obtained during the procedure should be inspected with SDS-PAGE to reassure that cell lyses did not occur. The abundant protein, methanol dehydrogenase, is a good marker for leakage of proteins from the periplasm due to disruption of the OM and should not be present in the fraction that constitutes the NaCl-extracted proteins (Fig. 11.4).

5. Biotin Labeling of Surface-Exposed OMPs 5.1. Biotin labeling of intact cells Protein labeling is an important tool for studying subcellular localization of proteins. N-hydroxysuccinimide (NHS) esters form amine bonds with primary amines and can be linked to biotin for labeling of proteins.

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The biotin-tagged proteins can be purified or visualized with, for example, avidin affinity chromatography or avidin-horseradish peroxidase (HRP) assays. Several NHS-biotin derivates have properties for keeping the biotinylation process at the cellular surface. The hydrophilic Sulfo-NHS-biotin cannot permeate lipid bilayers and has previously been shown to selectively biotinylate surface proteins of E. coli and Helicobacter pylori (Bradburne et al., 1993; Sabarth et al., 2002). We have used the commercially available EZ-LinkÒ Sulfo-NHS-biotin (Pierce) to selectively label and visualize surface proteins of M. capsulatus using a procedure combined from Turner et al. (1997) and Sabarth et al. (2002). It is recommended to do the surface extraction in rather small volumes to ease handling and minimize cell lysis (1–5 ml). 1. Approximately 109 M. capsulatus cells are harvested by centrifugation at 5000g for 5 min at 4  C. 2. The pellet is carefully resuspended in ice-cold phosphate-buffered saline (PBS), 1 mM CaCl2, 0.5 mM MgCl2, 1.6 mM biotin. 3. The cell suspension is incubated on ice for 15 min to saturate cells with biotin. If an active cellular uptake of biotin occurs, this preincubation will avoid transport of NHS-biotin into the periplasm and unwanted labeling of periplasmic proteins. 4. Cells are collected by centrifugation as described above and washed three times with PBS, 1 mM CaCl2, and 0.5 mM MgCl2. 5. The cell pellet is resuspended in PBS, 1 mM CaCl2, 0.5 mM MgCl2, and 200 mM freshly made Sulfo-NHS-biotin and incubated on ice for 15 min. 6. The cells are collected by centrifugation as described above and washed three times with 50 mM Tris–HCl, 1 mM MgCl2, and 1 mM CaCl2 to stop the reaction and remove unbound Sulfo-NHS-biotin. Cells can further be subjected to OM isolation or surface-protein extraction as described in Sections 2.2 and 4.1. 7. Proteins can be separated by either SDS-PAGE or 2D-PAGE and electrotransferred to a nitrocellulose/PVDF membrane. The membranes are blocked for 2 h at room temperature with 5% nonfat dried milk in PBS and 0.5% Tween-20. Further, the membrane is washed twice in PBS–Tween for 5 min, prior to incubation with streptavidine-biotinylated HRP conjugate (Pierce) diluted 1:3000 in PBS–Tween. After 1 h of incubation, the membrane is washed twice in PBS–Tween, and once in PBS, for 5 min of each wash. Biotinylated proteins are visualized with enhanced chemiluminescence (ECL). Biotin labeling of M. capsulatus whole cells combined with isolation of surface associated proteins is shown in Fig. 11.5.

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Figure 11.5 Biotin labeling of M. capsulatus surface-exposed proteins. (A and B) Lane 1, biotinylated whole cells; lane 2, biotinylated whole cells treated with 1 M NaCl; lane 3, 1 M NaCl extract of biotinylated cells. (A) Nitrocellulose membrane stained for total protein content using amidoschwarz. (B) The nitrocellulose membrane of (A) treated with streptavidin-biotinylated horseradish peroxidase complex (Pierce) and developed using ECL (GE healthcare) for detection of biotinylated proteins. Biotinylated proteins were detected prior to amidoschwarz staining. MopE, SACCP, MeDH, and bands corresponding to spots identified from the 2D-PAGE analyses are indicated and discussed in Karlsen et al. (2008). Molecular mass markers are indicated to the left. The figure was adapted from Karlsen et al. (2008) with kind permission from John Wiley & Sons, Inc.

REFERENCES Berven, F. S., Karlsen, O. A., Straume, A. H., Flikka, K., Murrell, J. C., Fjellbirkeland, A., Lillehaug, J. R., Eidhammer, I., and Jensen, H. B. (2006). Analysing the outer membrane subproteome of Methylococcus capsulatus (Bath) using proteomics and novel biocomputing tools. Arch. Microbiol. 184, 362–377. Bradburne, J. A., Godfrey, P., Choi, J. H., and Mathis, J. N. (1993). In vivo labeling of Escherichia coli cell envelope proteins with N-hydroxysuccinimide esters of biotin. Appl. Environ. Microbiol. 59, 663–668. Cvejic, J. H., Bodrossy, L., Kovacs, K. L., and Rohmer, M. (2000). Bacterial triterpenoids of the hopane series from the methanotrophic bacteria Methylocaldum spp.: Phylogenetic implications and first evidence for an unsaturated aminobacteriohopanepolyol. FEMS Microbiol. Lett. 182, 361–365. Feldman, R. A., Wang, E., and Hanafusa, H. (1983). Cytoplasmic localization of the transforming protein of Fujinami sarcoma virus: Salt-sensitive association with subcellular components. J. Virol. 45, 782–791.

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Fjellbirkeland, A., Kleivdal, H., Joergensen, C., Thestrup, H., and Jensen, H. B. (1997). Outer membrane proteins of Methylococcus capsulatus (Bath). Arch. Microbiol. 168, 128–135. Jahnke, L. L., Stan-Lotter, H., Kato, K., and Hochstein, L. I. (1992). Presence of methyl sterol and bacteriohopanepolyol in an outer-membrane preparation from Methylococcus capsulatus (Bath). J. Gen. Microbiol. 138, 1759–1766. Karlsen, O. A., Kindingstad, L., Angelskar, S. M., Bruseth, L. J., Straume, D., Puntervoll, P., Fjellbirkeland, A., Lillehaug, J. R., and Jensen, H. B. (2005). Identification of a copperrepressible C-type heme protein of Methylococcus capsulatus (Bath). A member of a novel group of the bacterial di-heme cytochrome c peroxidase family of proteins. FEBS J. 272, 6324–6335. Karlsen, O. A., Lillehaug, J. R., and Jensen, H. B. (2008). The presence of multiple c-type cytochromes at the surface of the methanotrophic bacterium Methylococcus capsulatus (Bath) is regulated by copper. Mol. Microbiol. 70, 15–26. Larsen, J., and Joergensen, L. (1996). Reduction of RNA and DNA in Methylococcus capsulatus by endogenous nucleases. Appl. Microbiol. Biotechnol. 45, 137–140. Murrell, J. C. (2010). The aerobic methane oxidizing bacteria (methanotrophs). In “Handbook of Hydrocarbon and Lipid Microbiology,” (K. N. Timmis, ed.). Springer-Verlag, Berlin. Prior, S., and Dalton, H. (1985). The effect of copper ions on membrane content and methane monooxygenase activity in methanol-grown cells of Methylococcus capsulatus. J. Gen. Microbiol. 131, 155–163. Sabarth, N., Lamer, S., Zimny-Arndt, U., Jungblut, P. R., Meyer, T. F., and Bumann, D. (2002). Identification of surface proteins of Helicobacter pylori by selective biotinylation, affinity purification, and two-dimensional gel electrophoresis. J. Biol. Chem. 277, 27896–27902. Schnaitman, C. A. (1971a). Effect of ethylenediaminetetraacetic acid, Triton X-100, and lysozyme on the morphology and chemical composition of isolate cell walls of Escherichia coli. J. Bacteriol. 108, 553–563. Schnaitman, C. A. (1971b). Solubilization of the cytoplasmic membrane of Escherichia coli by Triton X-100. J. Bacteriol. 108, 545–552. Silhavy, T. J., Kahne, D., and Walker, S. (2010). The bacterial cell envelope. Cold Spring Harb. Perspect. Biol. 2, a000414. Turner, M. S., Timms, P., Hafner, L. M., and Giffard, P. M. (1997). Identification and characterization of a basic cell surface-located protein from Lactobacillus fermentum BR11. J. Bacteriol. 179, 3310–3316. Whittenbury, R., Phillips, K. C., and Wilkinson, J. F. (1970). Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61(2), 205–218.

C H A P T E R

T W E LV E

Overexpression and Purification of the Particulate Methane Monooxygenase from Methylococcus capsulatus (Bath) Sunney I. Chan,*,† H.-Hoa T. Nguyen,‡ Kelvin H.-C. Chen,§ and Steve S.-F. Yu* Contents 1. Introduction 2. Overproduction of pMMO 2.1. A flow reactor for the overproduction of pMMO 2.2. Optimal conditions for the culturing of the bacteria 2.3. Isolation of pMMO-enriched membranes 3. Isolation and Purification of pMMO from the pMMO-Enriched Membranes 3.1. Direct transfer into detergent micelles followed by gel filtration (Method 1; Yu et al., 2003a) 3.2. Solubilization of the pMMO-enriched membranes by dodecyl b-D-maltoside and fractionation of the detergent–protein complex by ion-exchange chromatography (Method 2; Nguyen et al., 1998) 3.3. Reconstitution of the pMMO solubilized in dodecyl b-D-maltoside micelles into lipid vesicles 3.4. Methods of pMMO-purification employed by other research groups 4. Characterization of the Purified pMMO-Detergent Complex 4.1. Chemical composition of the purified pMMO-detergent complex 4.2. Specific activities of the purified pMMO Acknowledgments References

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* Institute of Chemistry, Academia Sinica, Taipei, Taiwan Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena, California, USA Transmembrane BioSciences, Pasadena, California, USA } Department of Chemical Biology, National Pingtung University of Education, Pingtung, Taiwan { {

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00012-7

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2011 Elsevier Inc. All rights reserved.

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Abstract The particulate methane monooxygenase (pMMO) is a multi-copper enzyme that mediates the facile conversion of methane to methanol in methanotrophic bacteria. As a membrane-bound multi-subunit metalloprotein, the highly active protein has been difficult to isolate and purify to homogeneity for biochemical and biophysical studies. In this chapter, we describe a method to overexpress pMMO with good specific activity in high yields in the intracytoplasmic membranes of the host organism, together with two protocols to isolate and purify the enzyme from pMMO-enriched membranes without loss of the copper cofactors and enzymatic activity.

1. Introduction The particulate methane monooxygenase (pMMO) is a membranebound enzyme in methanotrophic bacteria that mediates the facile conversion of methane to methanol using dioxygen as co-substrate (Balasubramanian et al., 2010; Chan and Yu, 2008). It is the first enzyme in the C1 metabolic pathway in these microorganisms. With a turnover frequency approaching 1 molecule of methane per second per enzyme, it is perhaps the most active catalyst known capable of accomplishing this difficult chemistry (Chan et al., 2007; Yu et al., 2003a). Accordingly, there is considerable interest in uncovering the methane C–H bond activation mechanism in order to develop a catalyst for industrial applications (Chen et al., 2007). Aside from methane, pMMO also hydroxylates a number of straight-chain hydrocarbons and epoxidates related alkenes with comparable efficiency (Elliott et al., 1997). The reaction is highly regio-specific and stereo-selective. It has been established that pMMO hydroxylates methane, ethane, and the secondary carbon(s) in propane, n-butane, and n-pentane with total retention of configuration at the carbon-atom oxidized (Elliott et al., 1997; Wilkinson et al., 1996; Yu et al., 2003b). Thus, the “O-atom” transfer from the activated dioxygen to the C–H bond in the organic substrate at the active site of the enzyme is concerted, that is, a nonradical mechanism. The epoxidation of propene, 1-butene, and cis- and trans-2-butene is also concerted and occurs without epimerization during the hydrophilic syn-addition to the C–C double bond (Ng et al., 2008). The pMMO is a highly complex enzyme comprised of three distinct subunits encoded by the genes pmoA, pmoB, and pmoC with an abg arrangement (Basu et al., 2003; Lieberman et al., 2003; Yu et al., 2003a). It is a multi-copper protein containing as many as 15 copper ions, although the number and type of copper cofactors is still under debate (Basu et al., 2003; Choi et al., 2003; Lieberman et al., 2003; Yu et al., 2003a).

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As a membrane-bound enzyme, it has been notoriously difficult to purify because of its instability outside the lipid bilayer and the tendency of the protein to lose its essential metal cofactors. For this reason, pMMO has resisted both initial identification and subsequent isolation and purification for biochemical and biophysical characterization. Nevertheless, good progress has been made in recent years toward elucidating the structure and function of this tantalizing enzyme. In-depth studies on a system as complex as the pMMO is only possible when one has access to non-limiting quantities of the functional protein purified to homogeneity. It goes without saying that the protein isolation and purification must be carried out with adequate controls to ensure the structural integrity before we can embark on biochemical and biophysical measurements. The purified protein must have all the subunits intact and structurally folded into the physiological functioning conformation. It must also contain all the essential metal cofactors. These requirements present significant challenges to the protein chemist for a membrane-bound metalloprotein like pMMO. In this chapter, we describe efforts to overexpress highly homogeneous pMMO in high yields in the intracytoplasmic membranes of Methylococcus capsulatus (Bath) as well as methods that we have been developed to obtain the pMMO with high specific activity in sufficient quantities for biochemical and biophysical studies.

2. Overproduction of pMMO M. capsulatus (Bath) harbors two sets of methane monooxygenase (MMO) genes (Kao et al., 2004). One set codes for the soluble methane monooxygense (sMMO), a nonheme iron-containing soluble cytosolic enzyme with diverse substrate specificity (Feig and Lippard, 1994; Lipscomb, 1994). The other encodes the pMMO, a multi-copper membrane-bound enzyme with narrow substrate specificity (Chan and Yu, 2008; Chan et al., 2004; Lieberman and Rosenzweig, 2005). The two MMO genes are differentially expressed. The sMMO is only expressed under conditions of low biomass and limiting copper ions in the growth medium. The expression of the sMMO gene is suppressed by copper ions. On the other hand, copper appears to be a transcriptional activator for the pMMO genes. When the copper concentration in the environment exceeds 2 mM, the pMMO is expressed, and the level of expression increases with the concentration of copper (Yu et al., 2003a). Quantitative proteomic studies of M. capsulatus (Bath) indicate that expression of the pMMO is accompanied by enhanced expression of the genes downstream of the pmmo gene cluster that are involved in the metabolism of the cell, such as those

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that regulate the biosynthesis of membrane lipids (Kao et al., 2004). The co-expression of these metabolic genes concomitant with the expression of the pmmo gene cluster ensures that the subunits of the overproduced pMMO are properly assembled in the milieu of a lipid membrane to accomplish the catalytic function with the specific activity expected. When the expression of the MMO genes in M. capsulatus (Bath) switches from sMMO to pMMO, the membrane lipids are produced in such great abundance that the overproduction of the pMMO is accompanied by the formation of an extensive network of intracytoplasmic membranes (Nguyen et al., 1994). As a membrane protein containing up to 15 copper cofactors, it is possible that some of the pMMO subunits produced are assembled without the full complement of copper ions when there is insufficient amounts of copper ions in the growth medium. This is evidently the case, as shown in Fig. 12.1. Here cultures of M. capsulatus (Bath) are grown in a fermentor at various initial concentrations of CuSO4 in the growth medium, and the residual copper concentration in the medium is assayed during the harvesting of the cells (vide infra). When the initial CuSO4 concentration is substantially <30 mM, all the copper in the growth medium is taken up by the cells. Under these conditions, the pMMO in the membrane is found to be heterogeneous in terms of the copper contents, with some protein molecules containing the full complement of copper ions, and others with fewer copper ions. The specific activity of the protein isolated from a preparation of these membranes reflects this heterogeneity in the copper content (Table 12.1). This is a common problem with the overexpression of

Copper concentration in the growth medium after harvest (mM)

35 30 25 20 15 10 5 0 0

10 20 30 40 50 Initial copper concentration in the NMS buffer (mM)

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Figure 12.1 Comparison between the copper contents in the NMS medium before the medium replenishment and during the harvesting of the cells. (Reproduced from Yu et al. (2003a). Copyright # American Society for Microbiology.)

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Table 12.1 The effects of copper concentration in the culture medium on the pMMO contents in the pMMO-enriched membranes, the average copper content of the pMMO proteins, and the specific activity of the pMMO-enriched membranes using NADH as the reductanta

a b

Cu (mM) in growth medium

pMMO/total protein (%)

Specific activity (nmol/ min/mg of protein)

Copper/ pMMOb

< 0.3 5 10 20 30 40 60

19 27 44 68 80 73 65

19.6 30.5 42.6 59.6 65.9 88.9 63.5

9.4 10.5 10.3 10.8 13.4 13.8 15.3

Taken from Yu et al. (2003a). Copyright # American Society for Microbiology. mol copper/mol of protein, determined assuming that the monomeric pMMO hydroxylase has a subunit composition of abg with a molecular mass of 99 kDa.

metalloenzymes. Of course, the missing copper cofactors could be replenished by simply adding additional CuSO4 to the harvested cells or to the isolated membranes. However, in the case of pMMO, which is a CuI enzyme (Chan and Yu, 2008; Chan et al., 2004, 2007), copper chaperones may be required to introduce the missing copper factors back to the unfilled sites.

2.1. A flow reactor for the overproduction of pMMO To obtain pMMO-enriched membranes from M. capsulatus (Bath) with high activity and in high yields, a method has been devised to process cell growth in a fermentor adapted with a hollow-fiber bioreactor that allows easy control and quantitative adjustment of the copper ion concentration in the NMS medium over the time course of the cell culture (Yu et al., 2003a). A schematic diagram of this flow reactor is shown in Fig. 12.2. This technical development allows M. capsulatus (Bath) cells to be grown in the presence of well-defined copper concentrations so that the cells are not subjected to gross variations in the copper ion levels during the time course of the culturing. In addition, the hollow-fiber membrane bioreactor offers a device to filter off used NMS medium in order to discard cellular waste and toxic metabolites from the cellular milieu, to replenish the cellular medium in order to maintain an essentially constant environment, and to adjust the copper concentration, if desired, in the growth medium in a controlled manner in increments that are not harmful to the cells. In the setup depicted in Fig. 12.2, the cell medium is continuously drained through the hollow-fiber membrane bioreactor consisting of a

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Valve (E) Valve (A)

Motor (G)

0.45 mM Filter (J)

0.45 mM Filter (K)

Peristaltic pump (M)

Air Valve (B)

Methane in air (20%) Valve (D) Agitate paddle (I)

Hollow-fiber membrane (F)

5-L Fermentor (H) Harvest output

Valve (C)

10-L Serum Bottle (L)

Filtrate

Figure 12.2 Schematic of a fermentor adapted with a hollow-fiber membrane bioreactor to culture M. capsulatus (Bath) for production of high-quality pMMO in high yields. (Reproduced from Yu et al. (2003a). Copyright # American Society for Microbiology.)

large number of tubular fiber membranes encased in a cylindrical shell with a pore size that is much smaller than the size of bacteria but large enough to allow passage of CuSO4 and small metabolites. In this dual reactor, the bulk of the bacterial cells are continuously flowing through the high-surface-area lumen of the fiber and the used NMS medium containing the waste and small molecules is filtered away from the cell medium across the hollowfiber membranes for removal. At the same time, the discarded medium is replenished with fresh growth medium with the controlled CuSO4 concentration to maintain the copper ions at a constant level. The combined fermentor-hollow-fiber-bioreactor has proven to be a high-capacity and a highly efficient system. With this system it is possible harvest more than 50 g of cells per day. In addition, the system is very robust, lasting for months.

2.2. Optimal conditions for the culturing of the bacteria Membranes enriched in the pMMO, with the three subunits of each protein properly assembled together with the full complement of 15 copper cofactors, are required before the purified enzyme could be obtained from these membranes with high activity and in high yields. The pMMO-enriched membranes can be obtained by growing the bacterial culture in a fermentor adapted with the hollow-fiber membrane bioreactor in a growth medium containing a copper concentration of 30–35 mM. At lower copper concentrations, there is insufficient copper in the growth

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medium to ensure that all the pMMO will be assembled with the full complement of copper ions. At copper concentrations in excess of 50 mM, the cells are not tolerant of the copper and do not grow well. 1. Start with cultures of M. capsulatus (Bath) (ATCC 33009). Before use in growing the bacteria, streak the cell cultures onto fresh Petri plates maintained in NMS medium (ATCC medium: 1306 nitrate mineral salts medium) and solidified with 1.7% agar under an atmosphere of 20% methane in air. 2. To initiate the bacterial growth, first transfer the cultures from the Petri plates to 250-mL flasks and subsequently to 2-L Erlenmeyer flasks containing, respectively, 30 and 300 mL of NMS medium containing a CuSO4 concentration of 30–35 mM in an atmosphere of 20% methane in air. 3. After 48 h incubation with continual shaking, use the resulting cell suspension to seed a fermentor (Fig. 12.2) with a 5-L vessel containing 3 L of NMS medium. When the turbidity of the culturing medium has reached 1.2–1.6 OD595, typically after a period of 24 h incubation, add additional NMS medium to increase the total culture volume to 5 L for further semi-continuous growth. It is usually required to replenish the cell-enriched medium with fresh NMS buffer by draining away the used medium in the bioreactor two to three times every 12–24 h. This media replenishment will keep the cell cycle in the middle-log phase to late-log phase over the said period. With M. capsulatus (Bath), it is possible to keep the cells growing at pH 6.8–7.4 for 12–24 h by agitating the cellular suspension in the range of 200–800 rpm and controlling the methaneand air-feeding rate (0.7–1.3 L/min) to maintain the dissolved oxygen content to 2–5% of the saturation. 4. After the cells have been cultured for 12–24 h and are ready for harvest, remove about 4 L of the medium with the assistance of the pre-equipped hollow-fiber filter. Drain three quarters of the resulting concentrated cell suspension out and centrifuge. Retain the harvested pellet for eventual isolation and purification of the pMMO. Filter the remainder of the concentrated cell suspension to almost dryness. Re-suspend these cells in the fermentor with 5 L fresh NMS medium injected through the inlet tube into the hollow-fiber filter for the next round of culturing. Typically, about 35–40 g of cells can be harvested during the above medium renewal process. During the next round of culturing, the cell density would usually recover to 10–12 g/L within the next 12 h.

2.3. Isolation of pMMO-enriched membranes 1. To isolate the pMMO-enriched membranes, suspend the harvested cell paste in 25 mM PIPES buffer at pH 7.0 (1.2 mL of buffer/g of cell paste) containing 0.01 mg/mL of DNase I.

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2. Pass the cell suspensions three times through a French pressure cell at 20,000 p.s.i. in order to separate the cytosolic and membrane fractions. Remove unlysed cells and cell debris by centrifugation at 27,000g for 20 min. Then, pellet the membrane fraction by ultracentrifuging the supernatant at 220,000g for 40 min. Discard the clear supernatant obtained after ultracentrifigation, namely, the cytosolic fraction. 3. The pelleted membranes often show distinct layers. Discard the minor bottom layer containing bluish and black materials and the thin white top layer. Collect only the middle cut, or the translucent intracytoplasmic membrane fraction, which constitutes the bulk of the membrane proteins. 4. Wash the translucent membranes by suspending them in washing buffer containing 25 mM PIPES, 5 mM ascorbate, 25 mg of catalase/mL (pH 7.0) using a Dounce homogenizer, re-pellet them by ultracentrifugation, and re-suspend in washing buffer of two to three times the volume of the original cell suspension. Repeat this process three to five times until the supernatant is virtually free of soluble proteins.

3. Isolation and Purification of pMMO from the pMMO-Enriched Membranes Following the above procedures, the pMMO is highly expressed in the membranes of M. capsulatus (Bath), accounting for  80% of the total intracytoplasmic membrane proteins and having a specific activity as high as 90 nmol of propylene oxide/min/mg of protein with NADH as the reductant (Yu et al., 2003a). The copper stoichiometry is 13 atoms per pMMO molecule. Analysis of other metal contents provided no evidence of zinc, and only traces of iron were present in the pMMO-enriched membranes. Two approaches have been developed to isolate and purify pMMO from the pMMO-enriched membranes (Nguyen et al., 1998; Yu et al., 2003a). Both methods begin with the solubilization of the membranes by the detergent dodecyl b-D-maltoside. In one protocol, the solubilization is followed by fractionation of the protein–detergent complexes according to molecular size of the micelles by gel filtration chromatography. In the other, the solubilized proteins are subjected to ion-exchange chromatography to separate membrane proteins of varying sizes and hydrophobicity. The gel filtration method is gentle and works well when the membrane protein of interest is present in abundance, which is the case when the pMMO-enriched membranes are obtained as described above. The second method is considerably harsher, but it has been shown to work, provided precautions are taken to ensure that the copper ions remain reduced and do not dissociate from the protein during the fractionation and chromatography steps.

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3.1. Direct transfer into detergent micelles followed by gel filtration (Method 1; Yu et al., 2003a) When high-quality pMMO accounts for 80–90% of the total proteins in the membranes, the bulk of the detergent–protein complexes will contain the competent pMMO. If a sufficient amount of detergent is used to solubilize the membrane so that [detergent]/[proteins]  1000, then the typical detergent–protein micelle particle would contain on the average only one protein molecule. The mass distribution of the micelles should also be fairly narrow given that most of the micelles would contain the abundant protein. Figure 12.3 shows the elution profile of the proteins after a preparation of the membranes from cells cultured and harvested at a copper concentration of 30 mM is solubilized in 5% (w/v) dodecyl b-D-maltoside (or 3 mg of detergent/mg of protein) and subjected to size exclusion column chromatography on a Sephacryl S-300HR column. The elution buffer contains 50 mM PIPES, 150 mM NaCl, 50 mM imidazole, 5 mM ascorbate, and 0.05% (w/v) dodecyl b-D-maltoside (pH 7.2). The strong band appearing around the elution volume of 50 mL corresponds to the purified pMMOdetergent complex. SDS-PAGE gels of cuts derived in the vicinity of this peak show the bands associated with the 45-, 27-, and 23-kDa (or a, b, and g) subunits of pMMO without contamination by other proteins. Thus, the gel filtration procedure outlined here leads to an apparently homogeneous preparation of pMMO-detergent complexes.

3.2. Solubilization of the pMMO-enriched membranes by dodecyl b-D-maltoside and fractionation of the detergent–protein complex by ion-exchange chromatography (Method 2; Nguyen et al., 1998) 1. The membrane suspension is first degassed by several vacuo/argon cycles. The membranes in storage buffer (in either low or high ionic strength buffer) are then treated with either solid or 20% stock solution of dodecyl b-D-maltoside (3–5%, w/v, or 2 mg of detergent/mg of protein). The mixture is mixed rigorously, and incubated on ice for 30 min to 1 h, then centrifuged at 18,000 rpm for 45 min to remove unsolubilized materials. The clear supernatant is the solubilized membranes, which is used in subsequent steps. 2. For rapid isolation of the pMMO (Method 2a), L-lysine agarose affinity chromatography could be used to remove positively charged and ironcontaining proteins. A lysine agarose column (Sigma) (20 cm2 cm) is first equilibrated with buffer containing 25 mM PIPES, 5 mM ascorbate, 200 mM CuSO4, and 0.05% (w/v) dodecyl b-D-maltoside (pH 7.2). Dithionite (5 mM) can be used in lieu of ascorbate; however, strict anaerobic protocol must be followed. The solubilized membrane

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97 kDa

α

66 kDa 45 kDa

β

30 kDa

20 kDa

γ

14 kDa Lane

Marker

A

B

C

D

E

↑

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300

Purified pMMO-detergent complex (~220 kDa)

200

20

40

60

80

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13.7 kDa

25 kDa

43 kDa

67 kDa

232 kDa

0

158 kDa

669 kDa

100 440 kDa

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400

120

Elution volume (mL)

Figure 12.3 The elution profile of the detergent-solubilized pMMO-enriched membranes from a Sephacryl S-300HR gel filtration column. Lane A: elution volume 46–48 mL; Lane B: elution volume 49–51 mL; Lane C: elution volume 52–54 mL; Lane D: elution volume 55–57 mL; Lane E: elution volume 58–60 mL. (Adapted from Yu et al. (2003a). Copyright # American Society for Microbiology.)

(2 mL) is then applied to the column and 0.5 mL effluent fractions are collected. Three to four fractions can be obtained. The flow-through fraction contains large pieces of the solubilized membranes and most of the positively charged proteins. A fast-moving fraction contains several

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proteins, including heme-containing proteins, but also pMMO (purity of  70% or higher). The next more slow-moving fraction constitutes the bulk of the solubilized membranes, and contains mostly the threesubunit form of the pMMO (purity of 90% or higher). Finally, a minor binding fraction can be eluted out of the column using buffer containing 50 mM PIPES, 100 mM NaCl (pH 7.2). This binding fraction consists of mostly heme-containing proteins, but also some residual pMMO. 3. For large-scale isolation of the pMMO (Method 2b), it is necessary to use anion-exchange chromatography to remove the positively charged and iron-containing proteins. The L-lysine agarose affinity column mentioned earlier is replaced by a DEAE-Sepharose Fast Flow column (Pharmacia). The column is first equilibrated with buffer containing 100 mM PIPES, 50 mM imidazole, 5 mM ascorbate, 200 mM CuSO4, 0.05% (w/v) dodecyl b-D-maltoside buffer at pH 7.2. Sucrose (200 mM) can also be included in this buffer but its effectiveness is not great. To carry out the subsequent purification of the pMMO under anaerobic conditions, the above setup and the gel must be degassed, dithionite (5 mM) must be added to the equilibrating buffer to remove dissolved residual dioxygen, and the isolation manipulation should be performed in an anaerobic chamber. Solubilized membranes are then applied to the column. The column is then washed with one column of the equilibrating buffer. The DEAESepharose Fast Flow column fractionates the solubilized membranes into four fractions. There are two flow-through fractions, a fast-moving fraction containing the so-called four subunit pMMO (or a mixture of two forms of pMMO) and slow-moving fraction containing positively charged proteins (proteins of high pI), a truncated form of pMMO, and other impurities. The binding proteins are eluted out using the above high ionic strength buffer with a NaCl (or NH4Cl) gradient from 0 to 200 mM. These proteins are separated into two fractions. The fraction eluted out at <100 mM NaCl contains mostly the pMMO as judged from SDS-PAGE assay (purity >90%). The second fraction elutes out of the column at higher salt concentration (>100 mM NaCl), containing a low pI, low molecular weight contaminant (22 kDa) and other minor impurities. The isolated pMMO proteins are then concentrated using Amicon ultrafiltration membranes (MW cutoff 50 or 100 kDa). The first binding fraction can be fractionated further using QEA-Sephadex A-50, albeit with a significant reduction in recovered activity. The QEA-Sephadex A-50 column is first equilibrated with buffer containing 50 mM PIPES, 50 mM NaCl, 50 mM imidazole, 200 mM sucrose, 5 mM ascorbate, 200 mM CuSO4, and 0.03% (w/v) dodecyl b-D-maltoside (pH 7.2). The binding fraction is then applied to the column. The pMMO is eluted out using a NaCl (or NH4Cl) gradient at around 200 mM NaCl as a light green band.

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3.3. Reconstitution of the pMMO solubilized in dodecyl b-D-maltoside micelles into lipid vesicles 1. To obtain the membrane lipids from the bacterial cells, the membranes are mixed with a methanol/chloroform (1:3, w/v) mixture, shaken vigorously, and decanted. The process is repeated a few times to ensure complete extraction. The extracts are combined, dried over anhydrous MgSO4, and decanted. After solvent removal, the crude yellowish lipids are dried and stored under vacuo. 2. Reconstitution of the pMMO is carried out as follows. A volume of the buffer containing 10–20 mg/mL of the isolated lipids is sonicated for 10–15 min to disperse the lipids and mixed with the purified protein in detergent-containing buffer (1:1 or 2:1, v/v, ratio). The resulting mixture is sonicated briefly for a few seconds to assure dispersion. As soon as the detergent-containing protein solution is added, the solution becomes clear. The mixture is then loaded into dialysis tubing (MW cutoff 50,000 or 100,000). The tube is dialyzed against a buffer at pH 7.2 containing 50 mM PIPES, 10 mM ascorbate, 200 mM CuSO4, 100 mM (NH4)2SO4 for 12 h with continuous stirring. The detergent can also be removed using BioBeads SM-2. A volume of 2–3 mL of the purified protein (30–50 mg/mL) is passed through a column (1 cm5 cm) of BioBeads and the eluate is concentrated using Amicon ultrafiltration membranes and mixed immediately with a sonicated lipid dispersion (10–20 mg/mL; lipid/protein 1:1 or 2:1, v/v). The mixture can be sonicated briefly for a few seconds to ensure dispersion. The reconstituted protein can be assayed immediately for activity and stored at 4  C or 80  C for later use.

3.4. Methods of pMMO-purification employed by other research groups Variations of Method 2b have been used by other research groups in the purification of pMMO from the membranes. Dispirito and coworkers have employed the DEAE-Sepharose Fast Flow column to separate pMMO from other proteins (Choi et al., 2003). The laboratory of Dalton has adopted the same protocol, but, to purify the pMMO further, this group has added an additional gel filtration step using a Superdex 200 column (Basu et al., 2003). Rosenzweig et al. begins the purification using a Source 30Q anionexchange column to remove contaminating proteins as well as lipids (Lieberman et al., 2003). Low molecular mass proteins are not resolved on the Source 30Q column, which are then separated by gel filtration on Sephacryl S200. In their isolation and purification of pMMO for protein crystallization and X-ray structural analysis, Rosenzweig and coworkers

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used ammonium sulfate precipitation followed by anion-exchange chromatography on a Source 30Q (Amersham Biosciences) column (Lieberman and Rosenzweig, 2005). In all of these protocols, the purification of the pMMO has typically not been carried out under anaerobic conditions. It is now evident that many of the CuI ions of the pMMO would become oxidized by the dioxygen in the buffer and be stripped off the protein during the column chromatography and ammonium sulfate fractionation/precipitation, with concomitant loss of enzymatic activity (Chan and Yu, 2008). Under these scenarios, the purified enzyme might contain 2–3 copper ions, and it is essentially inactive (Lieberman and Rosenzweig, 2005).

4. Characterization of the Purified pMMO-Detergent Complex 4.1. Chemical composition of the purified pMMO-detergent complex MALDI-TOF MS (Fig. 12.4) of the purified pMMO-detergent complex obtained by direct transfer of the pMMO from enriched membranes into detergent micelles followed by gel filtration (Method 1) reveals only three polypeptides with molecular masses of 42,785 kDa (subunit a or PmoB), 29,073 kDa (subunit g or PmoC), and 28,328 kDa (subunit b or PmoA), in excellent agreement with the peptide sequences predicted by the gene sequences in the National Center for Biotechnology Information database (Yu et al., 2003a). Assignment of the a, b, and g subunits on the SDS-PAGE gel to PmoB, PmoA, and PmoC, respectively, is made by gel in situ digestion peptide mass fingerprinting (Chan et al., 2004). The relative intensities of the peptide mass peaks are consistent with an equal stoichiometry of the three subunits in the pMMO-detergent complex. The purified pMMO-detergent complex elutes on the S-300HR size exclusion column at a position corresponding to a molecular mass of  220 kDa based on molecular mass calibration kits (Amersham Pharmacia Inc.). Evidently, the pMMO-detergent complex is isolated as an abg monomer with 240 detergent molecules. For comparison, there are 100 molecules per dodecyl b-D-maltoside micelle without the protein. Metal analysis of the purified pMMO-detergent complex obtained by Method 1 by ICP-MS yields average copper and iron contents of 0.143 and 0.002 mmol/mg of protein, respectively, or a copper content of 14.3 mol/ mol of protein and a copper-to-iron ratio of 80:1 (Yu et al., 2003a). Analysis of the CuKa X-ray absorption edge at 8984 eV indicates that 60% of the copper ions are CuI, as in the purified pMMO-enriched membranes (Nguyen et al., 1996; Yu et al., 2003a). 77 K EPR of the purified pMMO-detergent complex exhibits the same spectral features as the

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28,328.25

2000

% Intensity

1500

42,785.52

29,073.29

MALDI-TOF 1000

500

0 24,000

28,000

32,000

36,000

40,000

44,000

48,000

Mass (m/z)

Subunit

Molecular mass (predicted)

Molecular mass (measured)

PmoB1

42,786.04

42,785.52

PmoC1

29,063.66

29,073.29

PmoA1

28,376.16

28,328.25

Figure 12.4 Purified pMMO molecular mass identification by MALDI-TOF mass spectrometry. (Adapted from Yu et al. (2003a). Copyright # American Society for Microbiology.)

purified pMMO-enriched membranes, with a type 2 CuII signal of intensity corresponding to one CuII ion per protein molecule superimposed on an almost isotropic signal centered near 2.1 of similar intensity (Chen et al., 2004). Thus, the pMMO hydroxylase is a copper protein only, and as isolated, the copper cofactors consist of a mixture of CuI and CuII ions (Chan et al., 2004; Nguyen et al., 1996). The pMMO purified by ion-exchange chromatography (Method 2) shows the same subunit composition in SDS-PAGE as the protein purified by Method 1, consisting of the three subunits a, b, and g with an apparent molecular mass of 45, 26, and 23 kDa, respectively. The copper content of the pMMO prepared using DEAE-Sepharose procedure is 0.141 mmol/mg of protein, which corresponds to 14.1 Cu atoms per protein, assuming a molecular mass of 100 kDa for the apo-protein. The residual level of Fe in the purified pMMO corresponds to a Cu/Fe of 70–100/1. The CuKa X-ray absorption edge at 8984 eV indicates the same level of reduction of the copper ions in the protein (Nguyen et al., 1996). Low temperature EPR gives the same type 2 CuII feature superimposed on the isotropic CuII signal at g  2.1 as for the protein purified by Method 1.

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4.2. Specific activities of the purified pMMO The specific activities of the pMMO toward oxidation of propylene to propylene oxide (expressed as nmol of propylene oxide/min/mg of protein), measured using either NADH or duorquinol as the reductant, are compared among the detergent-solubilized membranes and the corresponding pMMO-detergent complexes purified by the various protocols described earlier in Table 12.2. These enzymatic activities are comparable to the specific activity of 88.9 nmol of propylene oxide/min/mg of protein obtained for the pMMO-enriched membranes mentioned earlier with NADH as the reductant (Chan and Yu, 2008; Chan et al., 2007). It is well known that specific activities of membrane proteins are strongly dependent on the physiochemical environment in which they are embedded, particularly the microscopic fluidity, and the specific activity of a given membrane protein is also expected to depend on the substrates used to assay the activity, and whether the protein is vectorially oriented in a phospholipid bilayer, solubilized in a phospholipid–detergent mixture, or encapsulated in a protein–detergent complex. Thus, the pMMO derived by both methods of purification of the enzyme from solubilized pMMOenriched membranes is functional, irrespective of whether NADH or duroquinol is used to assay the activity.

Table 12.2 The copper contents and specific activities of the pMMO in various preparations

Preparation

Cu content (atoms/protein) Reductant

pMMO-enriched membranes 12–15 Solubilized membranes 13.3 13.3 pMMO-detergent complex: Method 1d 14.3 14.3 Method 2ae 13.3 Method 2bf 14.1 a b c d e f

Specific activitiesa (nmol/min/mg of protein)

NADHb 88.9 NADHb 10.2 Duroquinolc 93.5 NADHb Duroquinolc NADH NADH

21.5 15.6 5.1 2.6

Conversion of propylene to propylene oxide. The concentration of NADH used is 5 mM. The concentration of duroquinol used is 1 mM. The pMMO is purified by direct transfer into detergent micelles followed by gel filtration using a Sephacryl S-300HR column. The pMMO is purified by ion-exchange chromatography using a L-lysine-agarose column. The pMMO is purified by ion-exchange chromatography using a DEAE-sepharose column.

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ACKNOWLEDGMENTS This work was supported by Academia Sinica and grants from the National Science Council of the Republic of China (NSC 95-2113-M-001-046, 97-2113-M-001-027, 98-2113-M001-026 to S. I. C; and 97-2113-M-001-006-MY3 to S. S.-F. Yu).

REFERENCES Balasubramanian, R., Smith, S. M., Rawat, S., Yatsunyk, L. A., Stemmler, T. L., and Rosenzweig, A. C. (2010). Oxidation of methane by a biological dicopper centre. Nature 465, 115–119. Basu, P., Katterle, B., Andersson, K. K., and Dalton, H. (2003). The membrane-associated form of methane mono-oxygenase from Methylococcus capsulatus (Bath) is a copper/iron protein. Biochem. J. 369, 417–427. Chan, S. I., and Yu, S. S.-F. (2008). Controlled oxidation of hydrocarbons by the membrane-bound methane monooxygenase: The case for a tricopper cluster. Acc. Chem. Res. 41, 969–979. Chan, S. I., Chen, K. H.-C., Yu, S. S.-F., Chen, C.-L., and Kuo, S. S.-J. (2004). Toward delineating the structure and function of the particulate methane monooxygenase from methanotrophic bacteria. Biochemistry 43, 4421–4430. Chan, S. I., Wang, V. C.-C., Lai, J. C.-H., Yu, S. S.-F., Chen, P. P.-Y., Chen, K. H.-C., Chen, C.-L., and Chan, M. K. (2007). Redox potentiometry studies of particulate methane monooxygenase: Support for a trinuclear copper cluster active site. Angew. Chem. Int. Ed. 46, 1992–1994. Chen, K. H.-C., Chen, C.-L., Tseng, C.-F., Yu, S. S.-F., Ke, S.-C., Lee, J.-F., Nguyen, H. T., Elliott, S. J., Alben, J. O., and Chan, S. I. (2004). The copper clusters in the particulate methane monooxygenase (pMMO) from Methylococcus capsulatus (Bath). J. Chin. Chem. Soc. 51, 1081–1098. Chen, P. P.-Y., Yang, R. B.-G., Lee, J. C.-M., and Chan, S. I. (2007). Facile O-atom insertion into C-C and C-H bonds by a trinuclear copper complex designed to harness a singlet oxene. Proc. Natl. Acad. Sci. USA 104, 14570–14575. Choi, D. W., Kunz, R. C., Boyd, E. S., Semrau, J. D., Antholine, W. E., Han, J. I., Zahn, J. A., Boyd, J. M., de la Mora, A. M., and DiSpirito, A. A. (2003). The membraneassociated methane monooxygenase (pMMO) and pMMO-NADH: Quinone oxidoreductase complex from Methylococcus capsulatus bath. J. Bacteriol. 185, 5755–5764. Elliott, S. J., Zhu, M., Tso, L., Nguyen, H. H. T., Yip, J. H. K., and Chan, S. I. (1997). Regio- and stereoselectivity of particulate methane monooxygenase from Methylococcus capsulatus (Bath). J. Am. Chem. Soc. 119, 9949–9955. Feig, A. L., and Lippard, S. J. (1994). Reactions of nonheme iron(II) centers with dioxygen in biology and chemistry. Chem. Rev. 94, 759–805. Kao, W.-C., Chen, Y.-R., Yi, E.-C., Lee, H., Tian, Q., Wu, K.-M., Tsai, S.-F., Yu, S. S.-F., Chen, Y.-J., Aebersold, R., and Chan, S. I. (2004). Quantitative proteomic analysis of metabolic regulation by copper ions in Methylococcus capsulatus (Bath). J. Biol. Chem. 279, 51554–51560. Lieberman, R. L., and Rosenzweig, A. C. (2005). Crystal structure of a membrane-bound metalloenzyme that catalyses the biological oxidation of methane. Nature 434, 177–182. Lieberman, R. L., Shrestha, D. B., Doan, P. E., Hoffman, B. M., Stemmler, T. L., and Rosenzweig, A. C. (2003). Purified particulate methane monooxygenase from Methylococcus capsulatus (Bath) is a dimer with both mononuclear copper and a copper-containing cluster. Proc. Natl. Acad. Sci. USA 100, 3820–3825.

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Lipscomb, J. D. (1994). Biochemistry of the soluble methane monooxygenase. Annu. Rev. Microbiol. 48, 371–399. Ng, K. Y., Tu, L.-C., Wang, Y.-S., Chan, S. I., and Yu, S. S.-F. (2008). Probing the hydrophobic pocket of the active site in the particulate methane monooxygenase (pMMO) from Methylococcus capsulatus (bath) by variable stereoselective alkane hydroxylation and olefin epoxidation. ChemBioChem 9, 1116–1123. Nguyen, H.-H. T., Shiemke, A. K., Jacobs, S. J., Hales, B. J., Lidstrom, M. E., and Chan, S. I. (1994). The Nature of the copper ions in the membranes containing the particulate methane monooxygenase from Methylococcus capsulatus (Bath). J. Biol. Chem. 269, 14995–15005. Nguyen, H.-H. T., Nakagawa, K. H., Hedman, B., Elliott, S. J., Lidstrom, M. E., Hodgson, K. O., and Chan, S. I. (1996). X-ray absorption and EPR studies on the copper ions associated with the particulate methane monooxygenase from Methylococcus capsulatus (Bath). Cu(I) ions and their implications. J. Am. Chem. Soc. 118, 12766–12776. Nguyen, H.-H. T., Elliott, S. J., Yip, J. H.-K., and Chan, S. I. (1998). The particulate methane monooxygenase from Methylococcus capsulatus (Bath) is a novel copper-containing three-subunit enzyme—Isolation and characterization. J. Biol. Chem. 273, 7957–7966. Wilkinson, B., Zhu, M., Priestley, N. D., Nguyen, H.-H. T., Morimoto, H., Williams, P. G., Chan, S. I., and Floss, H. G. (1996). A concerted mechanism for ethane hydroxylation by the particulate methane monooxygenase from Methylococcus capsulatus (Bath). J. Am. Chem. Soc. 118, 921–922. Yu, S. S.-F., Chen, K. H.-C., Tseng, M. Y.-H., Wang, Y.-S., Tseng, C.-F., Chen, Y.-J., Huang, D.-S., and Chan, S. I. (2003a). Production of high-quality particulate methane monooxygenase in high yields from Methylococcus capsulatus (Bath) with a hollow-fiber membrane bioreactor. J. Bacteriol. 185, 5915–5924, 10.1128/JB.185.20.5915-5924.2003. Yu, S. S.-F., Wu, L.-Y., Chen, K. H.-C., Luo, W.-I., Huang, D.-S., and Chan, S. I. (2003b). The stereospecific hydroxylation of [2, 2-2H2]butane and chiral dideuteriobutanes by the particulate methane monooxygenase from Methylococcus capsulatus (Bath). J. Biol. Chem. 278, 40658–40669.

C H A P T E R

T H I R T E E N

Metal Reconstitution of Particulate Methane Monooxygenase and Heterologous Expression of the pmoB Subunit Stephen M. Smith,*,†,1 Ramakrishnan Balasubramanian,*,†,1 and Amy C. Rosenzweig*,† Contents 1. Introduction 2. Preparation of apo pMMO Membranes and Metal Reconstitution 2.1. M. capsulatus (Bath) growth conditions 2.2. pMMO membrane isolation 2.3. pMMO activity assays 2.4. Metal removal with EDTA 2.5. Metal removal with cyanide 2.6. Metal reconstitution 3. Soluble Domain Constructs of the pmoB Subunit 3.1. Design of vectors 3.2. Expression of the soluble domains of pmoB 3.3. Isolation and purification of inclusion bodies 3.4. Refolding of inclusion bodies 3.5. Copper assays 3.6. Methods to assess refolding 3.7. Activity assays of the soluble pmoB domains Acknowledgments References

196 197 197 198 198 199 200 201 203 203 204 204 205 206 207 207 209 209

Abstract Particulate methane monooxygenase (pMMO) is a multisubunit metalloenzyme complex used by methanotrophic bacteria to oxidize methane in the first step of carbon assimilation and energy production. In this chapter, we detail methods * Department of Molecular Biosciences, Northwestern University, Evanston, Illinois, USA Department of Chemistry, Northwestern University, Evanston, Illinois, USA These authors contributed equally to this work.

{ 1

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00013-9

#

2011 Elsevier Inc. All rights reserved.

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to prepare metal free (apo) membrane-bound pMMO and to reconstitute apo pMMO with metal ions. We also describe protocols to clone, express, and refold metal-loaded soluble domain constructs of the pmoB subunit. These approaches were used to address fundamental questions concerning the metal content and location of the pMMO active site.

1. Introduction Particulate methane monooxygenase (pMMO) is a metalloenzyme that catalyzes the conversion of methane to methanol in methanotrophic bacteria (Hakemian and Rosenzweig, 2007). Understanding how pMMO oxidizes methane, the most inert hydrocarbon (C–H bond strength 104 kcal mol
1), could lead to new catalytic processes (Himes and Karlin, 2009). In addition, pMMO has potential applications in bioremediation and greenhouse gas removal (Semrau et al., 2010). However, studies of pMMO have been plagued by controversy surrounding the metal composition and location of the active site (Rosenzweig, 2008). It is widely accepted that there is a functional role for copper in pMMO since all active preparations contain some amount of copper and copper is linked to expression of the enzyme (Hakemian and Rosenzweig, 2007; Lieberman and Rosenzweig, 2004; Semrau et al., 2010). Some preparations of pMMO also contain iron, and it has been proposed that the active site is a dinuclear iron center (Martinho et al., 2007). However, some active preparations of pMMO have been reported to contain no detectable iron (Chan and Yu, 2008; Yu et al., 2003). To determine how many of which metal ions are required for pMMO activity, we developed a method to remove the metal ions from membrane-bound pMMO (Balasubramanian et al., 2010). All enzymatic activity is abolished by metal extraction, although SDS-PAGE analysis indicates that the three subunits, pmoB, pmoA, and pmoC, remain intact. Activity of pMMO, measured by propylene epoxidation or methane oxidation, can then be restored by titrating in 2–3 equivalents of copper per 100 kDa pMMO protomer (composed of one copy each of the three subunits). Addition of iron has no effect on activity (Balasubramanian et al., 2010). These data indicate that copper is the active site metal in pMMO and provide a means to study pMMO without the variable metal content observed with individual isolations. The crystal structures of pMMO from Methylococcus capsulatus (Bath) (Lieberman and Rosenzweig, 2005) and Methylosinus trichosporium OB3b (Hakemian et al., 2008) both contain dicopper centers in the soluble, periplasmic domains of the pmoB subunit. The histidine ligands to this dicopper site are highly conserved. This observation, combined with the

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stoichiometry of the copper-dependent activity data (Balasubramanian et al., 2010), is consistent with the possibility that the dicopper center is the active site. To test this hypothesis, we developed a recombinant system to express and characterize the soluble domains of pmoB (denoted spmoB) (Balasubramanian et al., 2010). This system provides for the first time the opportunity to address fundamental questions about pMMO by sitedirected mutagenesis. Using biochemical assays and spectroscopic techniques, we showed that spmoB assembles metal cofactors similar to those present in wild type, intact pMMO and, most importantly, that spmoB can oxidize propylene and methane. We then pinpointed the dicopper center as the active site of pMMO via analysis of site-directed spmoB variants (Balasubramanian et al., 2010). With these tools in hand, detailed mechanistic studies of pMMO are now possible. Here we detail the methods for generating metal free pMMO (apo pMMO) and reconstituting pMMO with metal ions. We also describe the design and expression of constructs of the pmoB subunit lacking the transmembrane regions (spmoB) and protocols to refold, metal load, and test activity of these spmoB proteins.

2. Preparation of apo pMMO Membranes and Metal Reconstitution 2.1. M. capsulatus (Bath) growth conditions M. capsulatus (Bath) cells are grown using a New Brunswick Scientific BioFlo 4500 Benchtop Fermentor/Bioreactor. 12 l of nitrate mineral salts (NMS) media (2 g/l KNO3, 1 g/l MgSO4 7H2O, 0.1 g/l CaCl2 2H2O) are combined with 6 ml of 20 trace elements solution (500 mg/l Na2EDTA, 200 mg/l FeSO47H2O, 10 mg/l ZnSO47H2O, 3 mg/l MnCl2 4H2O, 30 mg/l H3BO3, 20 mg/l CoCl26H2O, 1 mg/l CuCl2 2H2O, 3 mg/l Na2MoO4 2H2O), 6 ml of 20 sodium molybdate (500 mg/l Na2MoO4 2H2O), 120 ml of 10 phosphate buffer (pH 6.8; 53.4 g/l Na2HPO47H2O, 26 g/l KH2PO4), 6 ml of 0.1 M CuSO4 5H2O, and 9.6 ml of 0.1 M FeEDTA (ethylenediamine–tetraacetic acid ferric–sodium salt). The final Cu and Fe concentrations are 50 and 80 mM, respectively. Methane is bubbled through the media for 30 min prior to inoculation with 5–10 g of cell paste resuspended in sterile NMS. The fermentation is then maintained at 45  C with a 4:1 air:methane ratio and constant agitation of 300 rpm. Cells are harvested at an OD600 of 5–7, centrifuged at 8000g, washed in 25 mM Pipes, pH 6.8, recentrifuged, frozen in liquid nitrogen, and stored at
80  C.

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2.2. pMMO membrane isolation Frozen M. capsulatus (Bath) cells are resuspended and lysed either by sonication or microfluidization. Intracytoplasmic membranes are isolated by low speed centrifugation to remove cell debris, followed by several rounds of ultracentrifugation and homogenization to remove contaminating soluble proteins. 1. Frozen cells are thawed in 200–300 ml of freshly prepared lysis buffer (25 mM PIPES, pH 6.8–7.0, 250 mM NaCl) and transferred to a stainless steel beaker on ice. a. For sonication, the instrument is tuned and a stir bar is added to the resuspended cells. Cells are sonicated, with stirring, on ice at 4  C for 10 min with 1 s on/off pulses at 40% max power. b. For microfluidization, the heat-exchanging coil is submerged in an ice water bath and the instrument is flushed with ice cold lysis buffer. The cells are then passed through the microfluidizer three times at a constant pressure of 180 psi. 2. The lysate is centrifuged at 20,000g for 1.5 h to remove cell debris. 3. The supernatant is ultracentrifuged at 160,000g for 1 h and the pelleted membranes resuspended in fresh lysis buffer and homogenized using a Dounce homogenizer. This process is repeated three times to ensure complete removal of contaminating soluble proteins. 4. The protein concentration of the resulting pMMO-containing membranes is determined using the detergent-compatible Bio-Rad DC protein assay. The membranes are then diluted to 10–20 mg/ml, flash frozen in liquid nitrogen, and stored in a
80  C freezer.

2.3. pMMO activity assays pMMO activity is typically measured using a propylene epoxidation assay or a methane oxidation assay with duroquinol as the reductant. In general, freshly prepared duroquinol is combined with pMMO in a septum sealed vial. The assay is initiated by replacing a volume of the headspace gas with either propylene or methane and incubating in a shaking waterbath at 45  C. The propylene oxide or methanol produced is quantitated by gas chromatography and compared with known standards. 1. Frozen pMMO membranes are thawed on ice. Duroquinol is freshly prepared by following the method of Zahn and DiSpirito (1996), which involves reducing duroquinone with sodium dithionite and sodium borohydride in acidic ethanol.

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2. A small spatulaful of duroquinol (0.9–1.1 M final concentration, in excess for the purpose of the assay) is combined with 50–250 ml of pMMO membranes in a 1.5–3 ml septum sealed vial. 3. 1–2 ml of the headspace gas is replaced with propylene or methane, and the vial is placed in a shaking water bath set at 45  C and 200 rpm for 3 min. 4. The amount of propylene oxide or methanol produced is measured using a Hewlett Packard 5890A gas chromatograph. a. Propylene oxide is detected by injecting 50–250 ml of the headspace gas onto a Porapak Q packed column (Supelco) maintained at a constant temperature of 180  C and quantitated by comparison to a standard curve generated from 99% pure propylene oxide (Sigma Aldrich). b. For methanol, the vial is heated to 85  C for 10 min to stop the reaction and then cooled on ice. The assay mixture is then centrifuged for 2 min, and 3 ml of the clear solution is loaded onto an Rt-Q-BOND capillary column (Restek) maintained at a constant temperature of 75  C. As above, methanol production is quantitated by comparison to a standard curve generated using >99% pure methanol (Sigma Aldrich). M. capsulatus (Bath) cells grown with 50 mM CuSO4 and 80 mM FeEDTA in the media typically produce pMMO that contains between 6–20 Cu and 0.5–2 Fe per 100 kDa pMMO protomer, with average values of 10.2 Cu and 1.31 Fe, as measured by inductively coupled plasma optical emission spectroscopy (ICP-OES) (Fig. 13.1). Using the procedures detailed above, these isolated pMMO membranes exhibit specific activities of 50–200 nmol propylene oxidemin
1 mg pMMO
1 (Balasubramanian et al., 2010).

2.4. Metal removal with EDTA Several groups have used EDTA to remove metal from isolated pMMO membranes (Basu et al., 2003; Takeguchi et al., 1998). Using an EDTAcontaining buffer in a dialysis experiment, we were able to generate inactivated pMMO-containing membranes, but total metal removal was not achieved. ICP-OES analysis before and after EDTA treatment indicates that only  6 Cu and  1 Fe per 100 kDa pMMO protomer are removed using this method (Fig. 13.1). 1. Membrane-bound pMMO samples are digested in 5% trace-metal grade (TMG) nitric acid. The metal content (Cu, Fe, and Zn) is determined using ICP-OES by comparison with standard curves generated from atomic absorption standards diluted in 5% TMG nitric acid.

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Metal content (equivalents per 100 kDa pMMO)

16 14

10.2 ± 3.87

12 10 8 6 4.12 ± 0.15 4 2

1.31 ± 0.62 0.34 ± 0.11

0

As-isolated

EDTA

0.06 ± 0.03

0.35 ± 0.11

CN

Figure 13.1 Metal content of isolated pMMO membranes (seven independent isolations), EDTA treated membranes (three independent isolations) and CN
treated membranes (four independent isolations) expressed per 100 kDa pMMO protomer. The copper content is shown in white and the iron content in gray.

2. Three to 12 ml of pMMO membranes are placed in a 3500 MWCO Slide-A-Lyzer Dialysis Cassette (Thermo Scientific). 3. The cassette is initially dialyzed against 2 l of an EDTA chelating buffer (25 mM PIPES, pH 7, 250 mM NaCl, 250 mM Na2EDTA) with stirring at 4  C for 3 h before exchanging into fresh EDTA chelating buffer and dialyzing overnight. 4. After this overnight dialysis, EDTA is removed by dialyzing against 2 l of lysis buffer (25 mM PIPES, pH 6.8–7.0, 250 mM NaCl), exchanging every 2 h for a total of 12 l. 5. The metal content of these EDTA treated pMMO membranes is again measured by ICP-OES. EDTA treated pMMO membranes contain 4 Cu and  0.3 Fe per 100 kDa (Fig. 13.1). In order to achieve total metal removal, a second procedure involving cyanide was developed.

2.5. Metal removal with cyanide Prior to CN
treatment, the metal content of active pMMO-containing membranes is measured via ICP-OES. The amount of CN
used in the extraction buffer is based on the total copper concentration of the isolated membranes. To limit CN
exposure, the extraction buffer is maintained at

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a pH  8 and all open manipulations that involve CN
are performed inside a hood with the highest level of personal protective equipment (fluidresistant lab coat, nitrile gloves, goggles). The institutional office of research safety must be consulted about proper working and disposal procedures. A 10-fold molar excess of CN
and ascorbic acid, relative to the total copper concentration, is added to the buffer. The highly buffered solution does not change pH upon addition of ascorbate. 1. A membrane-bound pMMO sample (6–12 ml) of predetermined copper concentration (measured by ICP-OES) is ultracentrifuged at 160,000g for 1 h, resuspended, and homogenized in 25–50 ml of extraction buffer (50 mM MOPS, pH 8.0, 250 mM NaCl). 2. Solid KCN and L-ascorbic acid are added at 10-fold molar excess and the solution is covered with parafilm and stirred at room temperature for 30–60 min. 3. The CN
treated membranes are then ultracentrifuged 160,000g for 1 h with the CN
buffer appropriately disposed of per institutional procedures, followed by resuspension and homogenization in 25–50 ml extraction buffer (containing no KCN or ascorbic acid). 4. The pMMO membranes are ultracentrifuged, resuspended, and homogenized an additional three times, resuspending in lysis buffer to remove all traces of CN
. 5. The metal content of these CN
extracted pMMO membranes is then determined by ICP-OES. Following the metal extraction procedure detailed above, the resulting pMMO membranes contain, on average, 0.06 Cu and  0.35 Fe per 100 kDa pMMO (Balasubramanian et al., 2010). These values correspond to a total metal removal of 99% of the Cu and 73% of the Fe. Previously, we reported that purified pMMO contains a heme contaminant on the basis of an optical feature at 410 nm, X-ray absorption spectroscopic (XAS), and electron paramagnetic resonance (EPR) data (Lieberman et al., 2003, 2006). Interestingly, an optical spectrum of solubilized CN
treated pMMO also exhibits an absorption peak at 410 nm. Thus, some portion of the iron ( 0.05 Fe per 100 kDa pMMO or 14% of the total iron) that remains after the extraction procedure is from a contaminating iron-porphyrin.

2.6. Metal reconstitution Metals (Cu and/or Fe) are reconstituted into pMMO stoichiometrically on a mole-to-mole basis assuming a molecular mass of 100 kDa. The exact concentration of the metal stock solutions are determined using ICP-OES. To control for dilution errors, the concentration of the metal stock solution is made such that the volume increase from metal additions is negligible relative to the total volume of the assay (volume of apo pMMO membranes

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used in the activity assay  volume of metal stock solution added to the activity assay). After individual metal equivalents are added, the solution is mixed thoroughly and incubated for at least 30 min. The 30 min incubation time was chosen on the basis of time course experiments indicating that maximal activity is achieved after 30 min incubation with added metals (Fig. 13.2). Following this 30 min incubation, activity is measured as described above. 1. Determine pMMO concentration using the detergent-compatible BioRad DC protein assay. 2. Verify metal stock solution (CuSO45H2O, Fe(NH4)2(SO4)2 6H2O) concentration by ICP-OES. 3. Add appropriate molar equivalents of CuSO4 5H2O or Fe (NH4)2(SO4)26H2O to apo pMMO membranes, mix by pipetting or with a stir bar and incubate for at least 30 min. For consistency, all incubation times should be kept constant. 4. Add duroquinol, mix, and measure activity as detailed above (Section 2.3).

Activity (nmol propylene oxide min–1 mg pMMO–1)

Using this approach, we were able to recover 70% of our original propylene epoxidation activity and 90% of our methane oxidation activity by the addition of 2–3 equivalents of copper. The addition of iron has no effect on activity (Balasubramanian et al., 2010). Copper additions beyond 3

160 140 120 100 80 60 40 20 0 0

20

40

60

80 100 120 Time (min)

140

160

180

Figure 13.2 Reconstituted pMMO activity as a function of time incubated with added metal equivalents. A representative time course with 2 equivalents of added Cu is shown. Maximum activity is achieved after approximately 30 min incubation.

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molar equivalents produced an inhibitory effect. The nature of this inhibition is not well understood, but appears to derive from a hydrogen peroxide-producing side reaction between duroquinol and aqueous copper (Miyaji et al., 2009). This activity loss is reversible, however, and can be remedied with the addition of commercial catalase (Sigma Aldrich) (Balasubramanian et al., 2010).

3. Soluble Domain Constructs of the pmoB Subunit 3.1. Design of vectors Full-length M. capsulatus (Bath) pMMO contains three metal centers: a dicopper center and a mononuclear copper center, both coordinated by residues from the pmoB subunit, and a zinc center located within the membrane (Lieberman and Rosenzweig, 2005). The zinc derives from the crystallization buffer, and the nature of this site in vivo is not clear. Attempts to express full-length pMMO subunits in E. coli have not been successful. Based on the crystal structure and the aforementioned copperdependent activity data, we designed, cloned, and expressed three constructs that span the soluble domain of pmoB (denoted spmoBd1, spmoBd2, and spmoB) as potential truncated versions of pmoB that might contain the active site. The pET21b(þ) (Novagen) vector was chosen for the expression of the three domains. The following rationale and methods were used in the design of the expression constructs. 1. spmoBd1: the N-terminal domain of pmoB spanning residues 33–172. This domain contains all the ligands to the two copper centers. The dicopper center is coordinated by His 33, His 137, and His 139, and the monocopper center is coordinated by His 48 and His 72. A forward primer containing an NdeI site 50 -ggaattccatatgcacggtgagaaatcgcagg-30 and a reverse primer containing an HindIII site 50 -gtgatccaagctttccggtggtgacggggttgcgaa-30 are used for PCR amplification from M. capsulatus (Bath) genomic DNA. Both the PCR products and the vector are digested using NdeI/HindIII enzymes and ligated. 2. spmoBd2: the C-terminal domain of pmoB spanning residues 265–414. This domain does not bind any metal ions in the crystal structure. The forward primer 50 -gagaagcaagcttggaggaggacaggccgccggcaccatgcgtgg-30 and the reverse primer 50 -gagatcccaagcttacatgaacgacgggatcagcgg-30 are used for PCR amplification of the region from M. capsulatus (Bath) genomic DNA. As for generation of the spmoBd1 construct, NdeI/ HindIII enzymes are used to digest both the PCR products and the vector, which are subsequently ligated.

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3. spmoB: a third expression construct that connects the N-terminal (spmoBd1) and the C-terminal (spmoBd2) domains of pmoB via a GKLGGG linker instead of the two transmembrane helices present in native pMMO. For the generation of spmoB, we used spmoBd2 amplified with primers 50 -gagaagcaagcttggaggaggacaggccgccggcaccatgcgtgg-30 and 50 -gagatcccaagcttacatgaacgacgggatcagcgg-30 . Both of these primers contain HindIII restriction sites. HindIII digested spmoBd2 is ligated to NdeI/HindIII digested spmoBd1 DNA. The ligated DNA containing spmoBd1 and spmoBd2 results in the spmoB insert with NdeI and HindIII at the 50 and 30 ends, respectively. The NdeI-spmoBd1spmoBd2-HindIII is then ligated to the NdeI/HindIII digested pET21b(þ) vector. The coding regions of spmoBd1, spmoBd2, and spmoB were verified using DNA sequencing. A silent mutation was identified at position 1076, but did not change the amino acid.

3.2. Expression of the soluble domains of pmoB The expression of inserts in the pET21b(þ) vector is controlled by a T7 promoter. For protein expression, plasmid containing the appropriate insert is transformed into BL21(DE3) or Rosetta (DE3) pLysS strains of E. coli. The expression strains are plated on LB containing 50 mg/ml ampicillin. Colonies of E. coli appear on the plates after an overnight growth at 37  C. 1. An overnight culture is grown using three colonies from the plate. 2. Cells from an overnight culture are used as a starter culture for growth in 1 l of LB supplemented with 50 mg/ml ampicillin. 3. After 3 h of growth at 37  C, when the optical density of the cells reaches an OD600 of 0.6, 0.5 mM IPTG is added and the cells are grown for an additional 4–6 h post induction. 4. The cells are harvested by centrifugation at 5000g for 10 min. 5. The cell pellets are resuspended in a total of 100–200 ml of lysis buffer containing 20 mM Tris–Cl, pH 8.0, 50 mM NaCl, aliquoted into 50 ml falcon tubes, flash frozen in liquid nitrogen, and stored in a
20  C freezer until further processing.

3.3. Isolation and purification of inclusion bodies Both spmoBd1 and spmoB always express as inclusion bodies. Growth experiments performed at 18  C with an incubation time of 1 h produced partially soluble spmoBd2. However, for consistency, all growths are performed at 37  C with an incubation time of at least 4 h. The spmoBd1, spmoBd2, and spmoB proteins are then purified from inclusion bodies.

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1. Frozen cells expressing spmoBd1, spmoBd2, or spmoB are thawed in warm water. Cells are lysed by sonication for 10 min with a 10 s on and 30 s off pulse sequence on ice. The output power is set at 50%. 2. The cell lysate is centrifuged at 3000g for 30 min. This step separates inclusion bodies from the cell debris. The supernatant is discarded and does not contain any overexpressed protein. 3. The usually white inclusion body pellet is resuspended with 50–100 ml of a buffer containing 20 mM Tris–Cl, pH 8.0, 250 mM NaCl, 1% Triton X-100. A Dounce homogenizer is used to completely homogenize the solution. 4. The mixture is centrifuged at 10,000g for 15 min and the supernatant discarded. This wash procedure is repeated three times. The use of Triton X-100 containing wash buffers eliminates most of the contaminating proteins and produces inclusion bodies that are almost homogeneous. 5. As a final step, the inclusion bodies are washed using the same buffer, but without detergent. The yield of pure inclusion bodies from each liter of cell culture is 1–3 g. 6. For solubilization of inclusion bodies in urea, 20 ml of freshly prepared 8 M urea is added per gram of purified inclusion bodies. This ratio of urea to inclusion bodies results in almost complete solubilization. The mixture is completely resuspended using a Dounce homogenizer and left to stir at room temperature for at least 1 h. After approximately 1 h, the mixture turns transparent, indicating solubilization. Since the soluble domains do not contain any cysteines, no reducing agent was added. 7. Urea solubilized inclusion bodies are then centrifuged at 15,000g for 30 min to remove any insoluble material. 8. The supernatant from the urea solubilized inclusion bodies is aliquoted and stored at
80  C for long-term use or kept at 4  C for immediate refolding.

3.4. Refolding of inclusion bodies Urea solubilized inclusion bodies are refolded using a stepwise dialysis procedure against buffers with decreasing urea concentrations. All urea stocks are freshly prepared and are not heated. 1. The protein, at a concentration of  5 mg/ml in 8 M urea, is dialyzed against 7 M urea buffered with 50 mM Tris–Cl, pH 8.0, for 3 h. This process is repeated with decreasing urea concentrations (6, 5, 4, 3, 2, 1, 0.5 M) for at least 3 h in each buffer. 2. After dialysis against a buffer containing 0.5 M urea, the final dialysis is performed against 20–50 mM Tris–Cl, pH 8.0, or 20 mM PIPES, pH 7.0, containing 250 mM NaCl and no urea. The efficiency of refolding is

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estimated to be approximately 0.2% of the total protein. This estimate is based on comparing SDS-PAGE band intensities of the initial and final material. 3. After the final dialysis step, precipitates are removed by centrifugation at 20,000g for 30 min at 4  C. On occasion, the precipitates are removed by pelleting using an ultracentrifuge at 40,000g for 30 min. 4. Attempts at loading copper into refolded proteins resulted in complete protein precipitation. Therefore, a reconstitution procedure was developed in which copper is introduced into the protein in a stepwise fashion during the refolding process. Cu(II) (either CuSO45H2O or CuCl22H2O) is added to the 6 M urea dialysis buffer to a final concentration of 1 mM. A stepwise dialysis procedure using urea buffers without copper, similar to that described in step 1, is used for subsequent refolding. A final dialysis step against a buffer lacking both urea and copper likely eliminates all unbound copper.

3.5. Copper assays The copper content of the refolded proteins can be determined by ICP-OES or using a colorimetric bicinchoninic acid (BCA) method. 1. For analysis by ICP-OES, the total metal content (copper, zinc, and iron) is measured as described in Section 2.4. 2. In the second procedure, BCA is used. Two molecules of BCA exhibit strong absorption features at 360 and 562 nm upon coordination to Cu (I). The copper contents determined from the samples are compared to standard curves generated from dilutions of atomic absorption standards (Sigma). a. Standards of 0–60 mM are prepared from copper atomic absorption standards (Sigma). b. 125 ml of 30% trichloroacetic acid is added to 325 ml of the sample or the standard. This step precipitates the protein and releases all of the bound copper. c. To this mixture, 100 ml of a freshly prepared 1.7 mM ascorbate solution is added. Addition of ascorbate reduces all of the Cu(II) to Cu(I). d. In the detection step, 400 ml of BCA solution (prepared by mixing 6 ml of 1% BCA, 3.6 g NaOH, 15.6 g Hepes acid, and 84 ml water) is added and the mixture incubated at room temperature for 5 min (Brenner and Harris, 1995). e. The mixture is centrifuged at 14,000g for 5 min to remove precipitates and the absorbance measured at 360 and 562 nm. f. Molar ratios of protein bound copper are calculated by dividing the total copper concentrations by the protein concentrations determined

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using the theoretical extinction coefficients at 280 nm. Using this method, spmoBd1, spmoBd2, and spmoB bind 1.59 0.84, 0.24 0.09, and 2.84 0.66 copper ions, respectively (Balasubramanian et al., 2010). We used XAS to assess if the copper reconstituted into both CN
treated apo pMMO and the refolded soluble domain constructs forms a dicopper center. Best fits for the second shell scattering obtained from the extended X-ray absorption fine structure (EXAFS) analysis of copperreconstituted apo pMMO and spmoB suggest that a copper cluster similar to that in native pMMO is present (Balasubramanian et al., 2010; Hakemian et al., 2008; Lieberman et al., 2006).

3.6. Methods to assess refolding Two methods can be used to assess the effectiveness of the refolding and copper reconstitution procedure, circular dichroism, and size exclusion chromatography. 1. Using protein concentrations of 1–2 mM in 20 mM potassium phosphate buffer, pH 7.5, an average of 5 scans are collected at 1 nm resolution at 20  C using a 2 mm path length quartz cuvette. The spectra are similar to those measured for laccase, which has a typical, well-characterized cupredoxin fold (Balasubramanian et al., 2010). 2. A Superdex G75 or Superdex S200 column is equilibrated with degassed buffer containing 20 mM Pipes, pH 7.0, and at least 150 mM NaCl. A volume of 0.1–0.5 ml of protein at a concentration of 1 mg/ml is injected onto the column for analysis. Stokes radii of the samples are estimated by comparison of the elution profile to that of standard proteins with known molecular mass. Under these conditions, the soluble domains of pmoB elute both in the void volume and at volumes that correspond to molecular masses of the protein monomers. There is no indication of trimerization (Balasubramanian et al., 2010).

3.7. Activity assays of the soluble pmoB domains Activity assays are performed as described for pMMO (Section 2.3) with the following modifications. 1. Propylene epoxidation. a. Excess duroquinol is added to 350 ml of sample and mixed thoroughly. b. 2.5 ml of headspace gas is removed and replaced with 2 ml of propylene and 0.5 ml of air.

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c. The mixture is incubated in a shaking water bath at 45  C for at least 1 h prior to sampling the headspace gas for propylene oxide. Reliable detection of propylene oxide formed under these assay conditions for spmoB requires at least 40 min incubation time (Figure 13.3). d. In the literature, propylene oxide formation is always represented in specific activity units of nmol propylene oxidemin
1 mg protein
1. For the soluble pmoB constructs, activity is represented as nmol propylene oxidemin
1 mol protein
1 (Balasubramanian et al., 2010). Moles are used instead of mg to account for the differences in molecular masses between the spmoB proteins and native pMMO facilitating direct comparison. 2. Methane oxidation. a. 2 ml of the headspace gas in the reaction vial is replaced with 2 ml of methane. b. After a 1 hr incubation, the reaction vial is heated at 85  C for 10 min to stop the reaction and cooled on ice. c. The samples are then transferred to an eppendorf tube and centrifuged to remove any protein debris. d. 3 ml of the clear liquid is injected onto the capillary column that is held at a constant temperature of 75  C.

0.20

mM propylene oxide

0.15

0.10

0.05

0.00 20

40

60

80

100 120 140 160 180 200 220 Time (min)

Figure 13.3 Time course of spmoB activity. The activity can be measured reliably after a 40 min reaction incubation. For uniformity, all samples are measured after at least 1 h incubation.

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ACKNOWLEDGMENTS This work was supported by NIH grant GM070473. We thank Liliya Yatsunyk, Megen Culpepper, Swati Rawat, and Timothy Stemmler for assistance at various stages of this project.

REFERENCES Balasubramanian, R., Smith, S. M., Rawat, S., Stemmler, T. L., and Rosenzweig, A. C. (2010). Oxidation of methane by a biological dicopper centre. Nature 465, 115–119. Basu, P., Katterle, B., Andersson, K. K., and Dalton, H. (2003). The membrane-associated form of methane monooxygenase from Methylococcus capsulatus (Bath) is a copper/iron protein. Biochem. J. 369, 417–427. Brenner, A. J., and Harris, E. D. (1995). A quantitative test for copper using bicinchoninic acid. Anal. Biochem. 226, 80–84. Chan, S. I., and Yu, S. S. F. (2008). Controlled oxidation of hydrocarbons by the membrane-bound methane monooxygenase: The case for a tricopper cluster. Acc. Chem. Res. 41, 969–979. Hakemian, A. S., and Rosenzweig, A. C. (2007). The biochemistry of methane oxidation. Annu. Rev. Biochem. 76, 223–241. Hakemian, A. S., Kondapalli, K. C., Telser, J., Hoffman, B. M., Stemmler, T. L., and Rosenzweig, A. C. (2008). The metal centers of particulate methane monooxygenase from Methylosinus trichosporium OB3b. Biochemistry 47, 6793–6801. Himes, R. A., and Karlin, K. D. (2009). Copper-dioxygen complex mediated C-H bond oxygenation: Relevance for particulate methane monooxygenase (pMMO). Curr. Opin. Chem. Biol. 13, 119–131. Lieberman, R. L., and Rosenzweig, A. C. (2004). Biological methane oxidation: Regulation, biochemistry, and active site structure of particulate methane monooxygenase. Crit. Rev. Biochem. Mol. Biol. 39, 147–164. Lieberman, R. L., and Rosenzweig, A. C. (2005). Crystal structure of a membrane-bound metalloenzyme that catalyses the biological oxidation of methane. Nature 434, 177–182. Lieberman, R. L., Shrestha, D. B., Doan, P. E., Hoffman, B. M., Stemmler, T. L., and Rosenzweig, A. C. (2003). Purified particulate methane monooxygenase from Methylococcus capsulatus (Bath) is a dimer with both mononuclear copper and a copper-containing cluster. Proc. Natl. Acad. Sci. USA 100, 3820–3825. Lieberman, R. L., Kondapalli, K. C., Shrestha, D. B., Hakemian, A. S., Smith, S. M., Telser, J., Kuzelka, J., Gupta, R., Borovik, A. S., Lippard, S. J., Hoffman, B. M., Rosenzweig, A. C., and Stemmler, T. L. (2006). Characterization of the particulate methane monooxygenase metal centers in multiple redox states by X-ray absorption spectroscopy. Inorg. Chem. 45, 8372–8381. Martinho, M., Choi, D. W., DiSpirito, A. A., Antholine, W. E., Semrau, J. D., and Mu¨nck, E. (2007). Mo¨ssbauer studies of the membrane-associated methane monooxygenase from Methylococcus capsulatus Bath: Evidence for a diiron center. J. Am. Chem. Soc. 129, 15783–15785. Miyaji, A., Suzuki, M., Baba, T., Kamachi, T., and Okura, I. (2009). Hydrogen peroxide as an effecter on the inactivation of particulate methane monooxygenase under aerobic conditions. J. Mol. Catal. B 57, 211–215. Rosenzweig, A. C. (2008). The metal centres of particulate methane monooxygenase. Biochem. Soc. Trans. 36, 1134–1137.

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Semrau, J. D., Dispirito, A. A., and Yoon, S. (2010). Methanotrophs and copper. FEMS Microbiol. Lett. 34, 496–531. Takeguchi, M., Miyakawa, K., and Okura, I. (1998). Purification and properties of particulate methane monooxygenase from Methylosinus trichosporium OB3b. J. Mol. Catal. A 132, 145–153. Yu, S. S.-F., Chen, K. H.-C., Tseng, M. Y.-H., Wang, Y.-S., Tseng, C.-F., Chen, Y.-J., Huang, D. S., and Chan, S. I. (2003). Production of high-quality particulate methane monooxygenase in high yields from Methylococcus capsulatus (Bath) with a hollow-fiber membrane bioreactor. J. Bacteriol. 185, 5915–5924. Zahn, J. A., and DiSpirito, A. A. (1996). Membrane-associated methane monooxygenase from Methylococcus capsulatus (Bath). J. Bacteriol. 178, 1018–1029.

C H A P T E R

F O U R T E E N

Particulate Methane Monooxygenase from Methylosinus trichosporium OB3b Akimitsu Miyaji Contents 1. Introduction 2. Bacterial Cells Expressing pMMO 2.1. Culture medium 2.2. Culture of bacterial cells 2.3. Activity assay of pMMO in bacterial cells 2.4. Alkanes, alkenes, and their halogenated derivatives that are substrates for pMMO 2.5. Application for methanol production 3. Membrane-Bound Form of pMMO 3.1. Isolation of membrane fractions 3.2. Protein analysis 3.3. Electron donors for pMMO in membrane fractions 4. Purification of Detergent-Solubilized pMMO 4.1. Solubilization to water phase using detergents 4.2. Purification using column chromatography 4.3. Metals in purified pMMO 4.4. Crystallization of pMMO 5. Summary Acknowledgments References

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Abstract Particulate methane monooxygenase (pMMO) catalyzes methane hydroxylation to methanol at ambient temperature and pressure. pMMO from Methylosinus trichosporium OB3b is one of the two pMMOs for which the protein structure was determined by X-ray crystallography. Because purified pMMO is inherently instable in vitro, it is difficult to use for time-consuming analysis. Therefore, Department of Environmental Chemistry and Engineering, Tokyo Institute of Technology, Nagatsuta-cho, Midori-ku, Yokohama, Japan Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00014-0

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2011 Elsevier Inc. All rights reserved.

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investigations using crude enzyme preparations of pMMO are useful in some cases. In this chapter, methods for preparing pMMO from M. trichosporium OB3b to varying degrees of purity, including bacterial cells expressing pMMO, membrane fractions containing pMMO, and highly purified pMMO, are described.

1. Introduction Particulate methane monooxygenase (pMMO), which is found in nearly all methane-oxidizing bacteria (methanotrophs), is a membranebound enzyme catalyzing the hydroxylation of methane to methanol selectively at ambient temperature and pressure. Recently, pMMO from two strains, Methylococcus capsulatus (Bath) and Methylosinus trichosporium OB3b, was successfully crystallized and the structure examined by X-ray crystallography (Hakemian et al., 2008; Lieberman and Rosenzweig, 2005). According to the crystallographic analyses, the overall architecture of pMMO from the two strains is almost identical. pMMO is a a3b3g3 trimer comprising three subunits, (PmoB (a subunit), PmoC (b subunit), and PmoA (g subunit)). The N- and C-terminal subdomains of PmoB are hydrophilic, water-exposed regions, while the other regions of pMMO are predominantly hydrophobic transmembrane regions. pMMO contains copper ions that are required for its enzymatic activity. The pMMO from M. trichosporium OB3b contains two copper-binding sites, a dinuclear and a mononuclear site (Hakemian et al., 2008), whereas a third copper-binding site is found in the pMMO from M. capsulatus (Bath) (Lieberman and Rosenzweig, 2005). The dinuclear copper site is located in a water-soluble N-terminal subdomain of PmoB, where an N-terminal (His40) and two additional histidine residues (His144 and His146) are coordinated to the two copper ions. In contrast, the mononuclear site is found in a hydrophobic region of pMMO, where Asp129, His133, and His146 of PmoC and Glu200 of PmoA are coordinated to the copper ion. Although the amino acid residues in these two copper-binding sites are well conserved among pMMOs from other methanotrophic bacteria, the residues in the third copper-binding site of M. capsulatus (Bath) pMMO are not, suggesting that the two conserved sites are essential for the enzymatic function of pMMO (Hakemian and Rosenzweig, 2007). Recently, the dinuclear copper site has been shown to serve as an active center for the oxidation of methane to methanol (Balasubramanian et al., 2010); however, the role of the mononuclear copper site in the activity of pMMO is unknown. For accelerating studies for deciphering the catalytic mechanism of pMMO and for applying pMMO in industry, a recombinant pMMO

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expression system in suitable heterologous host cells, such as Escherichia coli, is a widely required tool. Although the expression of the pMMO gene in E. coli has not been achieved to date, pMMO from M. trichosporium OB3b can be expressed in Rhodococcus erythropolis LSSE8-1 (Gou et al., 2006). The LSSE8-1 cells expressing pMMO show activity for methane oxidation to methanol, although the amount of methanol produced is low. On the other hand, a soluble region peptide of pMMO from M. capsulatus (Bath) was synthesized in E. coli as an inclusion body, which could be activated in vitro by refolding of the peptide and reconstitution of the copper sites (Balasubramanian et al., 2010). In addition, our preliminary experiments show that the N-terminal soluble subdomain of pMMO from M. trichosporium OB3b where the dinuclear copper site is located can be expressed as a soluble protein. The advance in these techniques is essential to delineate methane hydroxylation by pMMO and to improve the function of pMMO for industrial application. In this chapter, we describe procedures for the preparation of pMMO from M. trichosporium OB3b to varying degrees of purity, such as cells expressing pMMO, membrane fractions containing pMMO, and highlypurified pMMO. These sources of pMMO are essential to study the catalytic mechanisms of pMMO from strain OB3b even once pMMO can be obtained by protein engineering techniques. The basic procedures described here can be applied for establishing the isolation procedures of recombinant pMMO.

2. Bacterial Cells Expressing pMMO The advantages of using whole bacterial cells as a source of pMMO are that the stability of the enzyme and the electron donor regeneration system, which is required to sustain the enzymatic reaction of pMMO, are effectively preserved. Therefore, cells of M. trichosporium OB3b expressing pMMO are typically used for applying pMMO to methanol production (Lee et al., 2004; Mehta et al., 1987; Takeguchi et al., 1997) and the degradation of haloalkanes (Lontoh and Semrau, 1998). M. trichosporium OB3b has two forms of methane monooxygenase (MMO): pMMO and soluble MMO (sMMO). The culture conditions largely affect the production of pMMO by bacterial cells. By adjusting the copper-to-biomass ratio in the culture medium, the expression of the pMMO and sMMO genes can be controlled (Murrell et al., 2000). The batch and continuous culture conditions for producing pMMO from M. trichosporium OB3b have been well established (Shah et al., 1992, 1995). The presence of copper ions and concentrations of nitrate is particularly important for not only the expression of the pMMO gene in strain

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OB3b cells, but also required for the activity of pMMO. Based on these previous studies, the ideal medium composition and culture conditions for the expression and activity of pMMO were decided and are described in the following sections.

2.1. Culture medium For the culturing of M. trichosporium OB3b cells, a 10-fold concentrated nitrate mineral salt (NMS) medium is first prepared (Tables 14.1 and 14.2), which can then be stably stored at room temperature. A 40 mM FeSO4 stock solution is prepared as follows: 0.112 g of FeSO4
7H2O and 0.5 mL of 0.25 M H2SO4 are added to distilled water, and the volume is adjusted to 100 mL. A 10 mM CuSO4 stock solution is prepared by dissolving CuSO4
6H2O in distilled water. The metal content of these two metal aqueous solutions can be accurately determined by atomic absorption spectroscopy (AAS) or inductive-coupled plasma atomic emission spectroscopy (ICP-AES). To prepare the working medium, the 10-fold concentrated NMS medium is appropriately diluted with distilled water, and 10 mM CuSO4 is then added to a final concentration of either 5 or 10 mM. After sterilization Table 14.1 Composition of NMS medium (10-fold concentrated) Component

Amount (per 5 L)

NaNO3 KH2PO4 NaHPO4 12H2O K2SO4 MgSO4 7H2O CaCl2 (0.35 g/mL-distilled water) Trace elements (see Table 14.2) 10 mM H2SO4

42.5 g 26.5 g 108.5 g 8.5 g 1.85 g 1 mL 100 mL 2.5 mL







Table 14.2 Composition of the trace element solution (100 mL) Trace element





ZnSO4 7H2O MnCl2 4H2O H3BO3 NaMoO4 2H2O CoCl2 6H2O KI







Amount (g)

0.287 0.209 0.062 0.048 0.048 0.083

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and cooling of the culture medium, the 40 mM FeSO4 solution is added using a 0.22-mm syringe filter to a final concentration of 80 mM.

2.2. Culture of bacterial cells To produce cells to be used as a source of pMMO, cells are grown in 3 L of NMS medium containing 10 mM CuSO4 and 80 mM FeSO4. To obtain the inoculum for the large-scale batch culture, a two-step small-scale culture is performed. The cells are first cultured in a 25-mL NMS medium containing 5 mM CuSO4 in a 200-mL baffled Erlenmeyer flask. After inoculation of the medium with a thawed 80  C glycerol stock of the bacterial cells, the gas phase of the flask is filled with a 0.22-mm filter-sterilized 1:4 mixture (v/v) of methane and air. The flask is then incubated at 30  C for 3–5 days with shaking at 120 rpm, and every 12 h, a 1:1 (v/v) gas mixture of methane and oxygen is supplied through a 0.22-mm filter. During this culture period, cells are in the logarithmic growth phase, and the optical density of the medium at 600 nm (OD600) reaches 1. At this point, all of the culture medium is transferred to a 200 mL NMS medium supplemented with 10 mM CuSO4 in a 500-mL baffled Erlenmeyer flask to initiate the second culturing step. During this stage, the culture conditions are identical to those in the first stage, and the 500-mL flask is incubated at 30  C for 2 days with shaking at 120 rpm. The cultured medium is used as the large-scale culture inoculum. The prepared inoculum is transferred to a 3 L NMS medium containing 10 mM CuSO4 in a 5-L fermentor. Methane and oxygen (1:1, v/v) are supplied through balloons attached to the head space of the fermentor, and the gas is circulated using a diaphragm pump. Every 12 h, the balloons are filled with methane and oxygen, respectively. After 3–4 days of culture at 30  C with agitation at 150 rpm, the OD600 reaches 1, 2.5 L of growth medium is then removed, and fresh sterile culture medium is added to re-initiate growth. Cells in the removed growth medium are collected by ultracentrifugation at 277,200g for 10 min at 4  C. To wash the cells, the resulting pellet is suspended in a 25 mM MOPS buffer (pH 7.0), followed by a final ultracentrifugation at 277,200g for 10 min at 4  C. The cell pellet is then suspended in 25 mM MOPS (pH 7.0) to be 0.5 g-cell pellet mLbuffer 1 and stored at 80  C frozen liquid nitrogen in small aliquots. Using this method, 2–4 g of wet cells can be obtained from 1 L of growth medium. From 1 g of wet cells, 0.1 g of dry cells can be obtained by lyophilization.

2.3. Activity assay of pMMO in bacterial cells For measuring pMMO activity in bacterial cells, the oxidation of propene to epoxypropene is used. The produced epoxypropene is not a substrate for methanol dehydrogenase in the cells, thus is not oxidized further.

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In addition, solubility of propene into water is higher than that of alkanes that can be oxidized by pMMO (see Section 2.4, Table 14.3). Therefore, propene epoxidation is a convenient reaction for measuring pMMO activity. For regeneration of the electron donor in the cells, sodium formate is used. Formate is oxidized to carbon dioxide by formate dehydrogenase, coupled with NADþ reduction to NADH in the cells. The assay protocol is as follows: A 2.5-mL reaction mixture, consisting of 0.1 g-wet cells weight mL 1 of bacterial cells suspended in a 25 mM MOPS buffer (pH 7.0), and 15 mM sodium formate, is placed in a 10-mL vial and sealed with a Teflon-sealed septum. The reaction mixture is incubated in a 30  C thermostatic water bath for 5 min. The reaction is initiated by the injection of 1 mL of propene into the reaction vial using a gas-tight syringe, and the vial is then incubated in a 30  C thermostatic water bath. The reaction continues almost linearly for 3 h. The amount of produced propene epoxide is determined by flameionized detector-gas chromatography. The activity is described as the amount of epoxypropane produced in 1 min by 1 mg of wet or dry cells weight (mol-epoxypropane min 1 mg-wet or dry cells weight 1).

2.4. Alkanes, alkenes, and their halogenated derivatives that are substrates for pMMO Substrates and products of oxidation by pMMO from strain OB3b were investigated in whole cells, or the cells treated with cyclopropanol to inhibit further oxidation of produced primary alcohol by methanol dehydrogenase. The products from oxidation of alkanes, alkenes, and their halogenated derivatives by pMMO from strain OB3b are summarized in Table 14.3. The substrate range is narrower than that of sMMO (Burrows et al., 1984); thus the substrate binding site of pMMO is more restricted for substrate than that of sMMO. The substrate binding site of pMMO has still not been identified, but probably is the hydrophobic area adjacent to the dinuclear copper site (Lieberman and Rosenzweig, 2005). Stereoselectivity of pMMO from strain OB3b in some reactions was also investigated, as summarized in Table 14.4. The selectivity for alkenes is not as high as for alkene monooxygenase (Weijers et al., 1988).

2.5. Application for methanol production Methanol production from methane at ambient temperature and pressure using strain OB3b-expressing pMMO has been investigated well by several groups. For this process, the further oxidation of methanol by methanol dehydrogenase should be inhibited. In cells, phosphate, cyclopropane, cyclopropanol, and sodium chloride are used as inhibitors for the further oxidation of methanol (Lee et al., 2004; Mehta et al., 1987; Shimoda and

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Table 14.3 Oxidation of alkanes, alkenes, and their halogenated derivatives by Methylosinus trichosporium OB3b expressing pMMO

Substrate

Products

Product distribution (%)

Alkane Methane

Methanola

100

Ethane

Ethanalb

100

Propane

2-Propanol Propanalb 2-Butanol Butanalb 2-Pentanol Pentanalb 3-Pentanol

84 16 91 9 31 69 0

1-Butene

Epoxyethane Epoxypropane Allyl alcohol 1-Propanol 1,2-Epoxybutane

100 95 4.6 0.4 100

1,3-Butadiene

1,2-Epoxybut-3-ene

100

cis-2-Butene

cis-2,3-Epoxybutane

100

trans-2-Butene

trans-2,3Epoxybutane trans-2-Butane-1-alb

41 59

1-Chloro-2-propanol 1-Propanola 1-Chloro-3-propanola 2-Chloro-1-propanol Acetonec 1-Bromo-2-propanol 1-Propanol 1-Bromo-3-propanol 2-Bromo-1-propanol Acetonec

64 29 7 29 71 72 24 4 n.d.d n.d.d

Butane Pentane

Alkene Ethylene Propylene

Haloalkane and haloalkene 1-Chloropropane

2-Chloropropane 1-Bromopropane

2-Bromopropane

Reference

Shimoda and Okura (1991) Burrows et al. (1984) Burrows et al. (1984) Burrows et al. (1984) Burrows et al. (1984)

– Shimoda et al. (1993b) Ono and Okura (1990) Ono and Okura (1990) Ono and Okura (1990) Ono and Okura (1990)

Shimoda et al. (1993a) Shimoda et al. (1993a) Shimoda et al. (1993a) Shimoda et al. (1993a) (continued)

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Table 14.3 (continued)

Substrate

1-Chloropropene

1-Bromopropene

a b c d

Product distribution (%)

Products

1-Chloro-2,3epoxypropane Allyl alcohol 1-Propanol 1-Bromo-2,3epoxypropane Allyl alcohol 1-Propanol

Reference

67 23 10

Shimoda et al. (1993b)

63 31 6

Shimoda et al. (1993b)

Further oxidation of primary alcohol by methanol dehydrogenase is inhibited by cyclopropanol. Alcanal is produced by the oxidation of primary alcohol with methanol dehydrogenase expressing in strain OB3b. Ketone is produced by the oxidation of secondary alcohol with NAD(P)þ-dependent alcohol dehydrogenase in strain OB3b. n.d., not described.

Table 14.4 Stereochemistry of alkene epoxidation by pMMO from M. trichosporium OB3b Selectivity (%) Substrate

Alkene Propene 1-Butene 1,3-Butadiene Halogenated alkane 1-Chloropropane 2-Chloropropane 1-Bromopropane 2-Bromopropane

S

Reference

Epoxypropane 57 1,2-Epoxybutane 36 1,2-Epoxybut-3-ene 36

43 64 64

Ono and Okura (1990) Ono and Okura (1990) Ono and Okura (1990)

1-Chloro-2-propanol 2-Chloro-1-propanol 1-Bromo-2-propanol 2-Bromo-1-propanol

30 74 30 74

Shimoda et al. (1993a) Shimoda et al. (1993a) Shimoda et al. (1993a) Shimoda et al. (1993a)

Chiral product

R

70 26 70 26

Okura, 1991; Takeguchi et al., 1997). Methanol produced from methane inhibits pMMO activity in methanotrophs. For the removal of produced alcohol from the reaction mixture to allow the reaction to proceed for extended periods of time at high rates, immobilization of cells and a semicontinuous system using ultrafiltration membranes were established (Furuto et al., 1999; Mehta et al., 1991).

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3. Membrane-Bound Form of pMMO As pMMO is an integral membrane protein, it is found in membrane fractions after cell disruption and centrifugation. pMMO constitutes the major protein (60% of total proteins estimated by SDS-PAGE analysis) in the membrane fractions of OB3b cells cultured, as described in Section 2. Membrane fractions prepared from strain OB3b cells represent a suitable source of pMMO for certain studies. As the pMMO activity in membrane fractions is relatively stable (80% of pMMO activity in membrane fractions can be retained for 1 day at 4  C under nitrogen atmosphere) compared to that of the purified enzyme, membrane fractions represent the optimum sample when enzyme stability is an issue. For instance, they can be used for some modification of pMMO that takes a long time. In addition, because it appears that membrane fractions contain a functional electron transport pathway, although it is unclear whether the pathway is retained completely, membrane fractions are also optimal for studying electron transfer to pMMO.

3.1. Isolation of membrane fractions Takeguchi et al. (1998a) established a procedure for the preparation of membrane fractions containing pMMO from M. trichosporium OB3b. In an attempt to obtain membrane fractions with higher and more stable pMMO activity, we slightly modified their procedure, as described in Sections 3.1.1 and 3.1.2. Due to these modifications, the specific activities of pMMO in our membrane fraction were 10 and 30 nmol-epoxypropane min 1 mg-protein 1 using duroquinol and NADH, respectively, as electron donors. 3.1.1. Cell disruption For the preparation of membrane fractions, 30 g of frozen cells suspension (prepared as described in Section 2) are first thawed in water at room temperature, followed by centrifugation to collect the cells. The resulting pellet (15 g) is suspended in 45 mL of 25 mM MOPS (pH 7.0) containing 10 mg L 1 DNase, 4 mM MgCl2, 1 mM benzamidine, and 300 mM CuSO4. At this stage, the addition of catalase increases the activity of pMMO in the membrane fractions (Miyaji et al., 2009). To disrupt M. trichosporium OB3b cells, ultrasonication is used. Prior to sonication, the cell suspension is transferred to a 100-mL glass beaker and continually gassed with nitrogen. The sonication of OB3b cells is performed at 80 W for 15 min under nitrogen on ice. It should be stressed that the sonication conditions significantly affect the stability of pMMO activity in the membrane fraction and that the temperature of the suspension should not exceed 4  C.

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3.1.2. Collection of membrane fractions by centrifugation Immediately following sonication, the suspension is centrifuged at 27,720g for 10 min at 4  C to remove unbroken cells. The resulting supernatant is then ultracentrifuged at 143,000g for 90 min at 4  C, and the pellet is collected. The pellet is washed twice with 25 mM MOPS (pH 7.0) containing 0.5 mM KCl using ultracentrifugation at 143,000g for 90 min at 4  C. The salt-washed membrane fractions are suspended in 25 mM MOPS (pH 7.0) and can be frozen in small aliquots at this point using liquid nitrogen. Membrane fractions stored at  80  C retain pMMO activity for at least 1 month. From 15 g of wet cell pellets, membrane fractions containing 100 mg of proteins can be obtained.

3.2. Protein analysis The concentrations of pMMO can be determined by the bicinchoninic acid (BCA) assay. The content of pMMO in proteins in membrane fractions can also be estimated from the densitometric analysis of SDS-PAGE gels stained with Coomassie Brilliant Blue (CBB). For preparing a sample for SDS-PAGE, 2 mg-protein mL 1 of membrane fraction is mixed with 2 SDS-loading buffer (2% (w/v) SDS, 2% (v/v) mercaptoethanol, 0.02% (w/v) bromophenol blue, and 0.5% (w/w) glycerol in 25 mM Tris–HCl (pH 6.8)), followed by incubation at room temperature for 30 min. Metal content of membrane fractions is measured by AAS. For preparing samples for AAS, membrane fractions are dissolved in 1 M NaOH, followed by heating at 95  C for 5 min. Membrane fractions contain 150 nmol mg-protein 1 of copper, 450 nmol mg-protein 1 of iron, and 1.5 nmol mg-protein 1 of zinc. For the calculation of yield of pMMO purification, propene epoxidation is used for the pMMO assay. Due to the presence of methanol dehydrogenase in the crude extract and membrane fractions, the propene epoxidation assay is useful to prevent further oxidation of product. As an electron donor for pMMO, NADH and duroquinol (tetramethyl-p-hydroquinone) are used. Duroquinol has low solubility into water, thus its concentration is limited. The assay protocol is as follows: A 500 mL reaction mixture, consisting of 1 mg-protein mL 1 of the pMMO sample (membrane fraction, solubilized fraction, or purified enzyme) suspended in a 25 mM MOPS buffer (pH 7.0), 1 mg mL 1 of catalase, and reductant such as NADH (5 mM) and duroquinol (1 mM), is placed in a 3-mL vial and sealed with a Teflon-sealed septum. The reaction is initiated by the injection of 0.3 mL of propene into the reaction vial using a gas-tight syringe, and the vial is then incubated in a 30  C thermostatic water bath. For a 3 min reaction, the amount of produced epoxypropane is determined by flame-ionized detector-gas chromatography. The specific

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activity is described as the amount of epoxypropane produced in 1 min by 1 mg of protein (mol-epoxypropane min 1 mg-protein 1).

3.3. Electron donors for pMMO in membrane fractions As described above, duroquinol and NADH are used for pMMO activity assay. Some quinone derivatives such as decyl-plastoquinol, reduced coenzyme Q1, and trimethylquinol can drive pMMO, though its activity is lower than that with duroquinol. Succinate-driven pMMO activity in the membrane fractions is observed, as reported previously (Cornish et al., 1985). Its specific activity is almost the same as NADH-driven pMMO activity. Mitochondria-like electron transfer systems in the bacterial membrane may provide electrons to pMMO in vitro and in the cells.

4. Purification of Detergent-Solubilized pMMO The purification of pMMO is complicated due to the inherent instability of pMMO in vitro. To avoid the loss of pMMO activity, purification is routinely performed at 4  C. The purification method developed by Takeguchi et al. (1998b) and our group, which involves the purification of detergent-solubilized pMMO, is described here.

4.1. Solubilization to water phase using detergents For the solubilization of pMMO from membrane fractions, dodecyl-b,Dmaltoside (DDM) was identified as an optimal detergent (Takeguchi et al., 1998b), similar to that found for M. capsulatus (Bath) pMMO (Drummond et al., 1989). The membrane fractions are first degassed by bubbling with nitrogen for 20 min and are then incubated for 45 min in the presence of 1–2% (w/v) DDM with stirring. After incubation with the detergent, the suspension is centrifuged at 143,000g for 90 min at 4  C, and the supernatant can be directly used as the solubilized pMMO sample. However, despite its effectiveness for solubilizing pMMO, we also found that DDM reversibly inhibits pMMO activity (Miyaji et al., 2002). Thus, exchanging DDM in the solubilized preparation to another detergent that does not affect pMMO activity results in higher yields of active pMMO. Among the detergents tested, we determined that both Brij 58 and Tween 20 did not inhibit pMMO activity. Although these detergents have an ability to prevent protein aggregation, they only have a weak ability to solubilize membrane proteins from cellular membranes due to its high hydrophile–lipophile balance (HLB) value.

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Several methods are used for exchanging detergents (Linke, 2009). One effective method for the removal of excess DDM from solubilized pMMO samples involves the use of detergent–adsorbent beads (Miyaji et al., 2002). In this approach, detergent–adsorbent BioBeads SM-2 (Bio-Rad) are added (1 g-adsorbent per 50 mg-detergent) to the target sample, followed by incubation for 45 min at 4  C with gentle stirring. BioBeads are then removed by centrifugation, and the supernatant is filtrated using a 0.45-mm syringe filter. By this procedure, the pMMO activity can be significantly increased. A second method for exchanging DDM detergent with Brij 58 can be achieved using an anion exchange column, as described in Section 4.2 (Miyaji et al., 2009).

4.2. Purification using column chromatography Following the detergent-solubilization of proteins in membrane fractions, pMMO can be purified using an anion exchange column, such as the POROS 20HQ (Applied Biosystems). In this approach, solubilized proteins are first applied to the POROS 20 HQ column (HR10/10), allowed to adsorb, and are then washed with a buffer containing 0.1% (w/v) Brij 58 to exchange DDM to Brij 58. The adsorbed proteins are then eluted using a concentration gradient of KCl from 0 to 1 M. The flow rate for applying, washing, and eluting proteins is 149 cm h 1. The typical elution profiles of fractions obtained by this method exhibit four peaks, and duroquinoldependent pMMO activity (as described in Section 3) is found in the first peak that is eluted with 150 mM KCl. Purified pMMO displaying an activity of 10 nmol-epoxypropane min 1 mg-protein 1 with duroquinol can be obtained. The purified pMMO shows no activity with NADH, although NADH-dependent MMO activity of membrane fraction is typically observed (30 nmol-epoxypropane min 1 mg-protein 1). The results of the purification procedure for pMMO from M. trichosporium OB3b are summarized in Table 14.5. The purification of pMMO using DDM instead of Brij 58 has also been attempted. Although the elution profile obtained is identical to that observed using Brij 58, pMMO activity is not found in any fraction. Although the recovery of pMMO activity is observed by the removal of DDM using BioBeads, the activity is lower than that of pMMO purified with Brij 58 as a detergent. To evaluate the purity of the purified pMMO enzyme, gel filtration and SDS-PAGE are generally used. Following gel filtration using a Superose 6 column HR10/50, purified pMMO exhibits a single, symmetrical peak. From the SDS-PAGE (10–15% gradient gel) of purified pMMO, three subunits of pMMO with apparent sizes of 40, 24, and 21 kDa are observed. On CBB staining, the 24 kDa band shows much clearer than that observed in the SDS-PAGE analysis of membrane fractions. The quantity and activity

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Table 14.5 Purification of pMMO from M. trichosporium OB3b

Step

Crude extract Membrane fractions Solubilization POROS 20 HQ column a b

Total activity Total (nmolprotein epoxypropane (mg) min 1)

Specific activity (nmolepoxypropane min 1 mg-protein 1)

Yielda Purityb (%) (%)

380 89

4180 890

11 10

100 21

 30  60

55 12

28 144

0.55 12

0.67 3.4

 60  90

Total activity at each step divided by that in the first step. Determined by scanning a CBB-stained SDS-PAGE (10–15% gradient gel).

of purified pMMO enzyme can also be analyzed in the identical manner described for the analysis of membrane fractions described in Section 3.

4.3. Metals in purified pMMO The purified enzyme contains 2–3 coppers and no irons per 100 kDa protomer of pMMO (Miyaji et al., 2009). Some increase in pMMO activity is observed by adding CuSO4, but not by adding FeSO4. All or some of these copper ions in pMMO show a type 2 copper signal (g// ¼ 2.23, jA//j ¼ 18.8 mT, g? ¼ 2.06) and a nine-splitting superhyperfine structure around g ¼ 2 by electron paramagnetic resonance (EPR) measurement. The type 2 copper can be oxidized by hydrogen peroxide, which inhibits pMMO activity reversibly (Miyaji et al., 2009). The reduction of the type 2 copper is observed by duroquinol when catalase is added to pMMO in order to remove hydrogen peroxide generated during isolation or/and by autooxidation of duroquinol (Miyaji et al., 2009). The reduction is not observed by NADH, which is consistent with the observation that purified pMMO does not show NADH-driven activity.

4.4. Crystallization of pMMO As many questions remain concerning the active site and conformation of pMMO, determining the protein structure of this unique protein at high resolution is critical. Recently, the successful crystallization and X-ray structural analysis of pMMO from M. trichosporium OB3b has been accomplished (Hakemian et al., 2008). According to this study, the crystal of pMMO can be grown within 2 weeks using the hanging drop method; however, for obtaining high-quality crystals for use in X-ray analysis, 3–6 months is required.

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5. Summary In summary, purification procedures of pMMO from M. trichosporium OB3b have been improved by the efforts of some researchers. For obtaining good quality of pMMO, it is important to optimize the culture conditions of cells for expressing pMMO, to choose a detergent for stabilizing pMMO in the water phase, and to keep an anaerobic and low temperature (4  C) condition during purification for retaining pMMO activity. Purified pMMO is inherently unstable in vitro and is thus difficult to use for timeconsuming analysis. Therefore, investigations using bacterial cells and membrane fractions are useful in some cases. Alternatively, the technique for obtaining recombinant pMMO will be established in the near future. By this technique, pMMO will be obtained from host cells more simply than from methanotroph cells, and studies for revealing the catalytic mechanism of pMMO and for applying pMMO to chemically difficult reactions will be accelerated. The basic techniques regarding purification of pMMO from methanotrophs such as M. trichosporium OB3b and M. capsulatus (Bath) will also help to handle recombinant pMMO.

ACKNOWLEDGMENTS Our research efforts on pMMO from M. trichosporium OB3b are partially supported by a Grant-in-Aid for Young Scientists (B) (no. 20760527) from the Ministry of Education, Culture, Sports, Science, and Technology.

REFERENCES Balasubramanian, R., Smith, S. M., Rawat, S., Yatsunyk, L. A., Stemmler, T. L., and Rosenzweig, A. C. (2010). Oxidation of methane by a biological dicopper centre. Nature 465, 115–119. Burrows, K. J., Cornish, A., Scott, D., and Higgins, I. J. (1984). Substrate specificities of the soluble and particulate methane monooxygenase of Methylosinus trichosporium OB3b. J. Gen. Microbiol. 130, 3327–3333. Cornish, A., MacDonald, J., Burrows, K. J., King, T. S., Scott, D., and Higgins, I. J. (1985). Succinate as an in vitro electron donor for the particulate methane monooxygenase of Methylosinus trichosporium OB3b. Biotech. Lett. 7, 319–324. Drummond, D., Smith, S., and Dalton, H. (1989). Solubilization of methane monooxygenase from Methylococcus capsulatus (Bath). Eur. J. Biochem. 182, 667–671. Furuto, T., Takeguchi, T., and Okura, I. (1999). Semicontinuous methanol biosynthesis by Methylosinus trichosporium OB3b. J. Mol. Catal. A Chem. 144, 257–261. Gou, Z., Xing, X.-H., Luo, M., Jiang, H., Han, B., Wu, H., Wang, L., and Zhang, Fei (2006). Functional expression of the particulate methane mono-oxygenase gene in recombinant Rhodocuccus erythropolis. FEMS Microbiol. Lett. 263, 136–141.

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Hakemian, A. S., and Rosenzweig, A. C. (2007). The biochemistry of methane oxidation. Annu. Rev. Biochem. 76, 223–241. Hakemian, A. S., Kondapalli, K. C., Telser, J., Hoffman, B. M., Stemmler, T. L., and Rosenzweig, A. C. (2008). The metal centers of particulate methane monooxygenase from Methylosinus trichosporium OB3b. Biochemistry 47, 6793–6801. Lee, S. G., Goo, J. H., Kim, H. G., Oh, J.-I., Kim, Y. M., and Kim, S. W. (2004). Optimization of methanol biosynthesis from methane using Methylosinus trichosporium OB3b. Biotechnol. Lett. 26, 947–950. Lieberman, R. L., and Rosenzweig, R. C. (2005). Crystal structure of a membrane-bound metalloenzyme that catalyses the biological oxidation of methane. Nature 434, 177–182. Linke, D. (2009). Detergent: An overview. Methods Enzymol. 463, 603–617. Lontoh, S., and Semrau, J. D. (1998). Methane and trichloroethylene degradation by Methylosinus trichosporium OB3b expressing particulate methane monooxygenase. Appl. Environ. Microbiol. 64, 1106–1114. Mehta, P. K., Mishra, S., and Ghose, T. K. (1987). Methanol accumulation by resting cells of Methylosinus trichosporium OB3b (I). J. Gen. Appl. Microbiol. 33, 221–229. Mehta, P. K., Ghose, T. K., and Mishra, S. (1991). Methanol biosynthesis by covalently immobilized cells of Methylosinus trichosporium: Batch and continuous studies. Biotechnol. Bioeng. 37, 551–556. Miyaji, A., Kamachi, T., and Okura, I. (2002). Improvement of the purification for retaining the activity of the particulate methane monooxygenase from Methylosinus trichosporium OB3b. Biotech. Lett. 24, 1883–1887. Miyaji, A., Suzuki, M., Baba, T., Kamachi, T., and Okura, I. (2009). Hydrogen peroxide as an effector on the inactivation of particulate methane monooxygenase under aerobic conditions. J. Mol. Catal. B: Enzymatics 57, 211–215. Murrell, J. C., McDonald, I. R., and Gilbert, B. (2000). Regulation of expression of methane monooxygenases by copper ions. Trends Microbiol. 8, 221–225. Ono, M., and Okura, I. (1990). On the reaction mechanism of alkene epoxidation with Methylosinus trichosporium (OB3b). J. Mol. Catal. 61, 113–122. Shah, N. N., Hanna, M. L., Jackson, K. J., and Taylor, R. T. (1992). Batch cultivation of Methylosinus trichosporium OB3b: II. Production of particulate methane monooxygenase. Biotechnol. Bioeng. 40, 151–157. Shah, N. N., Park, S., Talor, R. T., and Droege, M. W. (1995). Cultivation of Methylosinus trichosporium OB3b: III. Production of particulate methane monooxygenase in continuous culture. Biotechnol. Bioeng. 40, 705–712. Shimoda, M., and Okura, I. (1991). Selective inhibition of methanol dehydrogenase from Methylosinus trichosporium OB3b by cyclopropanol. J. Mol. Catal. 64, L23–L25. Shimoda, M., Seki, Y., and Okura, I. (1993a). Oxidation of ally compounds with Methylosinus trichosporium OB3b. J. Mol. Catal. 78, L27–L30. Shimoda, M., Seki, Y., and Okura, I. (1993b). Oxidation of halogenated propanes with Methylosinus trichosporium OB3b. J. Mol. Catal. 83, L5–L10. Takeguchi, M., Furuto, T., Sugimori, D., and Okura, I. (1997). Optimization of methanol biosynthesis by Methylosinus trichosporium OB3b: An approach to improve methanol accumulation. Appl. Biochem. Biotechnol. 68, 143–152. Takeguchi, M., Miyakawa, K., and Okura, I. (1998a). Properties of the membranes containing the particulate methane monooxygenase from Methylosinus trichosporium OB3b. Biometals 11, 229–234. Takeguchi, M., Miyakawa, K., and Okura, I. (1998b). Purification and properties of particulate methane monooxygenase from Methylosinus trichosporium OB3b. J. Mol. Catal. A Chem. 132, 145–153. Weijers, C. A. G. M., van Ginkel, C. G., and de Bont, J. A. M. (1988). Enantiomeric composition of lower epoxyalkanes produced by methane-, alkane, and alkene-utilizing bacteria. Enzyme Microb. Technol. 10, 214–218.

C H A P T E R

F I F T E E N

Production, Isolation, Purification, and Functional Characterization of Methanobactins David W. Graham* and Hyung J. Kim† Contents 1. Introduction: Copper, Siderophores, Chalkophores, and Methanobactin 2. Methanotroph Growth and Optimizing Methanobactin Production 3. Methanobactin Isolation and Purification 3.1. Initial methanobactin capture and concentration 3.2. UV–Vis detection of methanobactin 3.3. Purification of methanobactin 3.4. Mass spectral analysis 3.5. Assessing methanobactins based on toxicity and growth 4. Studying Methanobactins in Pseudonatural Environments 4.1. Methanobactin solubilization studies with copper minerals 4.2. MMO gene expression assays for assessing Cu uptake Acknowledgments References

228 231 233 233 235 236 237 238 239 239 240 243 243

Abstract Aerobic methane-oxidizing bacteria (methanotrophs) have a high conditional need for copper because almost all known species express a copper-containing particulate methane monooxygenase for catalyzing the conversion of methane to methanol. This demands a copper homeostatic system that must both supply and satisfy adequate copper for elevated needs while also shielding the cells from copper toxicity. After considerable effort, it was discovered that some methanotrophs produce small peptidic molecules, called methanobactins, which bind copper, mediate copper transport into the cell, and reduce copper toxicity. Unfortunately, isolating, purifying, and proving the functionality of these molecules has been challenging. In fact, until very recently, only one * School of Civil Engineering and Geosciences, Newcastle University, Newcastle Upon Tyne, United Kingdom Departments of Medicine and Biochemistry, University of Utah Health Sciences Center, Salt Lake City, Utah, USA

{

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00015-2

#

2011 Elsevier Inc. All rights reserved.

227

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complete structure had been reported for methanobactins. As such, there is a desperate need for more studies seeking such molecules. The purpose of this chapter is to describe methods used to isolate and purify the original methanobactin with a published complete structure, which is made by Methylosinus trichosporium OB3b. Methods are also included for assessing the function of such molecules under pseudonatural conditions such as growth on mineral copper sources. Special emphasis is placed on verifying that isolated molecules are “true” methanobactins, because recent work has shown that methanotrophs produce other small molecules that also bind metals in solution.

1. Introduction: Copper, Siderophores, Chalkophores, and Methanobactin Transition metal ions, such as iron, zinc, and copper, are required by almost all microorganisms because they often act as cofactors or are associated with enzymes that catalyze key redox reactions (Hughes and Poole, 1989). Unfortunately, these same metals are not readily bioavailable under many growth conditions and/or are toxic at low levels; therefore, microorganisms have developed an array of strategies for acquiring such metals while protecting themselves against toxic effects. A common strategy for metal acquisition and protection is through the production of specific and nonspecific organic metal-binding agents that mediate metal uptake for growth while also shielding the cell from the deleterious metal-catalyzed Fenton chemistry. For example, low-molecular-weight siderophores scavenge insoluble Fe(III) and mediate iron supply to cells. However, work has shown that siderophore-like molecules are not unique to iron and a parallel class of molecules, called methanobactins (Kim et al., 2004, 2005), also exist for copper, especially associated with aerobic methane-oxidizing bacteria (methanotrophs). Copper supply is key to these environmentally important organisms, because it is central to the structure and function of particulate methane monooxygenase (pMMO; Balasubramanian et al., 2010b), the most efficient MMO at oxidizing methane to methanol that is found in almost all known methanotrophs (Hanson and Hanson, 1996). Such metal-specific binding agents are usually produced and regulated by ambient available levels of their target metal. In the case of siderophores, the molecules are often produced in response to low available iron levels (Neilands, 1982). However, what regulates the production of methanobactins is less clear, although it is known that they strongly bind copper, promote copper internalization into the cells, protect cells from free copper toxicity, display redox activity and weak antibiotic properties, and are possibly associated with the pMMO (Balasubramanian and Rosenzweig,

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229

2008; Choi et al., 2008; DiSpirito et al., 1998; Fitch et al., 1993; Kim et al., 2004, 2005; Zahn and DiSpirito, 1996). Thus, methanobactins are analogs for copper as siderophores are for iron and we subsequently called them “chalkophores” (Greek for “copper handling”) for this reason (Kim et al., 2005). Unfortunately, the breadth of different chalkophores in biological systems is less well known, although recent work has shown that alternate methanobactins exist (El Ghazouani et al., 2011; Krentz et al., 2010). However, there is still a strong need for improved isolation and identification methods to improve our understanding of this group of molecules. This is especially true, given that methanotrophs produce other small molecules (Balasubramanian et al., 2010a) that may differ from “classic” methanobactins, which resemble peptidic siderophores (Behling et al., 2008). The purpose of this chapter is to describe methods for identifying, isolating, purifying, and verifying the functionality of methanobactins. General approaches will be presented first; however, focus will be placed here on methods for identifying and assessing function in methanobactins with previously purified forms (Behling et al., 2008; Kim et al., 2004, 2005). Specifically, methods for assessing the ability of methanobactins to solubilize and sequester copper from mineral sources will be provided, how methanobactins influence MMO gene expression (Knapp et al., 2007), and two additional assays for assessing other functional roles will be described. Other key issues addressed here include conditions for growing methanotrophs to produce adequate methanobactin for study and also methods for isolating and purifying these molecules. As a background, there is some confusion over what is and what is not a methanobactin, which has resulted from changes in what “methanobactin” has been called over time and also inadequate verification of isolated molecules’ functional roles. Given this confusion, we have chosen to focus solely on the methanobactin for which there is a complete published structure, which is produced by Methylosinus trichosporium OB3b (Kim et al., 2004). Figure 15.1 shows this methanobactin, which has a chemical formula of C45H56N10O16S5Cu
, an exact mass of 1215.1781, and the following structure: 1-(N-[mercapto-{5-oxo-2-(3-methylbutanoyl)oxazol-(Z)-4-ylidene} methyl]-Gly1-L-Ser2-L-Cys3-L-Tyr4)-pyrrolidin-2-yl-(mercapto-[5-oxo-oxazol-(Z)-4-ylidene]methyl)-L-Ser5-L-Cys6-L-Met7 (Behling et al., 2008). It should be noted, however, that different methanobactins are produced by other species (Knapp et al., 2007; Krentz et al., 2010), and our recommended approaches should be adapted as needed based on the molecule and organism of interest. In summary, this chapter has three parts: (1) methanotroph growth and methanobactin production; (2) methanobactin isolation and purification; and (3) methanobactin in the real world, including mineral–methanobactin interactions and assessing methanotroph growth and activity on mineral copper sources.

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A

NH(1) - OH - Om O Ala O NH CH C CH2 C Gly NH CH2 CH CH2 OH O O CH2 O NH C C CH2 CH C NH1 O CH2 C NH H HN N Ser CH N+ C CH2 HC CH OH HO O¢ Ser 2 CH2 O O¢ +3 NH C O Fe O¢ C N O¢ O¢ O C NH C O O¢ O HC CH Threo - β - OH - Asp CH OH C CH HN O NH Thr CH C CH 2 O

HOCH2 Ser OH

B

NH2

CH2

CH2

CH

NH

NH CH2 CH

CO CH2

CO

CH2

O NH

O

CH

NH

HC

CO

C

CH2

b - OH - Asp

Fe

CH

CH2

NH CO

O

O

O

NH HO

OH

CH

CO

HC NH C

N

CH OH CH

CO

CH2

NH2

CH3 NH CO

N

+3

+

NH NH

O

R

O

NH CO NH CO CH

C

N

O Nd - OH - Om

CH HO CH3 O

C

O

O

C

C

HO

CH3 CH2

C N NH – + CH2 Gly1 S Cu O – S C O N N 4 C NH Tyr C O CH2 NH Ser2 CH O CH2 CH NH CH2 CH Ser5 O O C C C O NH

CH Cys3 CH2

OH S

S

Cys5 CH CH2

CH

OH

NH

O

C

CH3

Met7 NH CH

O C

CH

CH2 CH2 S CH3

Figure 15.1 Structures of (A) Pyoverdin complexed with Fe3þ from Pseudonmonas putida (from DeMange et al., 1990), (B) Azotobactin d from Azotobacter vinelandii complexed with Fe3þ (from DeMange et al., 1988), and (C) Methanobactin from Methylosinus trichosporium (OB3b) (from Behling et al., 2008).

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Isolation and Characterisation of Methanobactins

2. Methanotroph Growth and Optimizing Methanobactin Production Methane-oxidizing bacteria cultures are typified by slow growth rates and variable cell yields under laboratory conditions. Relative to growing M. trichosporium OB3b, we favor the nitrate mineral salts (NMS) medium (see Table 15.1) under conditions suggested by Fox et al. (1990), which are common conditions for type II methanotrophic strains. However, when growing cells for the purpose of producing methanobactin, modifications are often needed. As a background, methanobactin can be found both in the bulk media and within cell membranes in methanotroph cultures (Zahn and DiSpirito, 1996). Unfortunately, successful retrieval of functional Table 15.1 Nitrate mineral salts (NMS) media recipe (Fox et al., 1990)

Components for stock solutions are as follows: Carbon source Instrument-grade methane Nitrate salts solution (100) 10.0 mM NaNO3 1.0 mM K2SO4 0.15 mM MgSO4 47.6 mM CaCl2 2H2O Phosphate buffer solution (pH 7) 100 3.9 mM KH2PO4 6.0 mM Na2HPO4 Adjust pH to 7 with H2SO4 Metals solution 500 2.0 mM ZnSO4 7H2O 1.6 mM MnSO4 7H2O 6.0 mM H3BO3 0.4 mM NaMoO4 6H2O 0.4 mM CoCl2 6H2O 1.0 mM KI Iron solution (1000) 40 mM FeSO4 7H2O Amounts from stock for media solution are as follows: Nitrate salts 10 mL/L Phosphate buffer (pH 7) 10 mL/L Metals 2 mL/L Iron solution 1 mL/L



 







85 g/L 17 g/L 3.7 g/L 0.7 g/L 53 g/L 86 g/L

0.287 g/L 0.223 g/L 0.062 g/L 0.048 g/L 0.048 g/L 0.083 g/L 11.2 g/L

Stock solutions should be prepared using deionized water, preferably with 18 MO resistance. The iron stock solution is sterile-filtered and added to the autoclaved medium to avoid precipitation.

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methanobactin from membranes has not been achieved; therefore, one must rely on harvesting methanobactin from the spent medium, although relationships between rates of methanobactin excretion and accumulation in spent media relative to growth conditions have not been completely understood. There are two general strategies for obtaining methanobactin from M. trichosporium OB3b. The first strategy is to grow the cells according to Fox et al. (1990) with no copper amendment. Care is needed in ensuring that copper contamination does not occur from reagents; therefore, only high-purity reagents are used along with 18 MΩ deionized water. Note that the growth rate of M. trichosporium under copper limitation is lower than under copper replete conditions, because this organism alternately expresses an iron-containing soluble methane monooxygenase (sMMO) that has lower catalytic activities than the pMMO (Murrell et al., 2000). As such, cultures grown under copper-free conditions must be maintained for relatively long durations to generate sufficient methanobactin for use, although such conditions are more likely to produce apo-methanobactin (copper-free) that requires less subsequent processing for copper binding and other characterization assays. Methanobactin yields are typically <5 mg/L using this approach from a culture with an optical density at 600 nm of 1.0 (OD600). Alternately, higher methanobactin yields can sometimes be obtained if one grows the cells at moderate copper levels, typically 2.0–5.0 mM Cu as CuCl2 or CuSO4, and harvesting the cells at higher densities. Empirical studies have shown that if one grows cells in the presence of copper, the cells initially express the copper-requiring pMMO and minimal methanobactin accumulates in solution, but as cell densities increase, copper is titrated out of the bulk media and the cells switch over to the sMMO (Murrell et al., 2000). During and after “switchover,” extraneous methanobactin accumulates in the spent media apparently because cellular copper needs decline when pMMO manufacture also declines. Depending upon cell density, copper level, and the specific culture, methanobactin yields can be higher using this method (up to 10 mg/L per 1.0 OD600 in our lab). It must be emphasized that optimal growth conditions for methanobactin production have not been fully understood, and we recommend that presumptive methanobactin levels be monitored regularly in the spent medium in any growing culture. Methanobactin levels can be estimated by collecting 2–3 mL of the culture, centrifuging cells from solution, and performing a UV–Vis spectrophotometric scan between 280 and 500 nm of the centrate (see Section 3.2). A Plot of peak absorbance at 342 nm over culturing time for M. trichosporium OB3b provides a crude estimate of methanobactin accumulation and aids in choosing an appropriate time for harvesting. The preferred OD600 for actual harvesting varies among research groups that study methanobactins (Balasubramanian et al., 2010b; Choi et al., 2008; Kim et al., 2005). However, our group typically does not

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grow cultures beyond an OD600 of 1.0, because spent media accumulates many nonmethanobactin small molecules that can confound and confuse isolation of the intended target. Recent work has hinted that small molecule yields in methanotroph cultures may also be influenced by iron conditions (Balasubramanian et al., 2010a). In reality, the possibility of iron effects is not surprising, given the close resemblance of the known methanobactin to pyoverdines and azotobactins, both peptidic siderophores. Pyoverdines are small yellow-green fluorescent compounds produced under iron limitation by the fluorescent subgroup of Pseudomonas (Atkinson et al., 1998; Demange et al, 1990; Teintze et al., 1981; Visca et al., 2007), whereas azotobactins are produced by Azotobacter vinelandii as a consequence of high cellular iron requirements associated with N2-fixation (Demange et al., 1988; Knosp et al., 1984). Figure 15.1 shows the similarity between selected siderophores and the methanobactin from M. trichosporium OB3b. Specifically, pyoverdines and azotobactins are chromopeptides like methanobactin and have a peptide chain of between 6 and 10 amino acids bound to a chromophore. As such, it is strongly recommended that approaches for the study of pyoverdines, azobactins, and related molecules also be considered when assessing suspected methanobactin-producing systems (Payne, 1994; Vraspir and Butler, 2009).

3. Methanobactin Isolation and Purification Growth conditions are critical for maximizing methanobactin yield, stability, and quality. We favor using growth media free of copper for methanobactin production that is harvested at comparatively low cell densities. This minimizes the accumulation of potentially confounding extraneous small molecules. In the following section, we describe the methods used to isolate and purify the methanobactin from M. trichosporium OB3b that was used to obtain its 3D structure via mass spectral and crystallographic methods (Kim et al., 2005). Alternate approaches that are also useful for characterizing methanobactins are provided elsewhere (El Ghazouani et al., 2011; Krentz et al., 2010).

3.1. Initial methanobactin capture and concentration Original isolation and purification studies on the methanobactin from M. trichosporium OB3b were performed using a BIOFLO 2.5-L bioreactor (New Brunswick) operated in batch mode with a 2-L culture volume and copper-free NMS growth media. Larger reactors have since been employed, which is highly advantageous simply because methanobactin yields can be low, and it is often necessary to grow multiple batches of

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cells to produce adequate methanobactin for subsequent preparations. Instrument-grade methane supplied at  25 mL/min is used as the sole carbon and energy source for the above reactor system. Care must be taken to exhaust the methane from the reactor to a ventilation hood to prevent methane buildup within the lab. Air is supplied and regulated to maintain soluble O2 levels 30% saturation in air, which was found optimal relative to growth rates (Kim and Graham, 2001). After inoculation from shake flask “overnights,” cultures are grown in the batch mode for 3–6 days, depending upon the initial inoculum size and density, and ideally harvested during the mid-exponential growth phase to yield optimal quality methanobactin (0.6 < OD600 < 0.8). If small bioreactors are used, about 90% of the culture is harvested and replaced with the same volume of fresh NMS medium, and the culture is regrown to an OD600 again between 0.6 and 0.8 for harvesting. This sequence is repeated until 8 L of the spent supernatant is collected, which is usually enough for further work depending on yields. This sequential harvesting process should only be performed for two or three cycles before using a fresh overnight, because as time proceeds, methanobactin yields often decline and extracellular “detritus” starts to accumulate. Collected bulk media is combined and centrifuged at 9000g for 15 min. The decanted centrate should be yellow-green, which is suggestive of copper-free methanobactin. To stabilize methanobactin and to decrease the formation of breakdown products, 1–10 mM CuCl2 is added at this stage to bind copper to methanobactin. However, this step is only performed if the methanobactin is for structural characterization studies. This copper-supplemented medium is stirred slowly for approximately an hour at room temperature or overnight at 4  C in the dark. Upon addition of this copper, methanobactin turns red, although in its dilute state in the spent media, the red tint still appears as a deep yellow. This solution is centrifuged again to remove precipitated copper, and this centrate or the copper-free version (see above) is vacuum-filtered through a 0.45-mm membrane filter (Gelman) to remove any residual suspended solids. At this stage, the spent medium can be stored at 4  C away from light, although it is preferred to concentrate crude methanobactin immediately after harvest using solid-phase extraction on a hydrophobic support. Reversed-phase C18 solid-phase extraction (SPE) Sep-Pak cartridges (Waters Associates) have proved suitable for this purpose. A Masterflex pump is used to load the spent medium and, as extraction proceeds, a brown band slowly appears at the front end of the cartridge. The bound material is washed extensively using deionized water and ultimate elution is achieved using 60% ACN. The eluate is immediately lyophilized for extended storage. This isolation procedure using Sep-Pak cartridges provides preparations that are relatively free of contaminants, and this “crude extract” can be used for some

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Isolation and Characterisation of Methanobactins

practical tests on methanobactin function (with appropriate controls). However, further purification is required for structural and other determinations.

3.2. UV–Vis detection of methanobactin Methanobactin from M. trichosporium OB3b has a very characteristic UV–Vis spectrum that is quite distinct from other copper-binding chromophoric ligands and proteins. As such, the spectrum can be used to approximate its accumulation in the growth medium and also to track the molecule during isolation and purification. Furthermore, apo-methanobactin and Cu–methanobactin each have distinct spectra, making the method helpful for detecting and distinguishing between methanobactin and other small molecules through a simple Cu(II) titration under air atmosphere. Specifically, methanobactin possesses three chromophoric moieties: two mercaptooxazolones that coordinate and reduce copper, and a tyrosine residue in the peptide backbone. The presence of the disulfide bridge (cystine) also contributes to the UV properties of methanobactin in the 255 nm range, albeit weakly. Figure 15.2 shows typical titration spectra of methanobactin from M. trichosporium OB3b with increasing concentrations of Cu(II). Copper-free 342 nm 258 nm

0.6

+Cu(ll)

Absorbance

331 nm

389 nm

0.4

+Cu(ll)

0.2

0.0 300

400

500

600

l (nm)

Figure 15.2 UV–Vis spectra of methanobactin from M. trichosporium OB3b showing successive spectral changes associated with sequential 0.25 equivalent Cu(II) as CuCl2 additions to the as-isolated form. Black, as isolated; red, 0.25 equiv. Cu; green, 0.5 equiv.; blue,  0.75 equiv.; purple, 1 equiv. Prominent as isolated features are at 342 and 389 nm. Upon Cu addition, a new feature appears at 331 with a loss of the 342 nm peak. The 388-nm peak is also lost. This response is characteristic of this methanobactin. Equivalent patterns with different spectra could be used to identify other methanobactins. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this chapter.)

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methanobactin possesses three absorption maxima at 388, 342, and 281 nm, indicative of the three chromophoric functions. Upon Cu(II) binding and autoreduction of the metal, a new peak at 331 nm is observed and a decrease in the 388 nm peak. The weak feature at 258 nm likely indicates cystine formation on Cu binding; the peak at 331 nm is observed when methanobactin is also fully loaded with Cu.

3.3. Purification of methanobactin The initial capture and isolation step using reversed-phase C18 sorbents results in methanobactin of reasonable purity and yield. This product has been successfully used for activity experiments in geochemical and other applied settings assessing methanobactin function and activity in complex matrices (Knapp et al., 2007). However, for analytical or structural characterization studies, methanobactin must be further purified using HPLC C18 chromatography or other methods that afford material of suitable quality for crystal growth. Prior to Kim et al. (2004, 2005), methanobactin had been purified using sequential Sep-Pak extraction and low-pressure, reversed-phase chromatography on a preparative column (15 RPC media by Pharmacia; DiSpirito et al., 1998). However, structural elucidation was not successful due to the presence of breakdown products in samples. Subsequent reinvestigation of the methods showed that low pH conditions induce the loss of the metal ion from the molecule that leads to its instability. For this reason, near neutral buffers are now used for the HPLC chromatography with ammonium or sodium acetate or sodium phosphate in the mobile phase. Reconstitution with Cu as mentioned prior to isolation greatly aids in preventing breakdown products. The procedure for HPLC purification is as follows. Lyophilized extract is dissolved in reagent-grade MQ water and purified using semipreparative Vydac 218 TP1010 column (10250 mm, 300 A˚) (or a similar semipreparative column). The mobile phase is composed of 10 mM ammonium or sodium acetate, pH 6.5 (solution A) and acetonitrile-10 mM ammonium or sodium acetate (80:20 mix, v/v, respectively) (solution B). Linear acetonitrile gradients should suffice and, in our work, methanobactin elutes at 30% B. We use a 20–40 % linear change over 20 min, then to 100% solution B in 2 min to regenerate the column. Absorbance is monitored at methanobactin’s prominent chromophoric peaks: that is, 388 and 342 nm, and major yellow fractions are collected using a fraction collector, lyophilized, and analyzed using Matrix-Assisted Laser Desorption/Ionization Time of Flight (MALDI-TOF; see below). The purified product can be dried in a speed vac or lyophilized; the latter is recommended for long-term storage. Fractions that contain the mass of interest are rechromatographed ˚ ) Vydac 218TP52 (C18) reversed-phase analytical on a (2.1250 mm, 300 A HPLC column using a 30–40% acetonitrile gradient over 20 min at 0.5 mL/ min. The resulting purification should result in a single peak.

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Isolation and Characterisation of Methanobactins

3.4. Mass spectral analysis Both purity and homogeneity of methanobactin is evaluated using mass spectral analysis, which should be performed during and after purification. MALDI-TOF is a convenient mass analysis technique and capable of detecting noncovalent transition metal complexes. Electrospray ionization (ESI) also has been used successfully for mass analysis of methanobactin. However, ESI generates multiply charged species; therefore, it is better suited to studies on mainly purified species. MALDI-TOF principally was used during the purification of methanobactin for crystallization and will be discussed here. Figure 15.3 provides an example negative-ion MALDI-TOF spectrum for methanobactin from M. trichosporium OB3b (Kim et al., 2005), which reveals two predominant ions differing in mass by 62. The peak with (M
H)
at m/z 1153 is assigned the molecular ion for the deprotonated methanobactin, while the most intense peak at m/z 1215 is the corresponding copper complex which can be attributed to the most abundant isotope of copper minus two protons [M
2H þ 63Cuþ 1]
. Additionally, this signal shows an isotopic distribution characteristic of copper and is identical when compared with the simulated isotopic distribution of copper-complexed with a peptide. Mass spectra are recorded on a PerSeptive Voyager-DE STR mass spectrometer (Applied Biosystems) with a reflector TOF mass analyzer. Analyses can be performed in linear, negative mode with time delayed extraction (150 ns) when sensitivity is an issue; however, reflector modes for improved resolution in both positive and negative polarities are 1215.46 1217.46

100

% Intensity

90 80

1216.42

70

1218.46

60

(M-H)

–

M-2H+Cu(l) 1153.46

50 40

1154.43

30

1220.40 1210.53

1155.38

1211.37

20

1119.42 1084.29

10 0 1021

1129.25

1159.38 1175.45 1189.53 1160.53

1231.49 1226.49

0

1075

1129

1183

1237

1291

Mass (m/z)

Figure 15.3 Negative-ion MALDI-TOF spectrum of methanobactin from M. trichosporium OB3b showing both the copper-complexed and the as-isolated (apo) forms (adapted from Kim et al., 2005).

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preferred. Acquisition mass range should be between 500 and 1500 Da. Alpha-cyano-4-hydroxy-cinnamic acid (CHCA), is the standard matrix for peptides in the mass range of methanobactin, but is strongly acidic (pH < 2). This strongly acidic condition disturbs the metal complex as evidence by a fairly substantial apo peak in the mass spectrum. This is especially pertinent to methanobactin due to its instability in acidic solutions indicated by HPLC purification experiments using running buffers with 0.1% trifluoracetic acid as ion pairing agents. Therefore, p-nitroaniline (p-NA) matrix solution (pH 6.5) is used as the matrix (Salih et al., 1998), which greatly improves ionization of methanobactin in positive and negative reflector modes (Kim et al., 2005). Furthermore, the isotopic clusters are consistently fully resolved. This is critical in examining the presence or absence of the metal complex in methanobactin and any other small molecules suspected to bind metals.

3.5. Assessing methanobactins based on toxicity and growth Methanotrophs produce an array of small extracellular molecules and it is necessary to verify that purified molecules are actually methanobactin or methanobactin-like. In fact, confusion over methanobactin and its role is partially due to incomplete structural and functional characterization of isolated molecules. The fact that a molecule binds copper does not necessarily mean it is a methanobactin because copper also binds to some siderophores and other inorganic and organic peptidic structures. Therefore, it is important to verify more than one of the identified functional roles associated with methanobactin, such as copper toxicity suppression, Cu-binding traits themselves (Section 3.2), and mediation of MMO expression in the presence of Cu-minerals or relatively unbioavailable copper sources (Section 4). A useful first assay for assessing methanobactin function is copper toxicity suppression. Although suppressing copper toxicity is not proof of methanobactin (other nonspecific copper-binding agents might also do this), it is useful for validating a key methanobactin function, especially if copper affinity coefficients also are determined. Toxicity suppression can be tested by growing 0.2–0.3 L of cells in copper-free NSM media to an OD600 of about 0.05–0.08 (usually in septum-capped, acid-washed 1-L flasks), verifying sMMO activity prior to use via the naphthalene–naphthol colorimetric assay (DiSpirito et al., 1998). Once significant sMMO activity is detected, one subdivides the culture into 20–40 mL aliquots and transfers them to a series of 125-mL septum-capped serum vials containing CuSO4 or CuCl2 provided at levels designed to attain a final solution concentration of 10 uM Cu. One then adds different quantities of methanobactin to each vial at differing Cu–mb ratios, ranging from no methanobactin to a 2:1 mb: Cu molar ratio. One then monitors OD600 over time, possibly for up to 7 days, and compares growth lags as a function of mb:Cu ratio.

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Methanobactin is implied if the Cu-induced growth lag is progressively suppressed as methanobactin level is increased.

4. Studying Methanobactins in Pseudonatural Environments Methanotrophic bacteria reside in natural aquatic, marine, and terrestrial settings. Although laboratory-based biochemical studies are essential for understanding physical and chemical characteristics of methanobactins, idealized laboratory experimental systems are very atypical of natural methanotroph habitat. This is especially true of the forms of copper previously used in laboratory studies relative to what is found in nature. In reality, we suspect that one of the primary roles of methanobactin in nature is its natural capacity to acquire copper from insoluble sources, such as minerals while simultaneously reducing copper toxicity during transport (Knapp et al., 2007; Kulczycki et al., 2007, 2011). As such, assessing the effect of methanobactin on cell activity in the presence of insoluble copper sources, such as minerals, is likely a key trait of methanobactins, and can be assessed using copper solubilization and gene expression assays. This section summarizes two methods for assessing methanobactin–copper mineral interactions, including assessing relative copper dissolution rates from minerals, and the expression of MMO and related genes during growth in the presence of relatively insoluble copper sources.

4.1. Methanobactin solubilization studies with copper minerals A common characteristic of methanobactin is its ability to bind and solubilize metals, including Cu(I) and Cu(II) (Choi et al., 2006; Hakemian et al., 2005). Fortunately, performing such assays is relatively straightforward, although care is needed in ensuring that background copper levels are minimized, both in the methanobactin itself and associated glassware. This can be done by using appropriate controls and high-sensitivity methods for copper detection. Solubilization assays are performed in the same manner regardless of copper source; however, the tendency of methanobactin to release copper upon exposure varies dramatically among minerals. Example minerals that provide contrasting copper release patterns include silicates, sulfides, oxides, and carbonates, with silicates and sulfides being more refractory to methanobactin-mediated copper release, oxides intermediate, and copper carbonates releasing copper quite readily (Chi Fru et al., in revision). If commercial copper minerals are available (e.g., Sigma-Aldrich), analytical grade minerals are recommended, whereas some minerals are better

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synthesized in-house, such as copper-silicates, due to greater control of mineral copper content (Knapp et al., 2007; Kulczycki et al., 2007, 2011). For a given assay, methanobactin level, and copper content and mineral type must be clearly defined. Typically, such assays do not require pure methanobactin, although knowing the amount of methanobactin in the crude extract is important for establishing appropriate stoichiometric ratios between methanobactin and copper for the assay. However, because such minerals are solids, one cannot always know what exactly the exposed copper level is, and our bias is to provide methanobactin in stoichiometric excess. The following procedure has been used successfully for solubilization experiments (Chi Fru et al., in revision). Prepare in triplicate 50-mL acidwashed (5% nitric acid) sample vials, each containing 20-mL of carbonatebuffered (5 mM) deionized water (pH 8). Depending on the experiment and purity of the methanobactin extract, amend each vial with adequate homogenized crude methanobactin to achieve molar ratios greater than 1:1 mb:Cu, but ideally up to 5:1. Controls for assay should include vials with mineral but no methanobactin, methanobactin with no mineral, and possibly, sterile controls. Agitate the vials at 30–50 rpm on a shaker table at a temperature consistent with the application and analyze released copper over time, preferably for over 24 h. Treatments with different mb:Cu ratios should be used to determine whether or not adequate methanobactin has been provided for each mineral. Various approaches exist for separating minerals and cells from different treatments to allow quantification of dissolved copper release (DiSpirito et al., 1998; Fitch et al., 1993; Yu et al., 2009). Regardless of the separation method, dissolved copper can be analyzed using inductively coupled plasma mass-spectroscopy (ICP-MS) or flameless atomic adsorption spectroscopy (AA). Copper solubilization patterns as a function of mineral type, copper level, and methanobactin level are compared over time.

4.2. MMO gene expression assays for assessing Cu uptake A powerful confirmation method for the verification of methanobactin is through the performance of gene expression assays that reflect biological responses to copper uptake. A useful assay developed for this purpose quantifies the upregulation of genes related to pMMO manufacture upon exposure to different forms of copper (Knapp et al., 2007). The recommended assay quantifies pmoA transcript levels using RTqPCR in M. trichosporium OB3b, although equivalent probes-and primers can be developed if pMMO-related gene sequences are available for design. Relative to M. trichosporium OB3b, the strategy is to grow the organism under copper-free conditions such that sMMO is the active form of MMO present. The culture is then challenged with different forms of copper, and pmoA transcript levels are quantified over time. Figure 15.4 presents typical

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Isolation and Characterisation of Methanobactins

A

10

0.0:1 mb: Cu 0.2:1 2.0:1

9 8 7 6 5 0.0 mM CuCl2

4 3

pmoA log (copy/mL)

B

10 9 8 7 6 5 5.0 mM CuCl2

4 3 C

10 9 8 7 6 5 5.0 mM Cu–Fe Oxide

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Figure 15.4 pmoA gene expression versus methanobactin (mb) to copper molar ratio in Methylosinus trichosporium OB3b for (A) 0.0 mM CuCl2, (B) 5.0 mM CuCl2, and (C) 5.0 mM Cu–Fe oxide supplements provided to cultures pregrown in “Cu-free” media (adapted from Knapp et al., 2007).

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results for pmoA expression in M. trichosporium OB3b where no copper, CuCl2, and Cu-doped iron oxide are provided to the organism. The figure shows that when no additional copper is provided, no change in pmoA transcript levels is detected over time (Fig. 15.4A), whereas when copper is provided as CuCl2, pmoA transcript levels increase substantially independent of methanobactin level (Fig. 15.4B). In contrast, when copper is provided as a solid mineral oxide, the level of methanobactin significantly alters transcript levels, ranging from no change without methanobactin to transcription level equivalent to copper salt with methanobactin in stoichiometric excess (Fig, 15.4C). The difference between pmoA transcript levels with no added methanobactin and methanobactin provided in excess is a demonstration of methanobactin function. The pMMO transcription assay involves three steps: culture pregrowth, assay performance, and transcript quantification. For M. trichosporium OB3b, stock cultures are grown in copper-free NMS media in 125-mL Tygonplugged serum vials (30-mL volume) under a 50% methane/50% air atmosphere at 30  C on an incubated orbital shaker table (200 rpm). After growth to an OD600 of 0.3, sMMO activity is verified using the naphthalene– naphthol assay (DiSpirito et al., 1998), and if sMMO activity is strongly apparent, the culture is centrifuged for 10 min at 10,000g at 4  C. The resulting pellet is washed and recentrifuged three additional times in fresh copper-free NMS media to remove extracellular methanobactin. The final pellet is retained in 30 mL of fresh media for subsequent assays. The above volumes can be modified depending upon the number of specific comparative assays to be performed, including controls, but one must remember that any transcription assay is very time-dependent and performing too many samples and replicates can reduce replicability among treatments. The 30-mL washed culture is divided into 10-mL aliquots and transferred to three 30-mL crimp-sealed glass vials and amended with 5 mL of research-grade methane. The vials are then placed on a shaker table at 30  C (200 rpm) and equilibrated for 30 min. Each vial is then provided with different treatments, such as no copper, 5 mM copper as CuCl2, and 5 mM Cu-doped iron oxide (or any other mineral), and/or further variations including differing mb:Cu supply ratios. Immediately before Cu-spiking, triplicate 300-mL samples are aseptically collected at time zero as controls and then, subsequent samples are collected for 30 min; pmoA transcript signals typically maximize after 10 min. Each withdrawn volume is transferred immediately to labeled microcentrifuge tubes containing 1 mL TrizolÒ LS reagent (Invitrogen, Carlsbad CA), and then immediately frozen on dry ice until completion of the experiment. All samples are then stored at
80  C prior to transcript quantification. Frozen samples are thawed on ice and homogenized for 20 s using a FastPrep (Qbiogene) cell disruptor. RNA is then isolated from Trizol reagent by incubating the samples with 0.2 mL chloroform for 5 min and centrifuging

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at 4  C for 15 min at 10,000g. The RNA-containing aqueous phase is purified using the RNeasyÒ MinEluteTM Cleanup Kit (QIAGEN), according to the manufacturer’s instructions for QIAzol extracted samples. Product RNA is eluted with 30 mL RNase-free (DEPC-treated) water into RNasefree microcentrifuge tubes with 30 units (1 mL) Prime RNase Inhibitor (Eppendorf), divided into two replicates, and stored again at
80  C. Transcription product for pmoA is detected using specific primers and fluorogenic probes designed for M. trichosporium OB3b (Knapp et al., 2007), although equivalent primers and probes can be designed for other species. It is recommended that 16S-rRNA housekeeping genes, or the equivalent, also be quantified to monitor general metabolic cell responses to the different treatments. Primer sequences for M. trichosporium 16S-rRNA were adapted from Gulledge et al. (2001) and Holmes et al. (1995). SBYR green is used to detect 16S-rRNA gene responses. For pmoA, RT-qPCR is conducted using the TaqManÒ EZ RT-PCR Kit (Applied Biosystems) according to the manufacturer’s specifications, although volumes must be scaled. A typical PCR program is 2 min at 50  C, 30 min at 60  C, 5 min at 95  C, and 40 cycles at 94  C (20 s), annealing temperature (60 s) and 72  C (30 s), although this must be optimized for each application. Procedures for generating reaction standards to monitor RT-qPCR responses are summarized in Knapp et al. (2007). Transcription signals are then compared statistically among treatments to determine the relative influence of methanobactin level on pMMO-related transcription as a function copper abundance and mineral source.

ACKNOWLEDGMENTS The authors thank Chris Dennison, Neil Gray, Abdelnasser El Ghazouani, Charles Knapp, Jennifer Roberts, Helen Talbot, and Dennis Winge for their valuable conversations pertinent to this chapter. This was supported by NERC grant NE/F00608X/1, NSF grant EAR 0433980, and EU Marie Curie Excellence Programme grant MEXT-CT-2006-023469.

REFERENCES Atkinson, R. A., Salah El Din, A. L., Kieffer, B., Lefe`vre, J. F., and Abdallah, M. A. (1998). Bacterial iron transport: 1H NMR determination of the three-dimensional structure of the gallium complex of pyoverdin G4R, the peptidic siderophore of Pseudomonas putida G4R. Biochemistry 37, 15965–15973. Balasubramanian, R., and Rosenzweig, A. C. (2008). Copper methanobactin: A molecule whose time has come. Curr. Opin. Chem. Biol. 12, 245–249. Balasubramanian, R., Levinson, B. T., and Rosenzweig, A. C. (2010a). Secretion of flavins by three species of methanotrophic bacteria. Appl. Environ. Microbiol. 10.1128/ AEM.00935-10.

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Balasubramanian, R., Smith, S. M., Yatsunyk, L. A., Stemmler, T. L., and Rosenzweig, A. C. (2010b). Oxidation of methane by a biological dicopper center. Nature 465, 115–119. Behling, L. A., Hartsel, S. C., Lewis, D. E., DiSpirito, A. A., Choi, D. W., Masterson, L. R., Veglia, G., and Gallagher, W. H. (2008). NMR, mass spectrometry and chemical evidence reveal a different chemical structure for methanobactin that contains oxazolone rings. J. Am. Chem. Soc. 130, 12604–12605. Chi Fru, E., Gray, N. D., McCann, C., Baptista, J. C., Christgen, B., Talbot, H. M., El Ghazouani, A., Dennison, C., and Graham, D. W. Effects of copper mineralogy and methanobactin on cell growth and sMMO activity in Methylosinus trichosporium OB3b. Biogeoscience. Submitted. Choi, D. W., Do, Y. S., Zea, C. J., McEllistrem, M. T., Lee, S.-W., Semrau, J. D., Pohl, N. L., Kisting, C. J., Scardino, L. L., Hartsel, S. C., Boyd, E. S., Geesey, G. G., et al. (2006). Spectral and thermodynamic properties of Ag(I), Au(III), Cd(II), Co(II), Fe(III), Hg(II), Mn(II), Ni(II), Pb(II), U(IV), and Zn(II) binding by methanobactin from Methylosinus trichosporium OB3b. J. Inorg. Biochem. 100, 2150–2161. Choi, D. W., Semrau, J. D., Antholine, W. E., Hartsel, S. C., Anderson, R. C., Carey, J. N., Dreis, A. M., Kenseth, E. M., Renstrom, J. M., Scardino, L. L., Van Gorden, G. S., Volkert, A. A., et al. (2008). Oxidase, superoxide dismutase, and hydrogen peroxide reductase activities of methanobactin from types I and II methanotrophs. J. Inorg. Biochem. 102, 1571–1580. Demange, P., Bateman, A., Dell, A., and Abdallah, M. A. (1988). Structure of azotobactin D, a siderophore of Azotobacter vinelandii Strain D (CCM 289). Biochemistry 27, 2745–2752. Demange, P., Bateman, A., Mertz, C., Dell, A., Piemont, Y., and Abdallah, M. A. (1990). Bacterial siderophores: Structures of pyoverdins Pt, siderophores of Pseudomonas tolaasii NCPPB 2192, and pyoverdins Pf, siderophores of Pseudomonas fluorescens CCM 2798. Identification of an unusual natural amino acid. Biochemistry 29, 11041–11051. Dispirito, A. A., Zahn, J. A., Graham, D. W., Kim, H. J., Larive, C. K., Derrick, T. S., Cox, C. D., and Taylor, A. (1998). Copper-binding compounds from Methylosinus trichosporium OB3b. J. Bacteriol. 180, 3606–3613. El Ghazouani, A., Basle, A., Firbank, S. J., Knapp, C. W., Gray, J., Graham, D. W., and Dennison, C. (2011). Copper-binding properties and structures of methanobactins and Methylosinus trichosporium OB3b. Inorg. Chem. DOI: 10.1021/ic101965j. Fitch, M. W., Graham, D. W., Arnold, R. G., Agarwal, S. K., Phelps, P., and Georgiou, G. (1993). Phenotypic characterization of copper-resistant mutants of Methylosinus trichosporium OB3b. Appl. Environ. Microbiol. 59, 2771–2776. Fox, B. G., Froland, W. A., Jollie, D. R., and Lipscomb, J. D. (1990). Monooxygenase from Methylosinus tricosporium OB3b. Methods Enzymol. 188, 191–202. Gulledge, J., Ahmad, A., Steudler, P. A., Pomerantz, W. J., and Cavanaugh, C. M. (2001). Family- and genus-level 16S rRNA-targeted oligonucleotide probes for ecological studies of methanotrophic bacteria. Appl. Environ. Microbiol. 67, 4726–4733. Hakemian, A. S., Tinberg, C. E., Kondapalli, K. C., Telser, J., Hoffman, B. M., Stemmler, T. L., and Rosenzweig, A. C. (2005). The copper chelator methanobactin from Methylosinus trichosporium OB3b binds Cu(I). J. Am. Chem. Soc. 127, 17142–17143. Hanson, R. S., and Hanson, T. E. (1996). Methanotrophic bacteria. Microbiol. Rev. 60, 439–471. Holmes, A. J., Owens, N. J. P., and Murrell, J. C. (1995). Detection of novel marine methanotrophs using phylogenetic and functional gene probes after methane enrichment. Microbiology 141, 1947–1955. Hughes, M. N., and Poole, R. K. (1989). Metals and Microorganisms. Chapman and Hall, London.

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Kim, H. J., and Graham, D. W. (2001). Effect of oxygen level on simultaneous nitrogenase and sMMO expression and activity in Methylosinus trichosporium OB3b and its sMMO(C) mutant, PP319: Aerotolerant N2 fixation in PP319. FEMS Microbiol Lett. 201, 133–138. Kim, H. J., Graham, D. W., DiSpirito, A. A., Alterman, M. A., Galeva, N., Larive, C. K., Asunskis, D., and Sherwood, P. M. A. (2004). Methanobactin, a copper-acquisition compound from methane-oxidizing bacteria. Science 305, 1612–1615. Kim, H. J., Galeva, N., Larive, C. K., Alterman, M. A., and Graham, D. W. (2005). Purification and physiochemical properties of methanobactin: A chalkophore from Methylosinus trichosporium OB3b. Biochemistry 44, 5140–5148. Knapp, C. W., Fowle, D. A., Kulczycki, E., Roberts, J. A., and Graham, D. W. (2007). Regulation of copper acquisition and MMO gene expression by methanobactin in methane-oxidizing bacteria in the natural environment. Proc. Natl. Acad. Sci. USA 104, 12040–12045. Knosp, O., von Tigerstrom, M., and Page, W. J. (1984). Siderophore-mediated uptake of iron in Azotobacter vinelandii. J. Bacteriol. 159, 341–347. Krentz, B. D., Mulheron, H. J., Semrau, J. D., DiSpirito, A. A., Bandow, N. L., Haft, D. H., Vuilleumier, S., Murrell, J. C., McEllistrem, M. T., Hartsel, S. C., and Gallagher, W. H. (2010). Comparison of methanobactins from Methylosinus trichosporium OB3b and Methylocystis strain SB2 predicts methanobactins are synthesized from diverse peptide precursors modified to create a common core for binding and reducing copper ions. Biochemistry 10.1021/bi1014375. Kulczycki, E., Fowle, D. A., Graham, D. W., and Roberts, J. A. (2007). Methanotrophpromoted weathering of Cu-substituted borosilicate glass. Geobiology 5, 251–263. Kulczycki, E., Fowle, D. A., Kenward, P. A., Leslie, L., Graham, D. W., and Roberts, J. A. (2011). Stimulation of methanotroph activity by Cu-substituted Borosilicate Glass. Geomicrobiol. J. 28, 1–10. Murrell, J. C., McDonald, I. R., and Gilbert, B. (2000). Regulation of expression of methane monooxygenases by copper ions. Trends Microbiol. 8, 221–225. Neilands, J. B. (1982). Microbial envelope proteins related to iron. Annu. Rev. Microbiol. 36, 285–309. Payne, S. M. (1994). Detection, isolation and characterization of siderophores. Methods Enzymol. 235, 329–344. Salih, B., Masselon, C., and Zenobi, R. (1998). Matrix-assisted laser desorption/ionization mass spectrometry of noncovalent protein-transition metal ion complexes. J. Mass Spectrom. 33, 994–1002. Teintze, M., Hossain, M. B., Barnes, C. L., Leong, J., and van der Helm, D. (1981). Structure of ferric pseudobactin, a siderophore from a plant growth promoting Pseudomonas. Biochemistry 20, 6446–6457. Visca, P., Imperi, F., and LamontI, L. (2007). Pyoverdine siderophores: From biogenesis to biosignificance. Trends Microbiol. 15, 22–30. Vraspir, J. M., and Butler, A. (2009). Chemistry of marine ligands and siderophores. Annu. Rev. Mar. Sci. 1, 43–63. Yu, Y., Ramsay, J. A., and Ramsay, B. A. (2009). Use of allylthiourea to produce soluble methane monooxygenase in the presence of copper. Appl. Microbiol. Biotechnol. 82, 333–339. Zahn, J. A., and Dispirito, A. A. (1996). The membrane-associated methane monooxygenase from Methyloccus capsulatus (Bath). J. Bacteriol. 178, 1018–1029.

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A Simple Assay for Screening Microorganisms for Chalkophore Production Sukhwan Yoon,*,1 Alan A. DiSpirito,† Stephan M. Kraemer,‡ and Jeremy D. Semrau* Contents 1. Introduction 2. Cu–CAS Assay for Chalkophore Detection 2.1. Preparation of Cu–CAS solution 2.2. Preparation of Cu–CAS agar plates 3. Fe–CAS Assay for Detecting Nonspecific Binding of Copper from Cu–CAS by a Siderophore 3.1. Preparation of liquid Fe–CAS 3.2. Preparation of Fe–CAS agar plates 4. Conclusions Acknowledgments References

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Abstract Recently, methanotrophs were found to exude a chalkophore, that is, a metal ligand with great affinity and specificity to copper. A rapid screening method for chalkophore expression was developed by adopting the chrome azurol S (CAS) assay originally used for detecting siderophore production in diverse groups of bacteria and fungi. In this assay, iron(III) chloride was replaced with copper(II) chloride. Both liquid and agar plate versions of the Cu–CAS assay can be used to examine the activity of either isolated methanobactin or to screen organisms for production of a chalkophore. Although here we describe the use of this assay to screen methanotrophs for chalkophore production, it can be modified as necessary to screen other organisms for chalkophore production as well. * Department of Civil and Environmental Engineering, The University of Michigan, Ann Arbor, Michigan, USA Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA { Department of Environmental Geosciences, University of Vienna, Althanstrasse, Vienna, Austria 1 Current Address: Department of Biogeochemistry, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany {

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00016-4

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Many siderophores can also bind copper in the presence of CAS. Therefore, this assay should be done in conjunction with the original iron–CAS assay to determine if any positive Cu–CAS assay results are due to nonspecific binding of copper by a siderophore. This inexpensive assay may also aid in analyses of the genetics of chalkophore synthesis.

1. Introduction It is well-known that copper plays a significant role in methanotrophic metabolism, not only controlling the relative expression of particulate and soluble methane monooxygenases (pMMO and sMMO, respectively), but also affecting the activity of pMMO, production of intracytoplasmic membranes, and expression of other enzymes, for example, different forms of the formaldehyde dehydrogenase (Semrau, et al., 2010). As such, it was long suspected that methanotrophs had a copper-specific uptake mechanism, or chalkophore, with the first example, methanobactin from Methylococcus capsulatus Bath, reported in 1996 as the copper-binding compound (cbc; Zahn and DiSpirito, 1996). This unique compound, analogous to a siderophore, was found to utilize oxazalone rings for metal binding (Behling, et al., 2008). Although some physicochemical properties of methanobactin, for example, affinity to copper and molecular structure, have been reported (Choi et al., 2006a,b, 2008; Kim et al., 2004), there has not been any significant progress in characterizing either the diversity of organisms that can express such compounds, or the genetics underpinning the synthesis of chalkophores, including methanobactin from Methylosinus trichosporium OB3b. Such data are important as empirical evidence suggests that methanobactin may be involved in regulation of expression of methane monooxygenases in M. trichosporium OB3b (Choi et al., 2010; Knapp et al., 2007; Morton et al., 2000). If methanotrophs produce different chalkophores with varying copper affinity, it may play a role in regulating methanotrophic community structure. Furthermore, it has been found that methanobactin from M. trichosporium OB3b can reduce Au(III) to Au(0), forming gold nanoparticles, and thus can have significant industrial potential (Choi et al., 2006a). The reported yields of methanobactin are less than 60 mg/L of spent medium, however (Choi et al., 2008), and the potential use of methanobactin in such industrial applications will require larger amounts of methanobactin. One strategy to increase the yield of methanobactin production would be the heterologous expression of genes involved in methanobactin synthesis in a less fastidious host such as Escherichia coli. The lack of simple and economic methods to screen for a wide variety of microorganisms has limited our ability to determine how wide-spread the

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ability to produce a chalkophore might be. For example, do all methanotrophs require such a mechanism to collect copper? Do closely related cells such as ammonia-oxidizing bacteria have a similar active copper-uptake system? Furthermore, the lack of a simple screening assay has limited our ability to deduce the genetics of methanobactin synthesis through the use of high-throughput techniques to screen putative chalkophore-minus mutants in organisms known to produce such a compound. To address these problems, we have devised a novel method to qualitatively screen for production of chalkaphores through a simple plate assay (Yoon et al., 2010). This assay was developed by adopting the chrome azurol S (CAS) assay for siderophore production (Schwyn and Neilands, 1987), to instead screen cultures for chalkophore production by substituting copper for iron. In the original assay, a blue complex is formed between iron and CAS in the presence of a detergent, hexadecyltrimethylammonium bromide (HDTMA). The removal of iron by a competing ligand, for example, a siderophore, results in a color change, typically from blue to orange. Here copper was substituted for iron, as CAS also has a high affinity for copper with a blue complex also being observed, with a similar color change observed when copper is abstracted by competing ligands.

2. Cu–CAS Assay for Chalkophore Detection The Cu–CAS assay was developed from the popular CAS assay originally developed to assay for the presence of siderophore production (Schwyn and Neilands, 1987). As in the original CAS assay, CAS binds copper with relatively high affinity (log K ¼ 13.2; Cha and Abruna, 1990) when there is no competing ligand in the medium. As the only characterized methanobactin to date has much higher affinity towards copper than CAS, that is, 3.3  1034  3.0  1011 M 1 (Choi, et al., 2006a), abstraction of copper from CAS should occur as methanobactin outcompetes CAS in binding copper. In the Cu–CAS assay described here, the colorimetric change associated with the loss of copper from CAS is used for detecting the presence of the methanobactin.

2.1. Preparation of Cu–CAS solution Cu–CAS solution for detection of methanobactin synthesis in methanotrophs is prepared with nitrate mineral salt medium (NMS; Whittenbury et al., 1970) and CAS and HDTMA (Sigma, St. Louis, MO). It should be noted that this assay can be easily modified for other organisms by substituting the appropriate growth medium as long as the color of the medium does not interfere with the color of Cu–CAS and no significant competition of

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trace nutrient metal ions for complexation by CAS occurs. Below we provide step-by-step instructions for preparing Cu–CAS solutions for screening methanotrophs for chalkophore production. 1. To produce 500 mL of Cu–CAS solution with 50 mM Cu2þ, first prepare NMS medium by adding 0.5 g MgSO47H2O, 0.5 g KNO3, 0.1 g CaCl22H2O, 50 mL of 3.8% (w/v) Fe–EDTA, 250 mL of 0.1% (w/v) NaMoO42H2O to 450 mL of distilled deionized water (>18 MOcm; Whittenbury et al., 1970). Then add 0.5 mL of trace element solution (50 mg/L FeSO47H2O, 40 mg/L ZnSO47H2O, 2 mg/L MnCl27H2O, 5 mg/L CoCl26H2O, 1 mg/L NiCl26H2O, 1.5 mg/L H3BO3, and 25 mg/L EDTA). 2. Prepare stock solutions of CuCl2 (Fisher Scientific, Pittsburg, PA), CAS, and HDTMA at concentrations of 5, 1.05, and 2.625 mM, respectively in distilled deionized water ( >18 MOcm). Stir these stock solutions until all solids are dissolved and the solutions are well-mixed. 3. Add 25 mL of the CAS stock solution to 5 mL of the CuCl2 stock solution. Next add 20 mL of HDTMA under stirring for final concentrations of 0.5, 0.525, and 1.050 mM of, Cu2þ, CAS, and HDTMA, respectively. This 10 stock solution of Cu–CAS should have a purple color at this stage. 4. Sterilize both the NMS growth medium and 10 Cu–CAS solution via autoclaving for 40 min. 5. Allow both the NMS growth medium and 10 Cu–CAS solution to cool to room temperature. 6. Pipette 50 mL of the 10 Cu–CAS concentrate into 450 mL of NMS growth medium. 7. Add 5 mL of sterile vitamin stock solution (20 mg/L biotin, 2.0 mg/L folic acid, 5.0 mg/L thiamin HCl, 5.0 mg/L Ca pantothenate, 0.1 mg/L vitamin B12, 5.0 mg/L riboflavin, and 5.0 mg/L nicotiamide; Lidstrom, 1988) to the combined NMS growth medium and Cu–CAS solution. 8. Buffer the mixture to pH  6.8 by adding 5 mL of sterile phosphate buffer (26 g/L KH2PO4 and 62 g/L Na2HPO47H2O). 9. Distribute the NMS Cu–CAS mixture as 5-mL aliquots into 20 mL vials. The vials can be sealed with rubber butyl stoppers (National Scientific Co. Duluth, GA) if isolation from the atmosphere is necessary. 10. Solutions of interest, for example, concentrates of isolated methanobactin or spent microbial growth medium can be screened at this point by adding volumes less than 500 mL. A color change from bright blue to yellow should be observed shortly after addition, as well as substantial changes in the UV/Vis absorption spectra, particularly at the wavelengths associated with the heterocyclic rings, for example, 340 and 394 nm for methanobactin from M. trichosporium OB3b (Fig.16.1).

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Figure 16.1 UV–visible absorption spectra of 50 mM of methanobactin from M. trichosporium OB3b (▪ ▪ ▪ ▪ ▪), 50 mM Cu–CAS (- - - - - -), and 50 mM methanobactin from M. trichosporium OB3b plus 50 mM Cu–CAS (————) after 5 min incubation at room temperature. Insert: (A) 50 mM Cu–CAS; (B) 50 mM methanobactin from M. trichosporium OB3b plus 50 mM Cu–CAS.

Growth of methanotrophs in liquid NMS–Cu–CAS solution is unlikely due to the toxicity of HDTMA. Thus, this liquid assay may best be used as an initial screen to check for methanobactin production of methanotrophs grown under different growth conditions.

2.2. Preparation of Cu–CAS agar plates As methanotrophic growth can be significantly compromised by the presence of the detergent used in the standard CAS assay, that is, HDTMA, the split-CAS assay devised by Milagres et al. (1999) was adapted for detecting the excretion of methanobactin from growing methanotrophs. Here split plates are made with one half containing Cu–CAS/NMS agar and the other with NMS agar only. Strains are streaked on the NMS agar, and methanobactin production is determined from color changes in the Cu–CAS/NMS agar due to diffusion of methanobactin. This assay does not inhibit methanotrophic growth and can be used to detect methanobactin production as the color changes are visible typically within 2 weeks. Below are step-bystep instructions for preparing split Cu–CAS/NMS and NMS agar plates to screen methanotrophs for chalkophore production. 1. Prepare 450 mL of NMS growth medium as described in step 1 of Section 2.1. Add 7.5 g Bacto agar (Bectron Dickinson, Franklin Lakes, NJ). 2. Prepare a 10 stock solution of Cu–CAS as described in steps 2 and 3 of Section 2.1. 3. Autoclave the NMS agar and 10 Cu–CAS solution as described in step 4 of Section 2.1.

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4. Allow both the NMS agar and 10 Cu–CAS solution to cool to 50  C. 5. Carefully add 50 mL of the 10 Cu–CAS concentrate to 450 mL NMS agar. 6. Add vitamin and phosphate buffer solutions as described in steps 7 and 8 of Section 2.1. 7. Pour the Cu–CAS/NMS agar into standard petri dishes. After the agar has completely solidified, carefully remove half with a sterilized razor. 8. Prepare an additional 450 mL of NMS growth medium as described above in Section 2.1. Add 7.5 g Bacto agar. Autoclave the NMS agar for 40 min and allow to cool to 50  C. 9. Add vitamin and phosphate buffer solutions as described in steps 7 and 8 of Section 2.1. 10. Add the desired copper concentration to the NMS agar using a sterile stock solution of 10 mM CuCl2. Previous results have shown that copper concentrations ranging between 0 and 10 mM do not have any impact on the results of the experiments (Yoon et al., 2010). It is recommended that at least 1 mM of copper be added to the NMS agar as copper limitation may result in repressed growth of some methanotrophic strains. 11. Carefully pour the NMS agar into the empty space in the agar plate created in step 7 above. The surface of both the Cu–CAS/NMS agar and NMS agar should be level. 12. Streak methanotroph(s) of interest on the NMS agar half of the plate. Streaking methanotrophs onto the NMS agar only will prevent inhibition of microbial growth by HDTMA as there is no direct contact. It is important that cells be streaked as closely as possible to the boundary of the Cu–CAS/NMS and NMS agars, however, to reduce the time required for any chalkophore produced to diffuse into the Cu–CAS/ NMS agar. 13. Incubate split Cu–CAS/NMS and NMS agar plates in a sealed container with a 1:1 air-to-methane ratio at the optimal growth temperature of the methanotroph(s) to be tested. A color change from blue to yellowish-orange should begin to appear on Cu–CAS/NMS agar within 2 weeks if a chalkophore is produced (Fig.16.2).

3. Fe–CAS Assay for Detecting Nonspecific Binding of Copper from Cu–CAS by a Siderophore Although much more specific to iron, some siderophores also bind copper with a relatively high affinity. For example, the bacterial siderophore deferoxamine-B forms a copper-complex with a 1:1 log formation constant

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6 Days

15 Days

Figure 16.2 An example of the Cu–CAS plate assay performed with M. trichosporium OB3b. The copper concentration in the NMS medium was 1 mM. Within  15 days from inoculation, color change from blue to yellow was obvious in the Cu–CAS/NMS agar due to the production and diffusion of methanobactin from M. trichosporium OB3b streaked on NMS agar.

of 14.1 (Martell et al., 2001) that is high enough to abstract copper from CAS (Yoon et al., 2010). Therefore, separate Fe–CAS assays are required to confirm that positive results from Cu–CAS assay are not due to production of siderophores. Some methanotrophs, for example, M. trichosporium OB3b and Methylomicrobium album BG8, have been found to produce siderophores (Yoon et al., 2010). Thus the standard Fe–CAS assay should be used to examine the possible production of a siderophore by methanotrophs as described below. We would like to note that the following procedures were originally developed by Schwyn and Neilands (1987) for detection of siderophore production, and are optimized here for screening of methanotrophs.

3.1. Preparation of liquid Fe–CAS 1. Prepare 450 mL of NMS solution as described in step 1 of Section 2.1. Add 15 g of PIPES and stir until completely dissolved. Adjust the pH of the NMS solution to 6.8 using 50% (w/v) NaOH. Note: Phosphate buffer cannot be used in Fe–CAS assays, as phosphate competes for Fe (III) bound to CAS and can cause a significant color change (Schwyn and Neilands, 1987). 2. Prepare stock solutions of CAS, FeCl3 (Fisher Scientific, Pittsburg, PA), and HDTMA at concentrations of 1.05, 5, and 2.625 mM, respectively using distilled deionized water ( >18 MOcm). Stir until all solids are dissolved and the solutions are well-mixed.

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3. Add 25 mL of the CAS stock solution to 5 mL of the FeCl3 stock solution. Next add 20 mL of HDTMA under stirring for final concentrations of 0.5, 0.525, and 1.050 mM of Fe3þ, CAS, and HDTMA, respectively. This 10 stock solution of Fe–CAS should have a dark blue color at this stage. 4. Sterilize the NMS medium and 10 Fe–CAS via autoclaving for 40 min. 5. Pipette 50 mL of the 10 Fe–CAS concentrate into 450 mL of NMS growth medium. 6. Distribute the Fe–CAS/NMS solution as 5-mL aliquots into 20 mL vials. The vials can be sealed with rubber butyl stoppers if isolation from the atmosphere is necessary. 7. Add materials to be tested as described in step 10 of Section 2.1. If a siderophore or other iron chelator, for example, deferoxamine-B, is present in the medium, the blue tint of the Fe–CAS solution will change to yellow, while when methanobactin from M. trichosporium OB3b is added, a greenish color is observed (Fig.16.3). It should also be noted that collecting the UV/Vis absorption spectra can also serve as an effective methodology to determine if a siderophore is present as the peaks associated with heterocyclic rings (340 and 394 nm for M trichosporium OB3b) are still apparent in the presence of Fe–CAS, while they diminish in the presence of Cu–CAS (Fig.16.1). 1.6

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Figure 16.3 UV–visible absorption spectra of 50 mM OB3b–mb (▪ ▪ ▪ ▪ ▪), 50 mM Fe– CAS (- - - - - -), 50 mM Fe–CAS plus 50 mM deferoxamine-B (—   —), and 50 mM methanobactin from M. trichosporium OB3b plus 50 mM Fe–CAS (————) after 5 min incubation at room temperature. Insert: (A) 50 mM Fe–CAS, (B) 50 mM Fe–CAS plus 50 mM deferoxamine-B, (C) 50 mM methanobactin from M. trichosporium OB3b plus 50 mM Fe–CAS.

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3.2. Preparation of Fe–CAS agar plates A Fe–CAS plate assay can be used for detecting siderophore production that may give false positive results for chalkophore production when using the Cu–CAS plate assay. Siderophore production can be repressed if sufficient iron is present in the growth medium (Crosa, 1989) and the standard NMS agar as used in the split Cu–CAS/NMS agar plates contains an appreciable amount of iron. Thus, it is unlikely that siderophores will interfere with the Cu–CAS assay, but such an assumption should be tested by screening for siderophore production using Fe–CAS/NMS agar plates. Below are stepby-step instructions for preparing split Fe–CAS/NMS and NMS agar plates to screen methanotrophs for siderophore production. 1. Prepare 450 mL of NMS growth medium as described in step 1 of Section 3.1. Adjust the pH of the NMS solution to 6.8 using 50% (w/v) NaOH. Add 7.5 g Bacto agar. 2. Prepare a 10 stock solution of Fe–CAS as described in steps 2 and 3 of Section 3.1. 3. Sterilize the NMS agar and 10 Fe–CAS via autoclaving for 40 min. 4. Allow both the NMS agar and 10 Fe–CAS stock solution to cool to 50  C. 5. Carefully add 50 mL of the 10 Fe–CAS stock solution to 450 mL NMS agar. 6. Add 5 mL of sterile vitamin stock solution (20 mg/L biotin, 2.0 mg/L folic acid, 5.0 mg/L thiamin HCl, 5.0 mg/L Ca pantothenate, 0.1 mg/L vitamin B12, 5.0 mg/L riboflavin, and 5.0 mg/L nicotinamide; Lidstrom, 1988) to the combined NMS growth agar and Fe–CAS solution. 7. Pour the Fe–CAS/NMS agar into standard petri dishes. After the agar has completely solidified, carefully remove half with a sterilized razor. 8. Prepare 450 mL of NMS growth agar as described in steps 8-10 of Section 2.2 with the appropriate copper concentration. As stated earlier, it is recommended that at least 1 mM of copper be added to the NMS agar as copper limitation may result in repressed growth in some methanotrophic strains. One may prepare NMS agar with and without Fe–EDTA, but the presence of iron in NMS agar will limit siderophore synthesis in most cases. Add 7.5 g Bacto agar to the NMS medium. 9. Carefully pour the NMS agar into the empty space in the agar plate created in step 7 above. The surface of the both the Fe–CAS/NMS agar and NMS agar should be level. 10. Streak methanotroph(s) of interest on the NMS agar half of the plate. Streaking methanotrophs onto the NMS agar only will prevent inhibition of microbial growth by HDTMA as there is no direct contact. It is important that cells be streaked as closely as possible to the boundary of the Fe–CAS/NMS and NMS agars, however, to reduce the time required for any siderophore produced to diffuse into the Fe–CAS/NMS agar.

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6 Days

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Figure 16.4 An example of the Fe–CAS plate assay performed with M. trichosporium OB3b. Ferric iron was present in the NMS agar as Fe–EDTA at a final concentration of 3.8  10 4% (w/v). After 15 days of inoculation, little disappearance of blue color in the Fe–CAS/NMS agar was observed, indicating a small amount of siderophore production.

11. Incubate split Fe–CAS/NMS and NMS agar plates in a sealed container with a 1:1 air-to-methane ratio at the optimal growth temperature of the methanotroph(s) to be tested. A color change from blue to yellowish-orange should begin to appear on Fe–CAS/NMS agar within 2 weeks if a siderophore is produced (Fig.16.4).

4. Conclusions Here we provide simple instructions for an assay that can be used to screen methanotrophs for chalkophore production. This assay can be modified to screen other cells for chalkophore production by substituting the appropriate growth medium for NMS agar. With this assay, the diversity of microorganisms that produce such copper-specific binding compounds can be more readily determined, and also can be used to help elucidate the genetics of chalkophore synthesis. As other biogenic metal binding compounds, for example, siderophores, can also abstract copper from this assay, parallel studies should be performed with both Cu–CAS and Fe–CAS split plates to determine if and under what conditions cells are producing siderophores that may give a false positive for chalkophore production when using Cu–CAS split plates.

ACKNOWLEDGMENTS Support from the Department of Energy (DE-FC26-05NT42431), the Carl Page Foundation, and the University of Vienna to JDS is gratefully acknowledged.

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REFERENCES Behling, L. A., Hartsel, S. C., Lewis, D. E., DiSpirito, A. A., Choi, D. W., Masterson, L. R., Veglia, G., and Gallagher, W. (2008). NMR, mass spectrometry and chemical evidence reveal a different chemical structure for methanobactin that contains oxazolone rings. J. Am. Chem. Soc. 130, 12604–12605. Cha, S. K., and Abruna, H. D. (1990). Determination of copper at electrodes modified with ligands of varying coordination strength: A preamble to speciation studies. Anal. Chem. 62, 274–278. Choi, D. W., Do, Y. S., Zea, C. J., McEllistrem, M. T., Lee, S. W., Semrau, J. D., Pohl, N. L., Kisting, C. J., Scardino, L. L., Hartsel, S. C., Boyd, E. S., Geesey, G. G., et al. (2006a). Spectral and thermodynamic properties of Ag(I), Au(III), Cd(II), Co(II), Fe (III), Hg(II), Mn(II), Ni(II), Pb(II), U(IV), and Zn(II) binding by methanobactin from Methylosinus trichosporium OB3b. J. Inorg. Biochem. 100, 2150–2161. Choi, D. W., Zea, C. J., Do, Y. S., Semrau, J. D., Antholine, W. E., Hargrove, M. S., Pohl, N. L., Boyd, E. S., Geesey, G. G., Hartsel, S. C., Shafe, P. H., McEllistrem, M. T., et al. (2006b). Spectral, kinetic, and thermodynamic properties of Cu(I) and Cu(II) binding by methanobactin from Methylosinus trichosporium OB3b. Biochemistry 45, 1442–1453. Choi, D.-W., Semrau, J. D., Antholine, W. E., Hartsel, S. C., Anderson, R. C., Carey, J. N., et al. (2008). Oxidase, superoxide dismutase, and hydrogen peroxide reductase activities of methanobactin from types I and II methanotrophs. J. Inorg. Biochem. 102, 1571–1580. Choi, D. W., Bandow, N., McEllistem, T. M., Semrau, J. D., Antholine, W. E., Hartsel, S. C., Gallagher, W., Zea, C. J., Pohl, N. L., Zahn, J. A., and DiSpirito, A. A. (2010). Spectral and thermodynamic properties of methanobactin from Mehylomicrobium album BG8 and Methylococcus capsulatus Bath: A case for copper competition on a molecular level. J. Inorg. Biochem. 104, 1240–1247. Crosa, J. H. (1989). Genetics and molecular biology of siderophore-mediated iron transport in bacteria. Microbiol. Rev. 53, 517–530. Kim, H. J., Graham, D. W., DiSpirito, A. A., Alterman, M. A., Galeva, N., Larive, C. K., Asunskis, D., and Sherwood, P. M. A. (2004). Methanobactin, a copper-acquisition compound from methane-oxidizing bacteria. Science 305, 1612–1615. Knapp, C. W., Fowle, D. A., Kulczycki, E., Roberts, J. A., and Graham, D. W. (2007). Methane monooxygenase gene expression mediated by methanobactin in the presence of mineral copper sources. Proc. Natl. Acad. Sci. USA 104, 12040–12045. Lidstrom, M. E. (1988). Isolation and characterization of marine methanotrophs. Antonie Leeuwenhoek 54, 189–199. Martell, A. E., Smith, R. M., and Motekaitis, R. J. (2001). NIST Critically Selected Stability Constants of Metal Complexes. NIST standard reference database 46, Version 6.0, NIST Gaithersburg, MD. Milagres, A. M. F., Machuca, A., and Napolea˜o, D. (1999). Detection of siderophore production from several fungi and bacteria by a modification of chrome azurol S (CAS) agar plate assay. J. Microbiol. Methods 37, 1–6. Morton, J. D., Hayes, K. F., and Semrau, J. D. (2000). Effect of copper speciation on wholecell soluble methane monooxygenase activity in Methylosinus trichosporium OB3b. Appl. Environ. Microbiol. 66, 1730–1733. Schwyn, B., and Neilands, J. B. (1987). Universal chemical assay for the detection and determination of siderophores. Anal. Biochem. 160, 47–56. Semrau, J. D., DiSpirio, A. A., and Yoon, S. (2010). Methanotrophs and copper. FEMS Microbiol. Rev. 34, 496–531.

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Whittenbury, R., Phillips, K. C., and Wilkinson, J. F. (1970). Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61, 205–218. Yoon, S., Kraemer, S. M., DiSpirito, A. A., and Semrau, J. D. (2010). An assay for screening microbial cultures for chalkophore production. Environ. Microbiol. Rep. 2, 295–303. Zahn, J. A., and DiSpirito, A. A. (1996). Membrane-associated methane monooxygenase from Methylococcus capsulatus Bath. J. Bacteriol. 178, 1018–1029.

C H A P T E R

S E V E N T E E N

Isolation of Methanobactin from the Spent Media of Methane-Oxidizing Bacteria Nathan L. Bandow,* Warren H. Gallagher,† Lee Behling,† Dong W. Choi,‡ Jeremy D. Semrau,§ Scott C. Hartsel,† Valerie S. Gilles,* and Alan A. DiSpirito* Contents 260 260 261 261 263 265 266 268 268

1. Introduction 2. Isolation of Methanobactin from the Spent Media of MOB 2.1. Maximizing yields in the spent media 2.2. Separation of mb from whole cells 2.3. Concentration of mb from spent media 3. Purification of mb 4. Sample Variability Acknowledgments References

Abstract Chalkophores are low molecular mass modified peptides involved in copper acquisition in methane-oxidizing bacteria (MOB). A screening method for the detection of this copper-binding molecule is presented in Chapter 16. Here we describe methods to (1) maximize expression and secretion of chalkophores, (2) concentrate chalkophores from the spent media of MOB, and (3) purify chalkophores.

* Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA Department of Chemistry, University of Wisconsin-Eau Claire, Eau Claire, Wisconsin, USA Department of Biological and Environmental Science, Texas A&M University-Commerce, Commerce, Texas, USA } Department of Civil and Environmental Engineering, The University of Michigan, Ann Arbor, Michigan, USA { {

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00017-6

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1. Introduction Chalkophores (Greek term for copper bearer or copper carrier) are a group of small molecular mass modified peptides secreted by methaneoxidizing bacteria (MOB) in response to copper deficiencies (Semrau et al., 2010). Currently, methanobactin (mb) from MOB is the only characterized molecule of this class of metal-binding chromopeptides. mb shows a number of structural similarities to amino acid-containing pyroverdin class of iron-binding siderophores (Ongena et al., 2001; Vossen and Taraz, 1999). Like pyroverdins, methanobactins are composed of 8–11 amino acids plus additional non-amino acid constituents. One distinguishing structural characteristic of methanobactins is the presence of two five-membered rings with an associated enethiol that is involved in metal coordination (Behling et al., 2008; Kim et al., 2004). This five-membered ring has been found to be either an oxazolone or an imidazolone depending on the mb characterized to date (Behling et al., 2008; Krentz et al., 2010). Other properties distinguishing chalkophores from iron-binding siderophores include (1) the preferential binding of copper over iron and other metals (Choi et al., 2006a), (2) the variety of metals bound by chalkophores (Choi et al., 2006b), and (3) copper displacement of iron and most other metals (Choi et al., 2006b). Similar to siderophores, which are produced in response to iron limitations, chalkophores are excreted by MOB in response to copper limitations (Choi et al., 2010). Chapter 16 presents a modification of the chromo azurol S (CAS) assay that can be used to identify and distinguish chalkophores from siderophores. This chapter focuses on methods to maximize chalkophore concentrations in the extracellular fraction and methods to purify this molecule from the spent medium of different MOB.

2. Isolation of Methanobactin from the Spent Media of MOB The following discussion is based on the isolation of mb from four different MOB, two capable of expressing both the soluble methane monooxygenase (sMMO) and membrane-associated or particulate methane monooxygenase (pMMO)—Methylosinus trichosporium OB3b and Methylococcus capsulatus Bath—and two MOB which do not have the genes for the sMMO and constitutively express the pMMO—Methylobacterium album BG8 and Methylocystis strain SB2.

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2.1. Maximizing yields in the spent media Copper-bound methanobactin (Cu-mb) can be isolated from whole cells, the washed membrane fraction of MOB expressing the pMMO or from the spent media (Zahn and DiSpirito, 1996). Isolation of mb from whole cells or from the washed membrane fraction requires an organic extraction with 100% N, N 0 -dimethylformamide (DMF) followed by chromatography on silica gel column and HPLC chromatography and is generally not recommended for a number of reasons. For example, the DMF extraction will solubilize a number of membrane components, which complicates purification. In addition, sample loss due to additional purification steps as well as sample breakdown is problematic. Last, when isolated from whole cells or washed membrane fraction, Cu-mb is the sole product, which requires extensive dialysis against Na-ethylene diamine tetraacetate to remove the majority (75–90%) of the bound copper. This procedure also results in an mb sample with altered copper-binding properties (Choi et al., 2006a,b; Kim et al., 2005). Thus, extraction from the spent media is recommended. In addition to avoiding the problems stated above, purification from the spent media is comparatively simple and the primary product is copper-free mb. The first and by far most time consuming step in purification of mb from the extracellular fraction of MOB is to maximize yields in the extracellular fraction. Copper concentration in the culture media is the only variable identified to date that influences excretion of mb (Choi et al., 2006a,b, 2008, 2010; Zahn and DiSpirito, 1996). As shown in Fig. 17.1, copper has a dramatic effect on the extracellular concentration of mb. In MOB capable of expressing both forms of the methane monooxygenase, the highest concentrations of mb in the spent media have been found to occur when the initial copper concentration is between 0.1 and 0.7 mM with cells expressing the sMMO. If the initial copper concentration is below 0.1 mM, the culture may initially secrete high concentrations of mb. However, concentration of mb in the spent media decreases rapidly in subsequent batches if the initial copper concentration is too low (Choi et al., 2006a,b, 2010). Surprisingly, a similar trend is also observed in MOB only capable of expressing the pMMO, where the highest concentrations of mb in the spent media are observed in cells cultured in media containing low (i.e., less than 1 mM Cu) amended copper (Bandow et al., unpublished results; Choi et al., 2010).

2.2. Separation of mb from whole cells 2.2.1. Method 1: Centrifugation and filtration Initially, mb is separated from whole cells via the following procedure (DiSpirito et al., 1998; Zahn and DiSpirito, 1996). This method is very labor intensive, but can be used in the absence of a tangential flow or hollow fiber filtration system required in method 2.

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pmoA

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40 1

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Figure 17.1 RT-PCR of pmoA (A) and extracellular concentrations of mb (B) from M. trichosporium OB3b cultured in media containing initial amended copper concentrations of 0.03, 0.2, 0.4, 0.7, 1.0, 3.0, 5.0 and 10.0 mM. The fermentor was run as a batch reactor, with 80% of the media replaced every 48 h. Open symbols designate cells expressing the sMMO and closed symbols for cells expressing the pMMO. (C) Spent media from cells cultured in NMS media amended with 0.7 mM CuSO4 (note yellow color from mb). (D) Spent media from cells cultured in NMS media amended with 10 mM CuSO4 (note: loss of color).

1. Cells are centrifuged at 10,000g for 20 min at 4  C to pellet cells. 2. The supernatant is decanted into a flask or centrifuge bottle and centrifuged a second time at 10,000g for 20 min at 4  C. 3. The supernatant is then filtered through a 0.2 mm membrane filter (Gelman Sciences, Inc., Ann Arbor, MI, USA) to remove any residual suspended solids.

2.2.2. Method 2: Tangential flow or hollow fiber filtration Methanobactin is separated from cells in the culture medium directly using a tangential flow or other forms of continuous filtration system. For example, we use a CentramateTM PE tangential flow filtration system (Pall Corporation, Framingham, MA, USA) containing either a OS010C10 Centramate 10,000 Da or a OS030C10 Centramate 30,000 Da molecular mass filter cassette (Choi et al., 2010). The key to using these systems is the use of molecular mass filter and not the 0.2 or 0.45 mM pore sized microbial filters. The microbial filters are designed to filter out low concentrations of cells and clog rapidly when separating high-density cell cultures. The molecular mass filters do not have this problem since the pore size is too small to trap cells and clogging is avoided. An additional value in using the smallest pore

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size possible is the elimination of higher molecular mass material present in cell cultures due to excretion and/or cell lyses. Last, even a small-scale tangential flow filtration system can filter 50–1000 ml/min.

2.3. Concentration of mb from spent media Although the mb concentrations in the spent media can be quite high (Fig. 17.1), the sample requires concentration before additional purification steps are undertaken unless one has access to a high-throughput HPLC system. Initial efforts to concentrate mb was by lyophilizing the spent media (Zahn and DiSpirito, 1996). However, lyophilization also concentrates media salts and other trace contaminants, which can cause problems in subsequent purification steps. 2.3.1. Method 1: Sep-Pak columns If the sample volume is small, the filtrate from Section 2.2.1 or 2.2.2 can be loaded onto reversed-phase C18 solid-phase extraction (SPE) Sep-Pak cartridges (Waters Corp., Milford, MA, USA). Prior to loading, the cartridges should be conditioned sequentially with 3 ml methanol, 3 ml 60% acetonitrile, 3 ml methanol, and then 6 ml H2O. The sample can then be added via a syringe or a syringe coupled to a peristaltic pump until a brown band accumulated at the front end of the cartridge (Sulpeco, Bellefonta, PA, USA; Choi et al., 2003; DiSpirito et al., 1998). The bound material is then washed three times with 6 ml H2O and the sample eluted with 60% acetonitrile. The eluant is then frozen by dropping into liquid nitrogen and lyophilized for mb concentration and removal of acetonitrile. Freezing mb by direct addition into liquid nitrogen results in frozen pellets and following freeze-drying cycle results in a yellow to orange powder depending on sample and metal composition. 2.3.2. Method 2: Dianion HP-20 For larger samples it is recommended that the sample be loaded on a Dianion HP20 column (Sulpeco, Bellefonta, PA, USA). Since the sample will be eluted with solvents, the column used should be solvent resistant. In this case, an old-fashioned glass column with a frit bottom is appropriate (Fig. 17.2). Solvent resistant commercial columns are available from most column supply companies, however, they are not necessary since the primary objective is to concentrate mb from the spent media and remove residual salts and the more aqueous components in the spent media. However, it should be noted that if done correctly this simple chromatography step can result in samples that are over 95% mb and the contaminants are often breakdown products of mb (Choi et al., 2006a,b). Dianion HP-20 from the manufacturer should first be activated with methanol for 15 min followed by extensive H2O washes as described on the

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Retentate Filtrate

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Figure 17.2 Schematic diagram of coupled tangential flow filtration of culture media and harvesting of mb from the spent media. X designates peristaltic pump.

product directions sheet and the column poured. If not used immediately, the column can be stored in 60% acetonitrile. Dianion HP20 can also be stored in the column in 60% acetonitrile at room temperature if the column is capped. Before use, the resin should be washed with 5–10 column volumes of H2O. If a peristaltic pump is used to drain the column, fast flow rates can be used without damage to the resin (Fig. 17.2). We typically use a 7  60 cm column with a bed volume of 7  20 cm and have used flow rates of 200 ml/min during loading and washing. Following column washing in H2O, the sample is loaded and eluted as follows: 1. Drain the column to the solvent resin interface. 2. Add spent media, slowly so as to avoid disturbing the resin. 3. Column flow rates can be increased once the sample volume accumulates 10–30 cm above the resin. At this point, flow rates can be increased to over 100 ml/min. 4. Following sample loading, drain column to the solvent resin interface and then add H2O as described in steps 1–3. Wash the sample with three to five column volumes of H2O and leave two to three bed volumes of H2O in the column. 5. Slowly add 60% acetonitrile, 1–3 ml/min, so as not to disturb the H2O– 60% acetonitrile interface. The objective is to maintain a defined interface between the two solvents (Fig. 17.3A). Once the desired interface is set, flow rates can be increased. 6. When the 60% acetonitrile fraction meets the resin solvent interface, slow flow rates to maintain a discrete interface (Fig. 17.3B). The solubility of air in water and acetonitrile differ, and gas bubbles in the column often accumulate and will disturb the resin (Fig. 17.3B). If the flow rate is maintained at a rate slow enough to maintain a tight colored band, for example, 3–8 ml/min, bubble formation will occur behind the eluting mb and the volume of the eluting mb can be kept to a minimum (Fig. 17.3C).

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Figure 17.3 Dianion HP-20 column highlighting the 60% acetonitrile:H2O interface (A), elution of mb and the degassing and column disruption due to degassing by acetonitrile (B), and mb as it elutes from the column (C).

7. The eluted sample is then frozen in liquid nitrogen and freeze-dried as described in Section 2.3.1. 8. Figure 17.2 shows a diagram of how we couple the tangential flow filtration described in Section 2.2.2 with sample collection on Dianion HP-20 column. Coupling the two procedures we can extract mb from 10 to 20 l of culture media and start the freeze-drying of the sample in less than 4 h.

3. Purification of mb Depending on the MOB mb is being extracted from, the sample or the culture conditions, additional purification of mb may be necessary. Sample purity should be checked to determine if additional purification is necessary, as previous studies have shown that this can generate a variety of breakdown products unless mb is complexed with Cu. (Kim et al., 2005). If additional purification is necessary, reverse-phase HPLC chromatography (e.g., Vydac 218 TP1010 (C18) column or Hamilton PRP-3) is suggested. Freeze-dried mb is resuspended in H2O, loaded, and the samples eluted with a H2O:methanol gradient containing 0.001% acetic acid. The HPLC chromatography step described above can improve sample purity of copper containing mb (Cu-mb). However, we have had very little success with the further purification of metal-free mb-OB3b because of the chemical instability of the oxazolone rings. Even concentrations of acetic acid as low as 0.001% result in the breakdown of the oxazolone B-ring during an HPLC chromatography. We have substituted 1 mM NH4-acetate in place of acetic acid but have still not improved on the purity of the metal-free material from the Dianion HP-20 column.

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4. Sample Variability An electrospray ionization time-of-flight (ESI-TOF) mass spectrum of the mb sample from Fig. 17.4 in the negative ion mode is shown in Fig. 17.4 (top). In addition to the expected [MþCuþ3Hþ]2 species at 607.04 1.0 ´ 104 1215.09

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Figure 17.4 ESI-TOF mass spectra of HPLC fractions in Fig. 17.5 of the mb from M. trichosporium OB3b. Top, unfractionated sample, prior to separation via HPLC; middle, fraction 1 from Fig. 17.5; bottom, fraction 2 from Fig. 17.5.

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and the [MþCuþ2Hþ]1 species at 1215.09, a 2 ion peak at 510.62 and a corresponding 1 ion peak at 1084.05 are also observed and have a calculated mass for M that is expected for an mb that is missing its C-terminal methionine (Met7). Figure 17.4 shows that both forms of mb can bind Cu ions. The two forms can be separated by loading on a reverse-phase HPLC column and eluting with an H2O:methanol gradient containing 0.1% acetic acid (Fig. 17.5). ESI-TOF analysis of the two fractions obtained shows the Cu-bound mb that is missing its Met7 elutes first (Fig. 17.4, middle), followed by the Cu-bound full-length mb (Fig. 17.4, bottom). Proton NMR experiments can be used to estimate the ratio of the two forms present in a particular preparation of mb. The ratio for the Lot B sample shown in Figs. 17.4 and 17.5 is of 48% to 52%, respectively. The Lot B sample shown in Figs. 17.4 and 17.5 represents the preparation showing the highest percentage of Cu-mb minus Met7 observed so far and is shown to demonstrate that more than one form of mb can be present in mb samples. The two forms of mb, mb and mb minus Met7, show slightly different spectral properties, which are reflected by the ratio of the absorption at 340 and 390 nm for the metal-free mb. This ratio can be used to estimate the percentage of each species (Fig. 17.6). At this time, we cannot predict whether the presence of multiple forms of mb is unique to the mb from M. trichosporium OB3b or is a common phenomenon.

A280

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Figure 17.5 Reverse-phase HPLC chromatography of mb Lot B from M. trichosporium OB3b after exposure to Cu(II) at a ratio of 0.7 Cu per mb. Before loading, the sample was dissolved in 10 mM phosphate buffer, pH 6.5, and CuSO4 added in 0.1 M increments, adjusting the pH to 6.5 after each addition. The sample was eluted with a 1–99% H2O:methanol gradient containing 0.001% acetic acid at a flow rate of 3 ml/m.

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1.2

Figure 17.6 (A) UV–visible absorption spectra of three different mb preparations from M. trichosporium OB3b. The percentages represent the ratio of metal-free mb to metal-free mb minus Met. (B) The absorption ratios at 340/390 nm of the three different mb preparations.

ACKNOWLEDGMENTS National Science Foundation Grants CHE-10112271 (A. D. S), CHE-085070 (W. H. G. and S. C. H), Department of Energy Grant DE-FC26-05NT42431 (J. D. S), and Grants from the Carl Page Foundation ( J. D. S) are gratefully acknowledged. The 400 MHz Bruker Avance II NMR spectrometer and Agilent 6210 ESI-TOF LC/MS used in these studies were funded with National Science Foundation Grants CHE-0521019 and CHE-0619296 to the UW-Eau Claire.

REFERENCES Behling, L. E., Hartsel, S. C., Lewis, D. E., DiSpirito, A. A., Masterson, L. R., Veglia, G., and Gallagher, W. H. (2008). NMR mass spectroscopy, and chemical evidence reveal a different chemical structure for methanobactin that contains oxazolone rings. J. Am. Chem. Soc. 130, 12604–12605. Choi, D.-W., Kunz, R. C., Boyd, E. S., Semrau, J. D., Antholine, W. E., Han, J.-I., Zahn, J. A., Boyd, J. M., de la Mora, A. M., and DiSpirito, A. A. (2003). The membraneassociated methane monooxygenase (pMMO) and pMMO-NADH:quinone oxidoreductase complex from Methylococcus capsulatus. Bath. J. Bacteriol. 185, 5755–5764. Choi, D. W., Do, Y. S., Zea, C. J., McEllistrem, M. T., Lee, S. W., Semrau, J. D., Pohl, N. L., Kisting, C. J., Scardino, L. L., Hartsel, S. C., Boyd, E. S., Geesey, G. G., et al. (2006a). Spectral and thermodynamic properties of Ag(I), Au(III), Cd(II), Co(II), Fe(III), Hg(II), Mn(II), Ni(II), Pb(II), U(IV), and Zn(II) binding by methanobactin from Methylosinus trichosporium OB3b. J. Inorg. Biochem. 100, 2150–2161.

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Choi, D. W., Zea, C. J., Do, Y. S., Semrau, J. D., Antholine, W. E., Hargrove, M. S., Pohl, N. L., Boyd, E. S., Geesey, G. G., Hartsel, S. C., Shafe, P. H., McEllistrem, M. T., et al. (2006b). Spectral, kinetic, and thermodynamic properties of Cu(I) and Cu(II) binding by methanobactin from Methylosinus trichosporium OB3b. Biochemistry 45, 1442–1453. Choi, D.-W., Semrau, J. D., Antholine, W. E., Hartsel, S. C., Anderson, R. C., Carey, J. N., et al. (2008). Oxidase, superoxide dismutase, and hydrogen peroxide reductase activities of methanobactin from types I and II methanotrophs. J. Inorg. Biochem. 102, 1571–1580. Choi, D. W., Bandow, N., McEllistem, T. M., Semrau, J. D., Antholine, W. E., Hartsel, S. C., Gallagher, W., Zea, C. J., Pohl, N. L., Zahn, J. A., and DiSpirito, A. A. (2010). Spectral and thermodynamic properties of methanobactin from Methylomicrobium album BG8 and Methylococcus capsulatus Bath: A case for copper competition on a molecular level. J. Inorg. Biochem. 104, 1240–1247. DiSpirito, A. A., Zahn, J. A., Graham, D. W., Kim, H. J., Larive, C. K., Derrick, T. S., Cox, C. D., and Taylor, A. (1998). Copper-binding compounds from Methylosinus trichosporium OB3b. J. Bacteriol. 180, 3606–3613. Kim, H. J., Graham, D. W., DiSpirito, A. A., Alterman, M. A., Galeva, N., Larive, C. K., Asunskis, D., and Sherwood, P. M. A. (2004). Methanobactin, a copper-acquisition compound from methane-oxidizing bacteria. Science 305, 1612–1615. Kim, H. J., Galeva, N., Larive, C. K., Alterman, M., and Graham, D. W. (2005). Purification and physical-chemical properties of methanobactin: A chalkophore from Methylosinus trichosporium OB3b. Biochemistry 44, 5140–5148. Krentz, B. D., Mulherron, H. J., Semrau, J. D., DiSpirito, A. A., Bandow, N. L., Haft, D. H., Vuilleumier, S., Murrell, J. C., Mc.Ellistrem, M. T., Hartsel, S. C., and Gallagher, W. H. (2010). A comparison of methanobactins from Methylosinus trichosporium OB3b and Mehylocystis strain SB2 predicts methanobactins are synthesized from diverse ribosomally produced peptide precursors modified to create a common core for binding and reducing copper ions. Biochemistry 49, 10117–10130. Ongena, M., Jacques, P., Thonart, P., Gwose, I., Ferna´ndez, D. U., Scha¨fer, M., and Budzikiewicz, H. (2001). The pyoverdin of Pseudomonas fluorescencs BTP2, a novel structural type. Tetrahedron Lett. 42, 5849–5851. Semrau, J. D., DiSpirio, A. A., and Yoon, S. (2010). Methanotrophs and copper. FEMS Microbiol. Rev. 34, 496–531. Vossen, W., and Taraz, K. (1999). Structure of pyoverdin PVD 2908–A new pyoverdin from Pseudomonas sp. Biometals 12, 323–329. Zahn, J. A., and DiSpirito, A. A. (1996). Membrane-associated methane monooxygenase from Methylococcus capsulatus Bath. J. Bacteriol. 178, 1018–1029.

C H A P T E R

E I G H T E E N

Measurements of Biosphere–Atmosphere Exchange of CH4 in Terrestrial Ecosystems Klaus Butterbach-Bahl,* Ralf Kiese,* and Chunyan Liu† Contents 1. Introduction 2. Chamber Measurements of CH4 Exchange 2.1. Closed chamber technique 2.2. Dynamic chamber technique 2.3. Determination of different emission pathways (plant-mediated transport, bubbles, gas diffusion) 3. Micrometeorological Measurements of CH4 Exchange Acknowledgment References

272 274 274 282 283 284 285 285

Abstract This chapter focuses on methods for measuring CH4 exchange between the biosphere and the atmosphere. In the context of the global importance of the biosphere as a source and a sink of atmospheric CH4, special emphasis is given to details of gas flux measurements. Due to their widespread use and suitability for targeted process studies, chamber techniques as a means for measuring CH4 fluxes are highlighted. Besides detailed recommendations for measurements of fluxes with chambers, potential problems of the chamber technique are also discussed, such as changes in environmental conditions due to chamber installations. Further, a short overview is provided of how different pathways of CH4 exchange, specifically plant-mediated transport, ebullition or diffusion, can be separated and quantified under field conditions. Finally, a short summary of micrometeorological CH4 measuring techniques such as the eddy covariance method is provided. This technique relies on fast-response

* Institute for Meteorology and Climate Research, Atmospheric Environmental Research (IMK-IFU), Karlsruhe Institute of Technology (KIT), Garmisch-Partenkirchen, Germany Institute of Atmospheric Physics, Chinese Academy of Sciences (IAP-CAS), Beijing, China

{

Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00018-8

#

2011 Elsevier Inc. All rights reserved.

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sonic anemometers and CH4 analyzers (
10 Hz) to make direct measurements of the vertical CH4 flux at a point above the vegetation surface.

1. Introduction In the past few decades, the scientific community became increasingly interested in methane (CH4) exchange between terrestrial ecosystems and the atmosphere. This interest is largely driven by the increasing public awareness of global climate change. CH4 contributes at present
18% to the observed global warming (Denman et al., 2007) and, thus, it is the second most important greenhouse gas following CO2. Besides being an important radiatively active atmospheric trace gas, CH4 also plays a key role in atmospheric chemistry by significantly affecting levels of ozone, water vapor, the hydroxyl radical, and numerous other compounds (Wuebbles and Hayhoe, 2002). Atmospheric CH4 concentrations have increased since 1750 from about 700 parts per billion by volume (ppbv ¼ nL L 1), as derived from ice cores, up to a global average concentration of about 1770 ppbv in 2005 (Isaksen et al., 2009). The growth rate in CH4 concentration has changed considerably since the early 1990s from a steady monotonic increase of
15 ppbv year 1 in the later decades of the twentieth century until the early 1990s to values close to zero (Fowler et al., 2009), though most recent measurements show renewed increases from the end of 2006 onwards (Rigby et al., 2008). CH4 exchange between terrestrial ecosystems and the atmosphere is mainly driven by soil microbial processes. Thereby, CH4 exchange is the net result of simultaneously occurring production and consumption processes, either carried out by methanogenic archaea or by methanotrophic bacterial communities (Conrad, 1996). CH4 production by methanogens represents the last step in anaerobic fermentation of organic substrates and, thus, waterlogged soils such as natural wetlands or rice paddies are with
32% or 160 Teragram (Tg ¼ 1012g) CH4 year 1 the main terrestrial sources within the global atmospheric CH4 budget (Fowler et al., 2009; Table 18.1). On the other hand, atmospheric CH4 can also be consumed by methanotrophs. Methanotrophs are ubiquitously found in upland soils, but are also present in wetland ecosystems where they inhabit aerobic microsites and oxic soil layers and oxidize significant amounts of the CH4 produced in anaerobic parts of the soil. Within the global atmospheric budget of CH4 (Table 18.1), uptake of atmospheric CH4 by soils is in a range of 15–45 Tg CH4 year 1, thus representing 3–9% of the total global CH4 sink strength, with the latter being dominated by the reaction of CH4 with OH radicals in the troposphere (Wuebbles and Hayhoe, 2002).

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Table 18.1 Sources and sinks of atmospheric CH4 (adapted from Wuebbles and Hayhoe, 2002)

Source/sink

Natural sources Wetlands Termites Oceans Marine sediments, geological sources, wildfires Foliar CH4 emissions from UV-irradiated pectin (Bloom et al., 2010) Anthropogenic sources Rice cultivation Ruminants Biomass burning Waste disposal Natural gas, coal mining, and other fuel-related sources Total sources Sinks for atmospheric CH4 Removal to the stratosphere Reaction with hydroxyl radicals in the troposphere Uptake by soilsa Total sinks a

Range of Global annual estimates Contribution emission/removal (Tg CH4 to source/sink strength (%) (Tg CH4 year 1) year 1)

100 20 4 21

92–232 2–22 0.2–20 12–48

19.9 4 0.8 4.2

0.6

0.2–1.0

< 0.2

60 81 50 61 106

25–90 65–100 27–80 40–100 46–174

11.9 16.1 9.9 12.1 21

504

410–660

40 445

32–48 7.9 360–530 87.8

22 507

15–45 4.3 430–600

Value for global atmospheric CH4 uptake by soils taken from Dutaur and Verchot (2008).

Recently, also plants themselves have been identified to be potential net emitters of CH4 (Keppler et al., 2006). However, here CH4 is not of microbial origin, but seems to be mainly produced by the UV radiationinfluenced destruction of structural components of plants, such as pectin, lignin, and cellulose (Vigano et al., 2008) as well as of fresh nonstructural photosynthates (Bru¨ggemann et al., 2009). In the original work on CH4 emissions by plants, it was estimated that plant CH4 emissions may account for 10–45% of the global atmospheric methane source strength. Based on new measurements and a reevaluation of available data it, has been concluded recently that plants are not a major source of atmospheric CH4 (Bru¨ggemann et al., 2009; Nisbet et al., 2009) and that the contribution of

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plant CH4 emissions to the global atmospheric CH4 budget may be as low as <0.2% (Bloom et al., 2010). For measurements of CH4 exchange between the biosphere and the atmosphere, one needs to consider that this exchange can occur via different pathways, namely molecular diffusion across the soil air/water–atmosphere boundary, pressurized ventilation (e.g., thermo-osmosis or Knudsen diffusion in porous media), plant-mediated transport of CH4 by aerenchymatic vascular plants or as dissolved CH4 via xylem water, and ebullition of gas bubbles from supersaturated sediments or soils. In view of these different pathways for biosphere–atmosphere CH4 exchange, different measuring techniques are used, which are outlined in the following section. Specific emphasis is given to the closed (nonsteady state) chamber method since this is the common technique applied for measuring CH4 fluxes (Conen and Smith, 1998), while micrometeorological techniques such as the eddy covariance technique are only briefly discussed.

2. Chamber Measurements of CH4 Exchange Chamber-based techniques are the most widespread for measurements of CH4 exchange rates between terrestrial ecosystems and the atmosphere. Two principal approaches are followed here: the closed chamber technique and the dynamic chamber technique, which includes exchange of chamber headspace air at a given flow rate. Both approaches involve the confinement of a surface area—with or without plants or plant parts—to restrict the gas volume and gas exchange and to increase the concentration of the gas in the headspace. The huge advantage of chambers as compared to other measuring techniques such as micrometeorological approaches is their easy deployment, low cost, and operation. Since power requirements are low and there is no strict need for online measurements, closed chamber measurements can be performed in remote regions as well. Further, they do not require large experimental areas and so they are portable and permit process studies and experiments with many treatments (Denmead, 2008).

2.1. Closed chamber technique Closed chambers are the most commonly used technique to measure soil– atmosphere exchange processes. They enclose gas tightly, a defined surface area such as a plant–soil system (e.g., sample plot of a rice field), plant parts (e.g., rice shoot), or bare soil. In the standard configuration, it consists of a frame which is driven into the soil prior to measurements and a top, which can be mounted on the frame in a gas-tight fashion and which encloses an area of typically 0.01–1 m2. Figure 18.1 provides an example of a principal

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A

B

C

D

Figure 18.1 Closed chamber measurements. (A) Insolated, gas-tight closed chamber of stainless steel (0.4  0.4  0.4 m) mounted on a stainless steel frame. Note the thermometer on top of the chamber for measurements of headspace air temperature and at the left-hand side the syringes for gas sampling as well as the battery pack (right-hand side) to power a ventilator inside the chamber to ensure homogenous mixing of the chamber’s headspace air; (B) chamber frame and chamber with rubber sealing to ensure gas-tight mounting; (C) time-dependent sampling of headspace gas concentrations (0, 10, 20, 30, 40 min) by plastic syringes with luer lock closure; (D) field setup of closed chambers to address spatial variability of CH4 fluxes from a feedlot in Inner Mongolia.

closed chamber measuring setup of gas fluxes. In this section, some general rules when using closed chambers are outlined and information on sampling as well as calculation of fluxes is provided. Most of this information also applies for dynamic chamber measurements. 2.1.1. Chamber effects on environmental conditions Putting a chamber on the ground will significantly alter environmental conditions. Consequently, it is likely that the flux that is intended to be measured will be somewhat affected by the chamber (Rochette and Eriksen-Hamel, 2008). Therefore, several precautions should be taken to minimize the chamber bias on measurements of biosphere–atmosphere exchange processes. Key issues are (a) soil disturbance due to installation and maintenance; (b) temperature, pressure, and humidity perturbations; (c) gas mixing; and (d) spatial and temporal variability of fluxes.

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2.1.1.1. Soil disturbance For ensuring gas-tightness to the ground, frames or anchors, on which the chambers are later mounted, are used. They may be driven 5–15 cm in the soil, depending on site conditions and requirements. However, during installation roots may get cut, affecting substrate availability for the soil microbial C and N turnover and consequently also the associated exchange of C and N trace gases (Keller et al., 2000). Moreover, if chambers are installed on top of the frame, air and consequently also soil temperatures may increase and rainfall may be missed. To minimize disturbance, closure times should be minimized and alternative positions for chambers should be available, such as having 2–3 more frames than chambers to minimize bias from long-term measurements of fluxes across a season or during a multiyear study (see e.g., Kiese et al., 2003). Further, soil compaction by trampling in the vicinity of the chambers needs to be minimized. For instance, chamber measurements of CH4 fluxes from rice paddies may get severely biased by approaching the chambers for gas sampling, since vibrations may force bubble emissions from the sediment. Boardwalks can significantly reduce this problem. 2.1.1.2. Temperature, pressure, and humidity perturbations Once closed, the headspace temperature of translucent chambers may increase by >15–20  C and air humidity quickly approaches saturation. Such increases affect biological activity, physical absorption, or dissolution of dissolved gases and the dilution of the gas of interest due to increased water vapor concentrations. Therefore, one should use opaque, insulated chambers. This is not always possible since translucent chambers have to be used for long-term installations and for running automated closed chamber systems enclosing plants (Holst et al., 2007). Again, keep closure times of chambers as short as possible to minimize the bias. Due to changes in the magnitude of fluxes, sampling closure time may be adapted across seasons. Pressure perturbations can also occur. Specifically, mounting of the chambers, wind effects, and gas sampling can affect soil–plant gas exchange. Even though pressure changes may only be in the range of a few pascals, these changes can significantly alter fluxes due to disturbance of the natural gradient, induced mass flow by sampling or release of gas bubbles from supersaturated sediments. The pressure effect of chamber mounting can be reduced if rubber seals are used instead of water sealing. For avoiding unintended mass flow due to gas sampling, chambers should have a vent, the dimension of which depends on wind speed and chamber volume (Hutchinson and Mosier, 1981). An effective vent design to ensure chamber pressure equilibrium has been introduced by Xu et al. (2006). 2.1.1.3. Gas mixing Molecular diffusion may be sufficiently rapid within the chamber headspace if chambers are <15 cm in height and do not enclose large amounts of vegetation. However, the latter is often the case

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if chamber-based fluxes are carried out in wetlands and rice paddies, where chambers are up to 1 m in height (Wassmann et al., 1996). Here, battery- or power-driven ventilators inside the chambers are needed to ensure homogeneous mixing of headspace during the sampling procedure. 2.1.1.4. Spatiotemporal variations Due to the biological nature of production and consumption processes underlying the net exchange of CH4 at the biosphere–atmosphere boundary, fluxes tend to be notoriously variable on spatial and temporal scales. For example, typical coefficients of variation for CH4 uptake by temperate and tropical forest or grassland soils range from 16 to 130 (Butterbach-Bahl et al., 2002; Kiese et al., 2003; Liu et al., 2009), with a median value of
25–40. To address the issue of site variability, multichamber measurements are needed. Davidson et al. (2002) reported that the means of eight randomly chosen flux measurements of soil respiration from a population of 36 measurements made with 300 cm2 diameter chambers in tropical forests and pastures were within 25% of the full population mean 98% of the time and were within 10% of the full population mean 70% of the time. Rates of CH4 exchange can easily vary over several magnitudes from microgram CH4–Carbon (C) m 2 h 1 to milligram CH4–C m 2 h 1 during the course of a year, and periods with net uptake of CH4 can be followed by net-emission periods (Butterbach-Bahl et al., 1997; Fiedler et al. 2005). Calculations of full annual budgets of CH4 exchange therefore require that measurements span at least an entire year or even multiyears to address interannual variability as driven by meteorological conditions or interannual and seasonal differences in depths of the groundwater table at wetland sites. For ecosystems with expected high net emissions of CH4 such as wetlands or rice paddies, subdaily (multiple times a day) measurements are recommended. Here diurnal variations, as driven by changes in topsoil temperature, can often reach a factor of two (Butterbach-Bahl et al., 1997). Subdaily flux measurements can be done either by automated chamber systems (Butterbach-Bahl et al., 1997) or if terrain or field sizes allow for it, by micrometeorological approaches (see below, Long et al., 2010). For systems being predominantly sinks for atmospheric CH4 such as steppe upland soils, weekly measurements using manual chambers and syringe sampling may prove sufficient to calculate annual budgets (Chen et al., 2010). Large variations in CH4 fluxes can also be induced by human management (e.g., tillage, fertilizer application, irrigation, and drainage) or changes in environmental conditions (air pressure changes, rainfall, snow melting, thawing; Rinne et al., 2008), so that the timing and frequency of measurements need to be adapted adequately.

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2.1.2. Chamber design Precaution needs to be taken that flux chambers are fabricated of nonreactive materials (stainless steel, aluminum, PVC, polypropylene, polyethylene, or plexiglass). To minimize temperature increases inside the chambers, they should be white, coated with reflective material, or insulated using, for example, styrofoam, at the outer side of the chamber. If automated closed chamber sampling systems are used and plants are enclosed, plexiglass chambers are needed, since such systems stay installed at a given plot for at least several days. In this case, lids need to open completely to allow rainfall to enter the area covered by the chamber (Barton et al., 2008). Further, chamber opening should be programmed in such a way that opening is ensured if the air temperature in the chamber exceeds a set threshold value (e.g., 40  C) or if rain is falling with a significant intensity (e.g., 5 mm/ 10 min; Barton et al., 2008). Areas covered by chambers should not be <175–200 cm2, or else insertion of frames will lead to compaction of soils, which is likely to affect exchange rates too. Typically, chambers cover areas of 0.1–1 m2. It should also be considered that the relative error in flux estimates associated with a poor chamber seal increases with decreasing chamber diameter since the flux is proportional to the source area while the gas leak is proportional to the perimeter (Table 18.2). Rochette and Eriksen-Hamel (2008) estimated that chamber area to perimeter ratios 10 cm (
cylindrical chamber with a diameter of 40 cm) are most suitable. No conclusion can be drawn at present if rectangular or cylindrical chamber designs are superior. Chambers should have a vent tube, preferably as described by Xu et al. (2006) in order to avoid pressure effects—that is, decreasing by gas sampling—on rates of gas exchange. Further, a port for gas sampling is needed such as a butyl rubber stopper or stopcock. It is also worthwhile to note that, for a given flux, the increase/decrease of the headspace CH4 mixing ratio is directly proportional to the height of the chambers: increasing the height of a chamber decreases the detection limit. 2.1.3. Sampling procedure Gas samples from the chambers’ headspace can be withdrawn by different means. Typically, gas-tight syringes or preevacuated vials are used. For flux rate calculations, a minimum of four gas samples should be taken, sequentially from time zero to 30–60 min (e.g., in intervals of 10 min) depending on the expected magnitude of fluxes. The gas volume taken will depend on the requirements for analytics. For CH4-concentration measurements in air samples, still gas-chromatographs equipped with a flame ionization detector (Butterbach-Bahl et al., 1997) are mostly used. Typical injection volumes on a packed mole sieve column such as Hayesep Q (3 m, 1/800 , 60/80 mesh) are 0.5–2 ml, but additional sample air (10–30 ml) is usually needed for flushing sample ports or for multigas analyses, including CO2 and N2O). However,

Table 18.2 Rating of closed chamber measurements based on design of chambers, calculation of fluxes, quality control measures, and gas sampling strategy Chamber characteristics

Unit

Good

Recommended

min days

< 10 <5 < 2.5 1 > 60 > 14

Base þ chamber Yes Yes Yes Yes Yes Yes Yes Yes Yes Extainers, vacutainers, on side analysis 10–40 Flexible height 5–20 (depend on soil type, roots) 2.5 –10 > 10 4 >4 20–60 < 20 3–10 Subdaily

days days days

>2 >2 > 30

1 2 5–30

Type of chamber Insulation Vent Pressurized sample (fixed volume containers only) Quality control sample Time zero sample Nonlinear model considered Zero slope tested Temperature corrections Pressure corrections Type of sample vial Height of chamber (depend on type of vegetation) Chamber base insertion depth Area/perimeter ratio Number of samples for calculating fluxes Duration of deployment Temporal resolution of measurement Duration of sample storage prior to analysis Plastic syringes Glass syringes Others

Poor

Push-in No No No No No No No No No Plastic syringe
< 0.5 <1 <5

The rating was based on a detailed evaluation of the importance of each characteristic in the measurement error and on the authors’ own judgment based on long-term experiences in closed chamber measurements (table modified following Rochette and Eriksen-Hamel, 2008).

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CH4 concentration changes in the headspace may also be measured directly at the site using on-site gas chromatography when using automated chamber systems (Breuer et al., 2000; Kiese and Butterbach-Bahl 2002) or portable analytical instruments such as infrared sensors or photoacoustic spectroscopes. For the latter, cross-sensitivities with water vapor and other gas compounds such as CO2 need always to be checked carefully by multipoint and multi-gas calibration procedures (i.e., various concentrations of CH4 water vapor and CO2) while using such instrumentation (Neftel et al., 2006). If standard gas sampling by vials or syringes is done, vials should be pressurized, for example, by injection of 20 ml gas volume in a 10 ml vial. Mixing of headspace gas by pumping the syringe before sampling should be avoided as this may cause pressure perturbations and/or excess dilution of headspace gas by entry of outside air through the vent tube (Hutchinson and Mosier, 1981). Note that withdrawal of gas from the headspace will create an influx of gas from the ambient air. The resulting dilution may need to be considered if the gas sampling volume to chamber volume ratio is >0.01. Moreover, quality control samples should be taken, such as gas samples from a calibration standard carried to the field. The control sample is treated and stored in the same manner as the other headspace gas samples (Table 18.2; Rochette and Eriksen-Hamel, 2008). Generally, it is recommended to analyze gas samples as quickly as possible, preferably within a few hours following sampling. Sampling time is another critical issue. Generally, CH4 fluxes do vary diurnally and seasonally due to changes in environmental conditions, including temperature, soil moisture, groundwater level, substrate supply, etc. Sampling needs to consider these variations and an adequate sampling strategy needs to be developed to address temporal variations. For estimating cumulative seasonal CH4 fluxes from rice paddies, Buendia et al. (1997) suggested that sampling at 06:00, 12:00, and 18:00 h is sufficient to capture most of the diurnal variation observed throughout the rice growing season and that sampling is most important where fluxes are most variable, between flowering and harvest. For a typical 100-day rice growing season, fluxes should ideally be measured at around 10, 20, 30, 50, 70, 77, 84, 91, and 98 days after planting. However, this is a rule of thumb which needs to be adapted to the site specific situation. 2.1.4. Flux calculations It should be noted that the CH4 flux at the soil surface is the result of simultaneous microbial production and consumption processes (Conrad, 1996) as well as gas diffusion or gas transport (e.g., thermo-osmosis), so that any major change in headspace gas concentrations is likely to feedback on diffusion gradients as well as soil processes and, thus, on net fluxes at the biosphere–atmosphere interface. Fluxes are calculated from the temporal change in gas concentrations in the chambers’ headspace. This change can be linear or nonlinear. Nonlinearity is a result of changes in the headspace

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CH4-concentration in the closed chamber [ppmv]

1.80

Atmospheric background

1.75

1.70

1.65

1.60 0

10

20

30

40

50

60

Time since chamber closure [min]

Figure 18.2 Typical example of temporal changes in headspace CH4 concentrations in a closed chamber at a temperate forest site where the soil predominantly functions as a sink for atmospheric CH4 (for details, see Butterbach-Bahl et al., 2002).

CH4 concentration with time so that in a net-emission situation the gradient between soil air CH4 concentration and headspace concentration will decrease, which results in a decrease of the flux either due to the diminishing gas-concentration gradient (Davidson et al., 2002) or due to stimulated microbial CH4 uptake (if the soil is partially oxic). For sites where the soils are net sinks for atmospheric CH4, decreasing headspace CH4 concentrations reduce the diffusive flux of CH4 into the soil, which affects the rate of CH4 uptake (Butterbach-Bahl et al., 2002; Fig. 18.2). Thus, the validity of using a linear approach for the calculation of CH4 fluxes needs to be proven by using the goodness of fit r2 values while comparing different calculation methods. It should be noted that use of linear fits as bases of flux calculations may result in an underestimation of real fluxes, so that the use of nonlinear regressions is recommended if temporal changes in headspace concentrations deviate from assumed linearity (Hutchinson and Mosier, 1981; Kroon et al., 2008; Pederson et al., 2001). Kroon et al. (2008) pointed out that there can be several reasons for still using linear regression methods: (a) linearity is assumed in the case of short measurement times, (b) linear regression method is much easier in use than all other methods, and (c) the uncertainty due to spatial and temporal variation in fluxes is assumed to be much larger than the biases due to linear regression. What one finally derives from linear or nonlinear regression is an estimate for the slope of headspace gas concentration changes over time

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(note: test for zero slope is recommended). This slope can then be used in the following formula to calculate the flux rate:
f CH4 mg C m2 h1
chamber volðm3 Þmol weight g mol1 slopeðppmv min1 Þ
¼ 60  106 chamber areaðm2 Þmol volume of CH4 m3 mol1 with 60 and 106 being constants used for converting minutes into hours and grams into micrograms, respectively. Considering the ideal gas law, the mol volume needs to be corrected for air pressure and temperature at the time of measurement:
273:15 þ Tempð CÞ mol volume m3 mol1 ¼ 0:02241 273:15 Atmospheric pressure at measurementðPaÞ  Atmospheric pressure at sea levelðPaÞ This indicates that necessary auxiliary measurements should always include headspace air temperature measurements as well as measurements of the atmospheric pressure. The latter can be also estimated from height above sea level inserted in the barometric formula. Other parameters always needed for interpretation of flux data are time courses of weather data (rainfall, air temperature, relative humidity, solar radiation), soil temperature, soil water content and possibly also soil nutrient concentrations (inorganic N, dissolved organic carbon (DOC)). Site characteristics should be noted, and specifically soil properties such as bulk density, texture, organic C and N concentrations, if possible, for all chamber positions. For all measuring systems, the accuracy of the determination of CH4 concentrations in a gas sample, that is, the closeness of agreement between a test result and the accepted reference value, should be determined. The detection limit, which provides an estimate of the lowest detectable flux, can be approximated by three times the accuracy. If a flux is less than the detection limit or the slope is not significantly different from zero, flux values should be set to zero.

2.2. Dynamic chamber technique Flux measurement with dynamic chambers or flow-through chambers refers to a system where the headspace air of a chamber is constantly exchanged at a given flow rate. From the concentration difference of CH4 in the incoming air and in the outflowing air, the gas exchange rate, and the area covered by the chamber, the CH4 flux can be calculated:

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f CH4 mg CH4  C m2 h1 ¼
mass flow m3 h1  ðCH4 in  CH4 out ÞðppmvÞ  mol weightðg mol1 Þ
chamber areaðm2 Þ  mol volume of CH4 m3 mol1 Again, the mol volume needs to be corrected for pressure and temperature effects (see above). Compared to static chambers, dynamic chambers have the advantage that the increase in CH4 concentrations above the atmospheric background can be controlled by the rate of gas exchange. This reduces the risk of altering CH4 fluxes from the plant–soil systems due to too high or too low CH4 concentrations in the headspace. However, when fluxes are small, the use of dynamic chambers can be limited by the small magnitude of the concentration changes (Denmead, 2008). Moreover, dynamic chamber measurements applied for flux measurements of CH4 are scarce because measuring system requirements such as pumps and flow controllers are expensive and demanding with regard to requirements for power supply and maintenance.

2.3. Determination of different emission pathways (plant-mediated transport, bubbles, gas diffusion) Most vascular wetland plants have developed an extensive aerenchyma system to provide their submerged root system with O2 for respiration. However, since in wetland soils gas concentrations of several gases such as CO2 and CH4 exceed atmospheric concentrations, gas transport in the inverse direction occurs from the soil to the atmosphere. This gas transport can be pure diffusion or in addition supported by pressurized gas flow (Colmer, 2003) due to thermo-osmosis or driven exchange (Schro¨der et al., 1986). In rice paddies and also in natural wetland ecosystems, plantmediated transport of CH4 from the soil to the atmosphere can be the major emission pathway. In Italian rice fields, the aerenchyma transport contributed 88–90% of the overall emission throughout the reproductive and ripening stage (Butterbach-Bahl et al., 1997) whereas the relative contribution of plant-mediated transfer was much lower under high organic inputs to rice paddies (Wassmann et al., 1996). For the determination of the contribution of different emission pathways of CH4 from the soil to the atmosphere in the field, including release of gas bubbles, diffusion through the floodwater column, and plant-mediated transport, Butterbach-Bahl et al. (1997) used a static two-chamber system. The upper chamber enclosed all aboveground plant parts and was sealed to the lower chamber, enclosing the area around the rice tillers, by the floodwater. Due to the exclusion of plant parts in the lower chamber, changes in the headspace CH4 concentration are only due to ebullition or

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diffusion via the water column. Ebullition fluxes can be measured by placing small water filled plexiglass chambers or funnels upside down on the soil surface. Gas bubbles get trapped at the top end of the sealed funnel and by determination of the amount and concentration of the trapped gas, bubble emissions can be estimated. The concentration of dissolved CH4 in soil pore water or flood water may be measured directly using a membrane inlet probe connected to a quadrupole mass spectrometer (Benstead and Lloyd, 1994) or by analyzing headspace CH4 concentrations following the gas equilibration of the water- and gas-phase of water samples in a gas-tight vessel (Wassmann et al., 1996).

3. Micrometeorological Measurements of CH4 Exchange Micrometeorological methods can be used to quantify the turbulent gas exchange between the biosphere and the atmosphere. For eddy covariance methods, the gas concentration and the vertical wind speed are determined simultaneously and at high temporal resolution (10–20 Hz) at a sampling point above the canopy, for example, in flat, homogenous terrain over grassland at 2 m height, thereby integrating fluxes for an area being
100 times greater than the height of the sensors above the canopy. Indirect methods such as the aerodynamic and Bowen ratio methods are based on quantifying the rate of diffusion along a concentration gradient. For many years, instrumental requirements such as fast-response sensors, high accuracy of measurements especially for CH4 and high power demands have hindered widespread applications of micrometeorological techniques for measurements of CH4 exchange. With recent developments in sensor techniques such as an open path CH4 sensor with low power demand and improved suitability and robustness of tunable laser instruments for field use, micrometeorological measurements of CH4 exchange over wetlands or rice paddies have increased (Rinne et al., 2007). Micrometeorological measurements are particularly valuable for heterogeneous ecosystems such as peat wetlands, where compaction by walking to a flux chamber may release gases into the chamber (Fowler et al., 2009). However, due to the integration of fluxes over areas of at least 1 ha, studies of treatment effects or process studies are difficult to realize. For further details on the micrometeorological measurement techniques for quantifying biosphere–atmosphere exchange of trace gases see Moncrieff et al. (1997) or Denmead (2008).

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ACKNOWLEDGMENT This work is a contribution to COST ES0804 “Advancing the integrated monitoring of trace gas exchange between biosphere and atmosphere”.

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Author Index

A Abdallah, M. A., 230, 233 Abecasis, G., 92 Abing, A. M., 41 Aboushadi, N., 92 Abruna, H. D., 249 Aebersold, R., 71, 179–180 Agarwal, S. K., 229, 240 Ahmad, A., 243 Akashi, H., 21 Alben, J. O., 190 Alberto, M. C. R., 277, 283–284 Alifano, P., 100 Ali, H., 106, 111, 123, 130 Ali, M. H., 32, 100, 136 Allen, E. E., 82, 87, 150 Allison, M. J., 3, 7 Alterman, M. A., 228–229, 232–233, 236–238, 248, 260–261, 265 Alves, A. M. C. R., 3, 7 Amann, R., 41, 87 Ambus, P., 272, 284 Amzel, L. M., 75 Anderson, D. J., 276, 278 Anderson, I. J., 82 Anderson, R. C., 229, 232, 248, 261 Andersson, K. K., 178, 188, 199 Andreev, L. V., 55 Angelskar, S. M., 171–172 Ansorge, W. J., 85, 90 Antholine, W. E., 178, 188, 196, 229, 232, 248, 260–263 Anzali, S., 16 Arah, J. R. M., 277, 280, 283–284 Aravind, L., 67 Argandona, M., 17 Arnold, R. G., 229, 240 Arp, D. J., 9 Asunskis, D., 228–229, 236, 248, 260 Atkinson, R. A., 233 Auman, A. J., 36–37 Aurela, M., 284 Ayres, E. K., 122 B Baani, M., 32, 121 Baba, T., 203, 219, 222–223 Babu, M. M., 67

Bagstevold, J. I., 63 Bagyan, I., 27 Baik, M.-H., 46 Bailey, M. J., 101 Baker, D. R., 157 Balasubramanian, R., 178, 195–197, 199, 201–203, 207–208, 212–213, 228–229, 232 Baltschevsky, H., 10 Baltschevsky, M., 10 Bandow, N. L., 229, 233, 248, 259–262 Banfield, J. F., 82, 87, 150 Bapteste, E., 7, 9–10 Baptista, J. C., 239–240 Baranyi, C., 46, 48, 56, 100 Ba¨r Gillisen, M.-J., 52 Barker, P. D., 76 Barnes, C. L., 233 Barns, S. M., 41 Barry, K. W., 83–90, 94–95 Barton, L., 278 Basle, A., 229, 233 Basu, P., 178, 188, 199 Bateman, A., 230, 233 Bates, D. M., 93 Bauer, M., 87 Bazylinski, D. A., 51 Beanan, M. J., 74 Beck, D. A. C., 22, 81, 99, 112 Becker-Birck, J., 151 Becking, J., 121 Beeson, K., 82, 89 Behling, L. A., 229–230, 248 Behling, L. E., 259–260 Belova, S. E., 32, 36–37, 46, 52, 54, 120 Bendall, D. S., 273 Bendinger, B., 46, 48, 56, 100 Benestad, R., 272 Benstead, J., 284 Berger, U., 50, 54 Bernard, L., 151 Bernard, T., 16 Bernhard, M., 17 Berntsen, T. K., 272 Berson, O., 106 Berven, F. S., 63, 66–70, 167, 169, 171–172 Beschastny, A. P., 3–4, 6, 8–10 Bhavsar, J., 82 Bidigare, R. R., 51 Bird, C. W., 64

289

290 Birkeland, N. K., 32, 47, 56, 100 Bisgaard, M., 55 Black, C. C., 3, 7 Blanco, C., 16 Blatny, J. M., 140 Blazzyk, J. L., 137 Bloom, A. A., 273–274 Blytt, H. J., 2, 7 Bodelier, P. L. E., 32, 52, 54 Bodrossy, L., 35, 47, 55–56, 64, 83, 168 Boetius, A., 46 Boffelli, D., 82 Bolstad, B., 93 Bonner, C. A., 21–22, 26 Borges, N., 17 Borken, W., 48 Bork, P., 82, 88 Borodina, E., 143 Borovik, A. S., 201, 207 Bosch, G., 94 Bottomley, P. J., 9 Bourne, D. G., 36 Bourneman, J., 136 Bourque, D., 36 Bousquet, P., 272 Bowerman, S., 157 Bowin, S., 3, 7, 10 Bowman, J. P., 33, 45, 47, 49–52, 56 Boyd, E. S., 178, 188, 239, 248–249, 260–261, 263 Boyd, J. M., 178, 188, 263 Bradburne, J. A., 174 Brandstetter, H., 137 Brantley, S. L., 47 Brasseur, G., 272 Brass, M., 273 Brautaset, T., 140 Brazeau, B. J., 137 Bremer, E., 16–17, 20–21 Brenner, A. J., 206 Breuer, L., 280 Brinkac, L. M., 74 Briones, J. A. M., 42 Briones, M. J., 83 Bruchhaus, I., 3, 7 Bru¨ggemann, N., 273, 276–277 Brunak, S., 67 Bruseth, L. J., 4, 9, 46, 64, 74, 171–172 Brusseau, G. A., 55 Bryan, B., 276 Buck, R., 278 Budzikiewicz, H., 260 Buendia, L. V., 280 Buenger, J., 16 Bueno, C., 277, 283–284 Bumann, D., 174 Burgess, L. W., 161–162, 164 Burrows, K. J., 216–217, 221

Author Index

Bursy, J., 17, 20 Burton, N. P., 140 Busse, H.-J., 48, 54–55 Bussmann, I., 50, 54 Butler, A., 233 Butler, M. K., 32 Butterbach-Bahl, K., 271, 276–278, 280–281, 283 C Cai, T., 277 Camacho-Carvajal, M. M., 71 Canovas, D., 17 Capote, N., 17 Carata, E., 100 Cardy, D. L. N., 136 Carey, J. N., 229, 232, 248, 261 Carey, V. J., 93 Carnal, N. W., 3, 7 Cattoir, H., 55 Cavanaugh, C. M., 243 Cayouette, M., 92 Chain, P. S. G., 9 Chakicherla, A. Y., 9 Chandler, R. E., 50 Chang, H. W., 51, 82, 88 Chan, M. K., 178, 181, 191 Chan, S. I., 177–182, 184–186, 189–191, 196 Chapman, J., 82, 87 Charoensilp, N., 280 Cha, S. K., 249 Chater, K. F., 122 Chen, C.-L., 178–179, 181, 189–191 Chen, K. H.-C., 82, 88, 177–179, 181–182, 184–186, 189–191, 196 Chen, L. L., 65 Chen, P. P.-Y., 178, 181, 191 Chen, W. W., 277 Chen, Y., 35, 48, 83, 100, 120 Chen, Y.-J., 178–182, 184–186, 189–190, 196 Chen, Y.-R., 179–180 Chetina, E. V., 3 Chi, A., 7 Chidthaisong, A., 52, 54 Chidthaisong, G., 272 Chi Fru, E., 239–240 Chisholm, S. W., 90 Chistoserdov, A. Y., 101, 106 Chistoserdova, L. V., 81, 83–84, 90, 94, 106, 112, 151–152, 156–159 Choi, D. W., 178, 188, 196, 229–230, 232, 239, 248–249, 259–263 Choi, J. H., 174 Chomczynski, P., 4 Chou, H. H., 85 Christensen, H., 55 Christgen, B., 239–240

291

Author Index

Christoffersen, L., 51 Chueng, C. Y., 151 Ciais, P. M., 272 Ciccioli, P., 284 Cimbleris, A., 151 Clark, W. A., 49 Colby, J., 120, 136 Cole, J. J., 151 Coleman, M. L., 90 Collins, W., 272 Colmer, T. D., 283 Conen, F., 274 Connon, S. A., 51 Conrad, R., 43, 272, 280 Copeland, A. C., 83–90, 94–95 Cormack, B., 110 Cornish, A., 216–217, 221 Cornish, S. L., 32, 36–37, 46, 54, 120 Corton, T. M., 280 Cossart, P., 90 Costello, A. M., 36–37, 64, 101, 106 Courties, C., 151 Cox, C. D., 229, 236, 238, 240, 242, 261 Cox, R. E., 272 Cox, T., 272 Craan, T., 20 Crill, P. M., 273 Crombie, A., 119 Crosa, J. H., 255 Csaki, R., 64 Cunnold, D. M., 272 Cvejic, J. H., 55, 168 D Daims, H., 46, 48, 56, 100 Dalsoren, S. B., 272 Dalton, H., 64, 136–137, 140, 145–146, 168, 178, 188, 199, 221 Darmaeva, T. D., 16 Daugherty, S. C., 74 Davagnino, J., 101 Davey, J. F., 64 Davidson, E. A., 277, 281 Davison, J., 122 de Beer, D., 32 De Bellis, L., 100 de Bont, J. A. M., 216 Deboy, R. T., 74 Dedysh, S. N., 31–33, 36–37, 41, 46–47, 50, 52, 54, 100, 120, 136 Dege, J. E., 137 Degelmann, D. M., 48 de Jonckheere, J., 2–3, 7 de la Mora, A. M., 178, 188, 263 De Ley, J., 55 Del Giorgio, P. A., 151 Dell, A., 230, 233 Delong, E. F., 90

Demange, P., 230, 233 Denart, M., 3, 7 Denman, K. L., 272 Denmead, O. T., 274, 283–284 Dennison, C., 229, 233, 239–240 Derrick, T. S., 229, 236, 238, 240, 242, 261 De Santis, D., 3, 7 De Stefano, M., 100 Detter, J. C., 82, 88 Dettling, M., 93 Dickinson, D., 272 Dijkhuizen, L., 3, 7 Dimitriov, G., 46 Dimitrov, G., 64, 74 DiSpirito, A. A., 32, 136, 178, 188, 196, 198, 228–231, 233, 236, 238, 240, 242, 247–248, 252–253, 259–263 Djordjevic, G., 92 Doan, P. E., 178, 188, 201 Dodson, R. J., 74 Doig, P., 69 Doronina, N. V., 16 Dorward, D. W., 76 Douffet, T., 277 Dow, C. S., 64 Downie, J. A., 76 Do, Y. S., 239, 248–249, 260–261, 263 Drachuk, S. V., 16 Drake, H. L., 48 Dreis, A. M., 229, 232 Driller, H., 16 Driskell, L. O., 122 Droege, M. W., 213 Drummond, D., 221 D’Souza, M., 21–22, 26 Dudoit, S., 93 Dumont, M. G., 32, 83, 100, 136, 143 Dunfield, P. F., 31–33, 36–37, 46–47, 50, 52, 54, 56, 100, 120–122, 136 Dunn, M. J., 76 Durand, P., 277 Durbin, R., 92 Durkin, A. S., 46, 64, 74 Dutaur, L., 273 Dutilh, B. E., 32 Duyzer, J., 272, 284 E Eberlein, C., 64 Edwards, R., 90 Eguchi, M., 36 Eichacker, L. A., 71–72 Eidhammer, I., 66–70, 167, 169, 172 Eisen, J. A., 74, 82, 89 El Ghazouani, A., 229, 233, 239–240 El Karoui, M., 122 Elliott, S. J., 178, 184–185, 189–190 Ellis, B., 93

292

Author Index

El-Sheikh, A. F., 9 Elvert, M., 55 Encalada, R., 2 Enomoto, T., 7 Erb, T. J., 121 Eriksen-Hamel, N. S., 275, 278–280 Eshinimaev, B. T., 3, 10, 16, 21, 26, 50, 54, 100–102, 113 Ettwig, K. F., 32, 100 Euverink, G. J. W., 3, 7 Exner, M. M., 69 Eyring, V., 272 F Fagerli, H., 272, 284 Falkow, S., 110 Feig, A. L., 179 Fei, Z., 213 Feldman, R. A., 172 Fennell, T., 92 Ferenci, T., 3, 64 Ferguson, S. J., 76 Ferna´ndez, D. U., 260 Fiedler, S., 277 Field, D., 90 Figurski, D. H., 107 Finch, R. J., 101 Firbank, S. J., 229, 233 Fischer, C., 280 Fisher, R., 273 Fitch, M. W., 229, 240 Fjellbirkeland, A., 67–71, 167–172 Flanagan, L. B., 277 Flechard, C., 280 Flikka, K., 66–70, 167, 169, 172 Floss, H. G., 178 Fouts, D. E., 82 Fowle, D. A., 229, 236, 239–241, 243, 248 Fowler, D., 272, 284 Fox, B. G., 136–137, 231–232 Fraser, P. J., 272 Frederick, C. A., 137 Frias-Lopez, J., 90 Friedrich, M. W., 83 Friesner, R. A., 46 Fritsch, E. F., 85, 91, 101, 106 Froland, W. A., 231–232 Fuchs, B., 49 Furtaw, M. D., 276, 278 Furuto, T., 213, 218 Fuse, H., 16, 50, 54, 100–102, 113 Fuzzi, S., 272, 284 G Gaasterland, T., 9–10 Galchenko, F. J., 55

Galeva, N., 228–229, 232–233, 236–238, 260–261, 265 Galinski, E. A., 16–17 Gallagher, W. H., 229–230, 233, 248, 259–262 Gallego-Sala, A. V., 273 Galperin, M. Y., 100 Gao, J., 65 Garcı´a-Estepa, R., 17 Garcia, R. L., 276, 278 Gatter, D., 278 Gauss, M., 272 Gautier, L., 93 Geenevasen, J. A. J., 52 Geesey, G. G., 239, 248–249, 260–261, 263 Gentleman, R. C., 93 Gentry, J., 93 Georgiou, G., 229, 240 Ge, Y., 93 Ghassemian, M., 92 Ghose, T. K., 213, 216, 218 Giffard, P. M., 174 Gilbert, B., 25, 137, 213, 232 Gilbert, J. A., 90 Gilles, V. S., 259 Gilna, P., 90 Giovannoni, S. J., 51 Glo¨ckner, F. O. B., 49, 82, 87 Gloerich, J., 32 Gloux, K., 16 Glukhov, A. S., 22–23, 107–109, 112 Godfrey, P., 174 Godfrey, W., 151 Gogarten, J. P., 65 Go¨ller, K., 17 Gomez-Alvarez, V., 82 Gonzalez, R. A., 51 Goo, J. H., 213, 216 Gordon, P., 9–10 Gorlenko, V. M., 47 Gottschalk, G., 4, 10, 16, 100–102 Gough, J., 67 Gou, Z., 213 Graf, R., 16 Graham, A., 276–277 Graham, D. W., 227–229, 232–234, 236–243, 248, 260–261, 265 Grammel, N., 17 Granier, C., 272, 284 Gray, D. R., 151 Gray, J., 229, 233 Gray, N. D., 239–240 Greco, S., 284 Green, P. N., 56 Green-Tringe, S., 83–90, 94–95 Green, W., 51 Greer, C. W., 36 Griffiths, R. I., 101

293

Author Index

Grigoriev, I., 83–90, 94–95 Groleau, D., 36 Grosse, W., 283 Grove, A., 22–23 Guckert, J. B., 55 Guenter, A., 83 Gulledge, J., 243 Guo, X., 113 Gupta, R., 201, 207 ¨ ., 56 Gustafsson, O Guzev, V. S., 47 Gwose, I., 260 H Haapanala, S., 284 Hackett, M., 94 Hafner, L. M., 174 Haft, D. H., 229, 233, 260 Hakemian, A. S., 136, 196, 201, 207, 212, 223, 239 Hales, B. J., 180 Halpern, A. L., 82, 89 Hamilton, J. T. G., 273 Hanada, S., 36 Hanafusa, H., 172 Han, B., 213 Hancock, R. E., 69 Hanczar, T., 64 Handelsman, J., 150 Handsaker, B., 92 Han, J. I., 178, 188, 263 Hanna, M. L., 213 Han, S., 277 Hansen, E. J., 122 Hanson, R. S., 55, 83, 136, 228 Hanson, T. E., 83, 136, 228 Han, X., 276–277 Hardt, M., 50, 54 Hargrove, M. S., 260–261, 263 Harhangi, H. R., 32, 47, 51, 56, 100 Harris, E. D., 206 Hartman, P. A., 3, 7 Hartsel, S. C., 229–230, 232–233, 239, 248–249, 259–263 Haugen, K., 140 Hauglustaine, C., 272 Hauser, L. J., 9 Haveman, S. A., 74 Hayes, K. F., 248 Hayhoe, K., 272–273 Hedman, B., 189–190 Heidelberg, J. F., 74, 82 Heidelberg, K. B., 82, 89 Heidrich, G., 27 Heijmans, K., 47, 51, 100 Heine, A., 20 Heinonen, J. K., 6

Heinze, E., 272 Heipieper, H. J., 64 Hektor, H. J., 3 Helinski, D. R., 107 Hemme, C. L., 74 Hemscheidt, T., 51 Hensel, R., 7 Hensen, A., 281 Hessels, G. I., 3 Hestnes, A. G., 50 Hewett, B., 276–277 Heyer, J., 50, 54 Higgins, I. J., 216–217, 221 Himes, R. A., 196 Hinz, C., 278 Hiramoto, M., 24, 26 Hirayama, H., 55 Hirsch, P., 48 Hoang, T. T., 122 Hochstein, L. I., 64, 168 Hodgson, K. O., 189–190 Hoffman, B. M., 178, 188, 196, 201, 207, 212, 223, 239 Hoffman, J. M., 82, 89 Holland, E., 272 Ho¨ll, B. S., 277 Holmes, A. J., 36–37, 101, 243 Holmes, M. H., 85 Holst, J., 276–277 Holzinger, R., 273 Homer, N., 92 Hommes, N. G., 9 Hondmann, D., 3 Honkasalo, S. H., 6 Hoppe, H., 151 Hordijk, K., 52 Horikoshi, K., 55 Hornibrook, E. R. C., 273 Hornik, K., 93 Horton, R. M., 140 Ho, S. N., 140 Hossain, M. B., 233 Hothorn, T., 93 Hou, S., 47, 50, 100 House, C. H., 47 Hristova, K. R., 9 Huang, D.-S., 178–179, 181–182, 184–186, 189–190, 196 Huang, J., 272 Huang, Y., 90 Hugenholtz, P., 82, 87–88, 150 Hughes, M. N., 228 Huntemann, M., 82 Hunt, H. D., 140 Huson, D. H., 90 Hutchinson, G. L., 276, 280–281 Hynes, A., 32, 47, 56

294

Author Index I

Iglesias-Guerra, F., 17 Iguchi, H., 52 Im, J., 32, 120 Imperi, F., 233 Ineson, P., 83–84 Isaksen, I. S. A., 272 Ishiguro, K., 151 Islam, T., 32, 47, 56, 100 Iturriaga, R., 151 Ivanova, N. N., 82–90, 92, 94–95 J Jackson, K. J., 213 Jacob, D., 272 Jacobs, S. J., 180 Jacobs, T., 3, 7 Jacques, P., 260 Jahnke, L. L., 55, 64, 168 Jasin, M., 122 Jasso-Chavez, R., 2 Javellana, J. M., 280 Jebbar, M., 16 Jensen, H. B., 63–64, 66–71, 74, 76, 167–173, 175 Jensen, R. A., 21–22, 26 Jensen, S., 100 Jeskulke, K., 151 Jetten, M. S. M., 32, 47, 51, 56, 100 Jiang, H., 100, 213 Jiang, L., 46, 64, 74 Jiang, P., 100 Joergensen, C., 69, 71, 168, 170–171 Joergensen, L., 168 Joint, I., 90 Jollie, D. R., 231–232 Jongejan, P. A. C., 281 Juncker, A. S., 67 Jungblut, P. R., 174 K Kahne, D., 168 Kalyuzhnaya, M. G., 4, 10, 16, 22–23, 27, 50, 54, 81, 83–90, 94–95, 99–102, 107–109, 112–113, 149, 151–152, 156–159 Kamachi, T., 203, 219, 221–223 Kamagata, Y., 36 Ka¨mpfer, P., 49 Kanagawa, T., 36 Kane, S. R., 9 Kang, K. H., 46, 64, 74 Kao, W.-C., 179–180 Kappelmeyer, U., 64 Karlin, K. D., 196 Karlsen, O. A., 63–64, 67–70, 74, 76, 167, 169, 171–173, 175

Karrasch, B., 151 Kato, K., 64, 168 Katterle, B., 178, 188, 199 Keeling, P. J., 3 Keller, M. A. M., 276 Keller, U., 17 Kempf, B., 16 Kemp, R. G., 7 Kenna, E. M., 36, 101 Kenseth, E. M., 229, 232 Kenward, P. A., 239–240 Keppler, F., 273 Ke, S.-C., 190 Khmelenina, V. I., 10, 50, 54 Khmelenina, V. N., 1, 3–4, 6, 8–10, 15–18, 20–27, 33, 41, 47, 50, 52, 54, 100–102, 107–109, 112, 114 Khouri, H., 46, 64, 74 Khunajakr, N., 24, 26 Kieffer, B., 233 Kiese, R., 271, 276–278, 280 Kim, C., 121 Kim, H. G., 213, 216 Kim, H. J., 227–229, 232–234, 236–238, 240, 242, 248, 260–261, 265 Kim, S. W., 213, 216 Kim, Y. M., 213, 216 Kindingstad, L., 171–172 King, T. S., 221 Kip, N., 47, 56 Kisting, C. J., 239, 248–249, 260–261 Kitaoka, S., 7 Kleivdal, H., 69, 71, 168, 170–171 Klemme, J.-H., 7 Klenk, H.-P., 7, 27 Klimont, Z., 272 Klotz, M. G., 9 Knapp, C. W., 229, 233, 236, 239–241, 243, 248 Knief, C., 32, 43, 100 Knittel, K., 46, 49 Knosp, O., 233 Knudsen, S., 113 Kobayashi, A., 82, 88 Kolb, S., 43, 48 Kolonay, J. F., 74 Ko¨mpfer, P., 48, 54–55 Kondapalli, K. C., 196, 201, 207, 212, 223, 239 Konopka, M. C., 149, 159 Koren, S., 85 Kosmach, D., 56 Kovacs, K. L., 64, 168 Kova, K. L., 55 Ko, W.-Y., 21 Kraemer, S. M., 247, 249, 252–253 Krentz, B. D., 229, 233, 260 Krogh, A., 67 Kroon, P. S., 281 Krummel, P. B., 272

295

Author Index

Kuhlmann, A. U., 17, 21 Kukko, E. I., 6 Kulczycki, E., 229, 236, 239–241, 243, 248 Kulichevskaya, I. S., 47 Kunte, H. J., 27 Kunz, R. C., 178, 188, 263 Kuo, S. S.-J., 179, 181, 189–190 Kurane, R., 36 Kuypers, M. M. M., 32 Kuzelka, J., 201, 207 Kyrpides, N. C., 82 L Laidler, V., 136 Lai, J. C.-H., 178, 181, 191 Lamer, S., 174 LamontI, L., 233 Land, M. L., 9 Lane, D. J., 41 Langenfelds, R. L., 272 Lantin, R. S., 277, 280, 283–284 Lanze´n, A., 90 Lapage, S. P., 49 Lapidus, A., 83–90, 94–95 Larimer, F. W., 9 Larive, C. K., 228–229, 232–233, 236–238, 240, 242, 248, 260–261, 265 Larsen, J., 168 Larsen, ., 46–47, 63–64, 74, 100 Larsen, Q., 4, 9 Latypova, E., 94 Leak, D. J., 64, 136 Lebaron, P., 151 Lebron, J., 101 Lee, H., 179–180 Lee, J. A., 9–10 Lee, J. C.-M., 178 Lee, J.-F., 190 Lee, S. G., 213, 216 Lees, V., 64 Lee, S.-W., 32, 239, 248–249, 260–261 Lee-Taylor, J., 273–274 Lefe`vre, J. F., 233 Legler, T. C., 9 Lentzen, G., 27 Leong, J., 233 Le Paslier, D., 32 Leslie, L., 239–240 Lessel, E. F., 49 Levine, S. R., 83–90, 94–95 Levinson, B. T., 229 Levin, W., 76 Levy, S., 82 Lewis, D. E., 229–230, 248, 260 Lewis, M., 46, 64, 74 Lidstrom, M. E., 16, 26, 36–37, 50, 54, 64, 83–84, 90, 100–102, 106–107, 109, 113, 124,

127–128, 149, 151–152, 156,–159, 161–162, 164, 180, 189–190, 250, 255 Lieberman, R. L., 178–179, 188–189, 196, 201, 203, 207, 212, 216 Liesack, W., 32, 41, 47, 52, 54, 121 Li, H., 92 Lillehaug, J. R., 64, 67–70, 74, 76, 167, 169, 171–173, 175 Lin, J.-L., 16, 100–102 Linke, D., 66, 222 Lippard, S. J., 46, 137, 179, 201, 207 Lipscomb, J. D., 136–137, 179, 231–232 Liu, C., 271, 276–277 Liu, Y., 137 Li, W., 90 Llenaresas, D., 277, 283–284 Lloyd, D., 284 Lo, C.-C., 21–22, 26 Loffler, C., 64 Lohmann, U., 272 Long, K. D., 277 Lontoh, S., 213 Lopez-Juez, E., 273 Louis, P., 17 Louis, S., 273 Lowry, D., 273 Lowry, R. K., 50 Lucas, S. M., 9 Ludwig, W., 48–49, 54–55 Luginbu¨hl, P., 92 Luo, M., 213 Luo, W.-I., 178 Lupas, A. N., 66 Ly, B. V., 47, 50, 100 Lynch, J. M., 64 M MacDonald, J., 221 Machuca, A., 251 Madan Babu, M., 67 Madera, M., 67 Madronich, S., 273–274 Madsen, E. L., 83 Madsen, R. A., 276, 278 Madupu, R., 74 Makarim, A. K., 280 Makarova, K. S., 100 Malfatti, S. A., 9 Mangenot, S., 32 Maniatis, T., 85, 91, 101, 106 Manolukas, J., 3, 7 Marin-Sanguino, A., 27 Martell, A. E., 253 Marth, G., 92 Martin, H. G., 87, 140 Martinho, M., 196 Marx, C. J., 26, 106–107, 124, 127–128

296 Massardo, D. R., 100 Masselon, C., 238 Masterson, L. R., 229–230, 248, 260 Mathis, J. N., 174 Mathur, E. J., 51, 82, 88 Mausezahl, I., 64 McCammon, S. A., 50–51 McCann, C., 239–240 McDermitt, D. K., 276, 278 McDonald, I. R., 16, 25, 32, 35–36, 50, 100–102, 136–137, 140, 213, 232 McEllistem, T. M., 248, 260–262 McEllistrem, M. T., 229, 233, 239, 248–249, 260–263 Mc Elwain, M. C., 3, 7 McEwan, A. G., 76 McHardy, A. C., 83–90, 92, 94–95 McIntire, W. S., 106 McLeod, A., 273–274 McMeekin, T. A., 50 McNamara, N. P., 83 McQuaide, S. C., 149, 161–162, 164 Medvedkova, K. A., 3 Mehrens, M., 151 Mehta, P. K., 213, 216, 218 Meier, R., 273 Meldrum, D. R., 161–162, 164 Merkx, M., 137 Merriman, B., 92 Mertens, E., 2–3, 7 Mertz, C., 230, 233 Mesbah, M., 52 Messenger, D. J., 273–274 Messing, J., 102 Methe, B. A., 74 Meyerdierks, A., 87 Meyer, T. F., 174 Michaels, G., 3 Miguez, C. B., 32, 100, 136 Milagres, A. M. F., 251 Miller, J. R., 85 Miquez, C. B., 36 Mishra, S., 213, 216, 218 Miyaji, A., 203, 211, 219, 221–223 Miyakawa, K., 199, 219, 221 Miyatake, K., 7 Molter, T. W., 161–162, 164 Moncrieff, J., 284 Moore, S. A., 7 Moreira, D., 7, 9–10 Moreno-Sanchez, R., 2 Morgan, H. W., 7 Morimoto, H., 178 Morton, J. D., 248 Mosier, A. R., 276, 280–281 Motekaitis, R. J., 253 Mountain, B. W., 47, 50, 100 Mulheron, H. J., 229, 233, 260

Author Index

Mu¨ller, M., 9–10 Muller, V., 21 Mu¨nck, E., 196 Murooka, Y., 24, 26 Murphy, D. V., 278 Murphy, K. C., 122 Murrell, C., 100–102 Murrell, J. C., 3–4, 8–9, 16, 22–26, 32–33, 35–37, 48, 64, 67–70, 83–84, 100, 106, 111, 119–121, 123, 135–137, 139–140, 143–144, 167–169, 172, 213, 229, 232–233, 243, 260 Mustakhimov, I. I., 4, 6, 8–10, 15–16, 21–26, 107–109, 112 Mutters, R., 55 Myers, C. R., 74 Myers, J. M., 74 Myhre, G., 272 N Nakagawa, K. H., 189–190 Nakorchevsky, A., 59 Napolea˜o, D., 251 Navarro, R., 277, 281 Neftel, A., 272, 280, 284 Neilands, J. B., 228, 249, 253 Nelson, K. E., 46, 64, 74, 82 Nelson, S. F., 92 Nelson, W., 74, 82 Nercessian, O., 84, 90 Neue, H. U., 277, 280, 283–284 Neufeld, J. D., 83 Newcomb, M., 46 Ng, K. Y., 178 Nguyen, H.-H. T., 177–178, 180, 184–185, 189–190 Nichols, P. D., 52, 55 Nichol, T., 143 Nielsen, H., 67 Nielsen, J. L., 100 Nielsenm, J. L., 46, 48, 56 Nielsen, P. H., 46, 48, 56, 100 Niemann, H., 55 Nieto, J. J., 17 Nikitin, D., 16, 100–102 Nimmo, R. H., 273 Nisbet, P. B. R., 273 Nisbet, R. E. R., 273 Norton, J. M., 9 Noyes, E., 84, 90 Nutricati, E., 100 O Oakley, C. J., 144 O’Brien, W. E., 3, 7, 10 O’Doherty, S., 272 O’Donnell, A. G., 101

297

Author Index

Oesterhelt, D., 27 Ofer, A., 17 Oh, J.-I., 213, 216 Oh, S. H., 122 Ojala, D. S., 22, 99, 149, 159 Okubo, Y., 113 Okura, I., 199, 203, 213, 216–219, 221–223 Olsen, J. E., 55 Omelchenko, M. V., 100 Ongena, M., 260 Ono, H., 24, 26 Ono, M., 217–218 Op den Camp, H. J. M., 32, 47, 51, 56, 100 Ostle, N., 83 Owens, N. J. P., 243 ystein, A., 55 P Page, W. J., 233 Palmer, P. I., 273–274 Pancost, R. D., 50 Panikov, N. S., 41 Pan, J. M., 50 Pan, Y., 47, 56 Papen, H., 277–278, 280–281, 283–284 Parekh, N. R., 83–84 Park, S., 213 Paulsen, I. T., 74, 82 Payne, S. M., 233 Pease, R. L., 140 Pedersen, P. L., 75 Pederson, A. R., 281 Peeples, T. L., 64, 106 Pelletier, D. A., 41 Pelletier, E., 32 Peplies, J., 49 Petersen, S. O., 281 Peters, R., 17 Petit, M., 151 Petzel, J. P., 3, 7 Pfeffer, M., 55 Pfeiffer, F., 27 Pfeiffer, H. P., 16 Pfleiderer, C., 7 Pfluecker, F., 16 Phelps, P., 229, 240 Philippe, H., 7, 9–10 Phillips, K. C., 4, 33, 49, 101, 170, 249–250 Phipps, D., 151 Pica, N., 17 Piemont, Y., 230, 233 Pierik, A. J., 17 Pihlatie, M., 284 Pilegaard, K., 272, 284 Pilkington, S. J., 145 Pineda, E., 2 Pirt, F. J., 64

Pittelkow, M., 20 Podar, M., 51, 82, 88 Pohl, N. L., 239, 248–249, 260–263 Pol, A., 32, 47, 51, 56, 100 Pollack, J. D., 3, 7 Pomerantz, W. J., 243 Pontieri, P., 100 Poole, R. K., 228 Poptsova, M. S., 65 Poret-Peterson, A. T., 9 Prabhu, J., 17 Prati, M. V., 100 Premachandran, U., 52 Priefer, U., 102 Priestley, N. D., 178 Prigent, Y., 277 Prinn, R. G., 272 Prior, S. D., 64, 136, 168 Priya, M. L., 67 Pruesse, E., 49 Pu¨hler, A., 102 Pullen, J. K., 140 Puntervoll, P., 171–172 Q Qi, J., 90 Quast, C., 49 Quayle, J. R., 3, 64 R Radajewski, S., 32, 83–84, 100, 136 Rahalkar, M., 50, 54 Raivonen, M., 272, 284 Ramachandran, S., 272 Rampp, M., 27 Ram, R. J., 82, 87 Ramsay, B. A., 240 Ramsay, J. A., 240 Rasche, F., 55 Ratkowsky, D. A., 50 Ravel, J., 82 Rawat, S., 178, 196–197, 199, 201–203, 207–208, 212–213 Reay, D. S., 273–274 Reeves, R. E., 2, 7, 9 Reichardt, W., 42 Reichart, G.-J., 47, 56 Reichenauer, T. G., 55 Reid, W. W., 64 Reigstad, L. J., 47, 100 Reina-Bueno, M., 17 Reisinger, V., 71–72 Remington, K., 82, 89 Remmert, M., 66 Rennenberg, H., 277–278, 283–284 Renstrom, J. M., 229, 232 Ren, Y., 47, 50, 100

298

Author Index

Reshetnikov, A. S., 4, 6, 8–10, 15–18, 20–26, 107–109, 112 Reuter, K., 20 Reynaerts, A., 55 Richardson, P. M., 9, 82, 87 Richardson, T. H., 92 Richter, M., 82 Ridgway, H. F., 151 Rigby, M., 272 Rigoutsos, I., 87 Rijpstra, I. C., 52 Rinne, J., 277, 284 Riutta, T., 284 Rivera, M. M., 276 Roberson, R. S., 7 Roberts, J. A., 229, 236, 239–241, 243, 248 Rochette, P., 275, 278–280 Ro¨ckmann, T., 273 Rodriguez, G. G., 151 Rohmer, M., 55, 168 Rokhsar, D. S., 82, 87 Ronimus, R. S., 7 Rosenberg, F. A., 51 Rosenzweig, A. C., 136, 178–179, 188–189, 195–197, 199, 201–203, 207–208, 212–213, 223, 228–229, 232 Rosenzweig, R. C., 212, 216 Rossello´-Mo´ra, R., 48, 54–55 Rothe, A., 277, 281 Rothstein, R. J., 122 Rozova, O. N., 1, 3–4, 6, 8–10, 24–26 Ruan, J., 92 Rubin, E. M., 82, 87 Rusch, D. B., 82, 89 Ruse, C. I., 59 Russell, D. W., 4 Ryan, D., 76 Ryan, M. D., 21 S Saavedra, E., 2 Sabarth, N., 174 Sacchi, N., 4 Sakai, Y., 52 Sakharovsky, V. G., 4, 10, 16, 100–102 Sakwa, J., 4, 9, 46, 64, 74 Salah El Din, A. L., 233 Salameh, P. K., 272 Salamov, A. A., 82–90, 94–95 Salih, B., 238 Salmond, G. P. C., 64, 136–137 Salyuk, A., 56 Sambrook, J., 4, 85, 91, 101, 106, 129 Sankaran, K., 67 Santos, H., 3, 7, 17 Sauer, T., 16 Saum, S. H., 21

Savagea, K., 277, 281 Sawada, K., 24, 26 Saw, J. H., 100 Scanlan, D., 46, 64, 74 Scanlan, J., 140 Scardino, L. L., 229, 232, 239, 248–249, 260–261 Scha¨fer, A., 121 Scha¨fer, M., 260 Schagger, H., 71 Schamel, W. W., 71 Schauwecker, F., 17 Schimmel, P., 122 Schink, B., 50, 54 Schjoerring, J. K., 272, 284 Schleper, C., 90 Schmidt, R., 9 Schmidt, T. M., 82 Schnaitman, C. A., 168, 170 Schnitzler, J. P., 273 Scho¨ning, B., 46, 48, 56, 100 Schouten, S., 32, 47, 56 Schreiber, F., 32 Schro¨der, P., 283 Schultz, A., 10 Schulz, G. E., 65, 69 Schuster, S. C., 27, 90 Schweisfurth, R., 48 Schwibbert, K., 27 Schwyn, B., 249, 253 Scott, D., 216–217, 221 Scow, K. M., 9 Sealy, J. A., 36 Seeliger, H. P. R., 49 Seiler, W., 277, 283–284 Seitz, H., 27 Seki, Y., 217–218 Selvan, A. T., 67 Semiletov, I., 56 Semrau, J. D., 32, 41, 101, 136, 178, 188, 196, 213, 229, 232–233, 239, 247–249, 252–253, 259–263 Senin, P., 47, 50, 100 Sensen, C. W., 9–10 Serrano, R., 2, 7 Servais, P., 151 Seshadri, R., 74 Seufert, G., 284 Severin, J., 16 Shafe, P. H., 260–261, 263 Shah, N. N., 213 Shakhova, N., 56 Shapiro, H. J., 82 Shaw, J. H., 47, 50 Shchukin, V. N., 10, 16, 21 Sherwood, P. M. A., 228–229, 236, 248, 260 Shiemke, A. K., 180 Shigematsu, T., 36 Shimoda, M., 216–218

299

Author Index

Shin, M. W., 9 Shinmyo, A., 24, 26 Shishkina, V. N., 3 Shi, Y., 90 Short, J. M., 51, 82, 88 Shrestha, D. B., 178, 188, 201, 207 Shuckburgh, E. F., 273 Siebers, B., 7 Sigmund, J., 47 Silhavy, T. J., 168 Silver, W. L., 276 Simmonds, P. G., 272 Simon, R., 102 Simpson, D., 272, 284 Sinninghe Damste´, J. S., 47, 52, 56 Sivam, D., 113 Skerman, V. B. D., 49 Skerratt, J. H., 50–52 Skovran, E., 113 Slade, S. E., 140 Slamovits, C. H., 3 Sly, L. I., 47, 52 Smirnova, A. V., 47, 50, 100 Smirnova, K. V., 47, 52, 54 Smith, K. A., 274 Smith, R. M., 253 Smith, S. M., 178, 195–197, 199, 201–203, 207–208, 212–213, 221, 228, 232 Smith, T. J., 100, 135–136, 140, 143, 146 Smolders, A. J. P., 47, 56 Sneath, P. H. A., 49 Soding, J., 66 Sokolov, A. P., 3, 8 Solonin, A., 16, 100–102 Solovyev, V. V., 82, 87 Sorek, R., 90, 92 Sorokin, D. Y., 16, 50, 54, 100–102, 113 South, D. J., 2, 7 Sriskantharajah, S., 273 Stackebrandt, E., 47 Stafford, G. P., 140 Stahl, D. A., 41 Stainthorpe, A. C., 64 Stanley, S. H., 64, 136 Stan-Lotter, H., 64, 168 Starkenburg, S. R., 9 Starostina, N. G., 10, 100–102 Steele, L. P., 272 Stege, J. T., 92 Steigner, D., 273 Steimle, V., 71 Stein, L. Y., 9 Stemmer, M., 55 Stemmler, T. L., 178, 188, 196–197, 199, 201–203, 207–208, 212–213, 223, 228, 232, 239 Steudler, P. A., 243 Stirling, D. I., 136

Stoecker, K., 46, 48, 56, 100 Stokes, A. N., 50 Stolyar, S., 36–37, 64, 106 Stott, M. B., 32, 47, 50, 56, 100 Stralis-Pavese, N., 83 Straume, A. H., 67–70, 167, 169, 172 Straume, D., 171–172 Stro¨m, T., 3, 64 Strous, M., 100 Strovas, T. J., 107, 109, 159, 161–162, 164 Stubner, S., 43 Suchorolski, M. T., 161–162, 164 Suciu, D., 83–90, 94–95 Sugimori, D., 213, 218 Susenbeth, A., 277 Susskind, B. M., 9 Sutton, G., 82, 85, 89 Sutton, M. A., 272, 284 Suzina, N. E., 3–4, 10, 16, 21, 32–33, 41, 47, 50, 54, 100–102 Suzuki, M., 203, 219, 222–223 Svenning, M. M., 50 Szeto, E., 82–90, 94–95 T Takano, M., 24, 26 Takeguchi, M., 199, 213, 218–219, 221 Takeguchi, T., 218 Talbot, H. M., 50, 239–240 Talibart, R., 16 Talor, R. T., 213 Tamaoka, J., 55 Tannich, E., 3, 7 Taraz, K., 260 Tas, B., 277 Taylor, A., 229, 236, 238, 240, 242, 261 Taylor, R. T., 213 Taylor, S. C., 64 Teal, T. K., 82 Tedesco, D., 47, 51, 100 Teeling, H., 82, 87 Teintze, M., 233 Telser, J., 196, 201, 207, 212, 223, 239 Theisen, A. R., 32–33, 100, 120–123, 136 Thestrup, H., 69, 71, 168, 170–171 Thomas, P. E., 76 Thonart, P., 260 Tian, Q., 179–180 Tiedje, J. M., 41 Timms, P., 174 Tinberg, C. E., 239 Tindall, B. J., 48, 54–55 Toa, T., 24, 26 Toenshoff, E. R., 46, 48, 56, 100 Toledo, G., 51 Tourova, T. P., 16 Toyama, H., 121–122

300

Author Index

Tran, B., 82, 89 Trapido-Rosenthal, H. G., 51 Tredici, S. M., 100 Treude, T., 48 Tringe, S. G., 82, 88, 92 Trotsenko, Y. A., 1, 3–4, 6, 8–10, 15–18, 20–27, 33, 41, 46, 50, 52, 54, 100–102, 107–109, 112–113, 136 Troussellier, M., 151 Tru¨per, H. G., 16–17 Trust, T. J., 69 Tsai, S.-F., 179–180 Tseng, C.-F., 178–179, 181–182, 184–186, 189–190, 196 Tseng, M. Y.-H., 178–179, 181–182, 184–186, 189–190, 196 Tsien, H. C., 55 Tsirigos, A., 87 Tso, L., 178 Tsyrenzhapova, I. S., 26 Tuittila, E. S., 284 Tu, L.-C., 178 Tuovinen, J. P., 284 Turner, M. S., 174 Tyson, G. W., 82, 87, 90, 150 U Ullrich, S., 151 Urich, T., 90 V Valdivia, R., 110 Valentini, R., 284 Valla, S., 140 van Alen, T., 100 van den Bulk, W. C. M., 281 van de Pas-Schoonen, K. T., 100 van der Helm, D., 233 van der Vlag, J., 3 van Ginkel, C. G., 216 Van Gorden, G. S., 229, 232 Van Praag, E., 3, 7 van Schaftingen, E., 2–3, 7 van Vliet, A. H., 90 van Weelden, H., 273 van Winden, J. F., 47, 56 Vargas, C., 17, 76 Veglia, G., 229–230, 248, 260 Venter, J. C., 82 Ventosa, A., 17 Verchot, L. V., 273, 277, 281 Vergez, L. M., 9 Vermeulen, A. T., 281 Vieira, J., 102 Vigano, I., 273 Vigliotta, G., 100 Vinther, F. P., 281

Visca, P., 233 Visser, J., 3 Vo¨lker, H., 48 Volkert, A. A., 229, 232 Von Heijne, G., 67 von Jagow, G., 71 von Mering, C., 82, 88 von Tigerstrom, M., 233 Vorobe´v, A. V., 32, 36–37, 46, 54, 120 Vossen, W., 260 Vraspir, J. M., 233 Vrijbloed, J. W., 3 Vuilleumier, S., 8, 10, 229, 233, 260 W Wackett, L. P., 55, 136 Wagner, M., 46, 48, 56, 100 Wagner, T., 50 Walker, S., 168 Wallar, B. J., 137 Wampler, D. E., 27 Wanfang, L., 280 Wang, E., 172 Wang, J., 47, 50, 100 Wang, L., 213 Wang, T., 94 Wang, V. C.-C., 178, 181, 191 Wang, Y.-S., 178–179, 181–182, 184–186, 189–190, 196, 276 Wang, Z., 280 Ward, B. B., 9 Ward, N., 4, 9, 46, 64, 74 Warnecke, F., 92 Warren, L. G., 2, 7, 9 Wartiainen, I., 50 Wassmann, R., 277, 280, 283–284 Watzinger, A., 55 Weijers, C. A. G. M., 216 Weisburg, W. G., 41 Weiss, R. F., 272 Wessels, H. J. C. T., 32 West, C. A., 136–137 Westhead, E. W., 27 White, D. C., 55 Whiteley, A. S., 101 Whitman, W. B., 52 Whittenbury, R., 4, 33, 49, 64, 101, 137, 170, 249–250 Whittington, D. A., 137 Wilkinson, B., 178 Wilkinson, J. F., 4, 33, 49, 64, 101, 170, 249–250 Wilkinson, S. P., 22–23 Willenbrock, H., 67 Williams, M. V., 3, 7 Williamson, S., 82, 89 Williams, P. G., 178 Winter-Larsen, H. C., 140 Woermann, D., 283

301

Author Index

Wohlfarth, A., 16 Wolf, B., 277 Wolf, Y. I., 100 Wollscheid, B., 71 Wommack, K. E., 82 Wood, A. P., 121 Woodford, S. K., 56 Wood, H. G., 3, 7, 10 Wood, T. K., 121 Woyke, T., 82 Wu, D., 74, 82, 89 Wuebbles, D. J., 272–273 Wu, H., 213 Wu, K.-M., 179–180 Wu, L.-Y., 178 Wu, M., 74 Wysoker, A., 92 X Xie, G., 21–22, 26 Xing, X.-H., 100, 213 Xu, L., 276, 278 Y Yamamoto, M., 24, 26 Yang, R. B.-G., 178 Yanish-Perron, C., 102 Yao, Z., 276–277 Yates, J. R., 59 Yatsunyk, L. A., 178, 212–213, 228, 232 Yi, E.-C., 179–180

Yip, J. H.-K., 178, 184–185 Yoon, S., 32, 136, 196, 247–249, 252–253, 260 Yooseph, S., 82, 89 Yue, J., 276 Yue, S., 151 Yurimoto, H., 52 Yuryev, A., 47, 50, 100 Yu, S. S.-F., 177–182, 184–186, 189–191, 196 Yusupov, V., 56 Yutin, N., 100 Yu, Y., 240 Z Zabinsky, R., 157 Zahn, J. A., 178, 188, 198, 229, 231, 236, 238, 240, 242, 248, 260–263 Zea, C. J., 239, 248–249, 260–263 Zealey, G. R., 122 Zedelius, J., 32 Zenobi, R., 238 Zerkle, A. L., 47 Zhang, C., 100 Zheng, X., 276–277 Zhou, Z., 47, 50, 100 Zhu, C., 50 Zhu, M., 178 Ziebis, W., 48 Zimmer, I., 273 Zimmermann, R., 151 Zimny-Arndt, U., 174

Subject Index

A Aerobic methanotroph characterization, for systematic research, 46. See also Methanotrophic bacteria, characterization and description Agar plates preparation, chalkophore production Cu–CAS assay, 251–253 Fe–CAS assay, siderophores, 255–256 Antibiotic resistance markers, for Methylomicrobium spp., 106 Aspartate kinase (Ask), ectoine biosynthesis, 26–27 ATP-dependent 6-phosphofructokinase (ATP-PFK), 2–3, 7 B Bacterial cells, pMMO activity assay of, 215–216 alkanes, alkenes, and halogenated derivatives, 216–218 culture medium, 214–215 culture of, 215 methanol production, application for, 216, 218 b-Barrel outer membrane protein predictor (BOMP), 66 Biosphere–atmosphere exchange, methane (CH4) closed chamber technique design of, 278–279 flux calculations, 280–282 gas mixing, 276–277 sampling procedure, 278, 280 setup, of gas fluxes, 275 soil disturbance, 276 spatiotemporal variations, 277 temperature, pressure, and humidity perturbations, 276 dynamic chamber technique, 282–283 emission pathways, determination of, 283–284 micrometeorological measurements of, 284 plants in, 273–274 role of, 272 soil microbial processes and, 272 sources and sinks of, 273 Biotin labeling, 173–175 Blue-native polyacrylamide gel electrophoresis (BN-PAGE), inner membrane protein complexes

detergent extraction of, 71–72 large-scale 2D BN-/SDS-PAGE first-dimension, 72–73 second-dimension, 73–75 Triton X-100 solubilized membranes, 72–73 C Chalkophore production, microorganisms screening Cu–CAS assay for agar plates preparation, 251–253 solution preparation, 249–251 UV-visible absorption spectra of, 251 Fe–CAS assay, siderophores agar plates preparation, 255–256 preparation of liquid, 253–254 UV-visible absorption spectra of, 254 role of, 248 Chamber measurements, of CH4 exchange closed chamber technique design of, 278–279 flux calculations, 280–282 gas mixing, 276–277 sampling procedure, 278, 280 setup, of gas fluxes, 275 soil disturbance, 276 spatiotemporal variations, 277 temperature, pressure, and humidity perturbations, 276 dynamic chamber technique, 282–283 emission pathways, determination of, 283–284 Chemotaxonomy, methanotrophic bacteria, 52, 55 Column chromatography, detergent-solubilized pMMO, 222–223 Conjugal transformation, of Methylomicrobium strains, 106, 108 Conjugation Methylocella silvestris BL2, 123 Ms. trichosporium OB3b, 141–144 Copper CAS assay, for chalkophore production agar plates preparation, 251–253 solution preparation, 249–251 UV-visible absorption spectra of, 251 methanobactin solubilization studies, 239–240

303

304

Subject Index

Copper (cont.) pmoB subunit, soluble domain constructs, 206–207 uptake, MMO gene expression assays, 240–243 Copper-bound methanobactin (Cu-mb), 261 c-type heme proteins, Methylococcus capsulatus, 74–75 identification of, 76–77 in-gel staining, 76 staining on protein transfer membranes, 76 Cyanide role, in metal reconstitution, 200–201 D Detergent-solubilized pMMO, purification column chromatography, 222–223 crystallization of, 223 metals in, 223 water phase, 221–222 Diaminobutyric acid (DABA) acetyltransferase, 24–25 aminotransferase, 23–24 Dianion HP20 column, methanobactin (mb) isolation, 263–265 Dodecyl-b,D-maltoside (DDM), 221–222 Dynamic chamber technique, CH4 exchange, 282–283 E Ectoine biosynthesis, in halotolerant methanotrophs aspartate kinase, 26–27 DABA acetyltransferase, 24–25 diaminobutyric acid aminotransferase, 23–24 ectoine synthase, 25–26 halophilic/halotolerant bacteria, 27 PCR-based approach, identification of aspartate kinase, 21 conserved regions of, 19 ectABC-ask, 19 ectABC operon, 17–18 oligonucleotide primer sequences, 18 orfs of, 18–19 sequencing strategies, 20 types of, 21 vectorette unit, 21 transcriptional regulation of, 22–23 Ectoine synthase, 25–26 EDTA role, in metal reconstitution, 199–200 Electroporation, Methylocella silvestris competent cells preparation, 124 of linear DNA, gene deletion, 123–124 method, 127 plasmid DNA transformation by, 123 Electrospray ionization time-of-flight (ESI-TOF) mass spectra, 266

F Facultative and obligate methanotrophs alphaproteobacterial, 32 culture purity, tests for complex organic media, 38–39 dilution-extinction growth experiments, 42 methane monooxygenase-coding genes, quantification of, 42–43 phase-contrast and electron microscopy, 39 16S rRNA gene clone library analysis, 41–42 whole-cell hybridization, with fluorescent probes, 39–41 identification of culture conditions, 33–34 growth dynamics on methane, 34–35 intracytoplasmic membrane structures, 35 methane monooxygenase encoding genes, 35–37 pmoA and mmoX genes amplification, PCR primers, 36 Methylocella species, 32 particulate methane monooxygenase enzyme (pMMO), 32–33 substrate utilization tests, 37–38 Fae peptides, phylogenetic diversity, 87, 89 Fe–CAS assay, chalkophore production agar plates preparation, 255–256 preparation of liquid, 253–254 UV-visible absorption spectra of, 254 Flow cytometry-based redox sensing for actively metabolizing microbes, 156–158 cell physiological state, evaluation of, 152, 156 principle, 151 redoxsensor green dye, 151–152 summary of, 153–155 Fluorescent probes, whole-cell hybridization, 39–41 Fructosebisphosphate aldolase (FBA), 6 Functional metagenomics, of methylotrophs database and DNA sequencing and assembly, 85–86 gene-centric analysis, 85, 87 organism-centric analysis, 87–89 stable isotope probing (SIP) DNA extraction, isopycnic centrifugation, and labeled DNA recovery, 84–85 sample collection and cell labeling, 83–84 strategy for, 83–84 ultrashort read-based data processing, 91–93 metatranscriptome coverage and specificity, 93–95 mRNA enrichment and (optional) cDNA synthesis, 91 principle and strategy, 90 RNA isolation, 90–91

305

Subject Index G Gene-centric analysis, functional metagenomics functional genes in Fae peptides, phylogenetic diversity of, 87, 89 tetrahydromethanopterin-linked formaldehyde oxidation in, 87–88 phylogenetic markers, 85, 87 Gene deletion efficiency of, 127–128 by homologus recombination, 124–126 of isocitrate lyase, 128 of linear DNA, by electroporation, 123–124 Genetic systems Methylocella silvestris BL2 competent cell preparation, 124 conjugation and electroporation, 123 electroporation, 127 electroporation, gene deletion by, 123–124 gene deletion, efficiency of, 127–128 growth of, 122–123 homologus recombination, gene deletion by, 124–126 isocitrate lyase deletion, 128–130 for moderately halo (alkali)philic bacteria (see Methylomicrobium spp., genetic systems) Glycolysis, 2 H Halo (alkali)philic bacteria. See Methylomicrobium spp., genetic systems HPLC, purification of methanobactin, 236 hpp genes, 9–10 I Inclusion bodies, pmoB subunit isolation and purification of, 204–205 refolding of, 205–206 Intracytoplasmic membrane structures, facultative and obligate methanotrophs, 35 Isocitrate lyase complementation, 128–129 deletion of, 128 phenotype of, 129–130 L Lipoproteins, Methylococcus capsulatus, 67 M MALDI-TOF spectrum, of methanobactin, 237 Mass spectral analysis, methanobactin, 237–238 Membrane-bound form, of pMMO electron donors for, 221 isolation of

cell disruption, 219 centrifugation, membrane fractions collection, 220 protein analysis, 220–221 Metal reconstitution, of pMMO apo pMMO, preparation of activity of, 202 assays of, 198–200 cyanide, 200–201 EDTA, 199–200 membrane isolation, 198 Methylococcus capsulatus (Bath) growth conditions, 197 copper role in, 196 crystal structures of, 196–197 Metatranscriptomics, ultrashort sequence reads, 90–95 Methane (CH4) biosphere–atmosphere exchange (see Biosphere–atmosphere exchange, methane (CH4)) facultative and obligate methanotrophs growth dynamics, 34–35 oxidation analysis flow cytometry-based redox sensing, 151–158 respiration detection system, 161–165 respiration response imaging, 158–161 oxidation, soluble pmoB domains, 208 Methane monooxygenase (MMO) encoding genes identification of, 35–37 quantification of, 42–43 gene expression assays, Cu uptake, 240–243 pmoA gene expression, 241–242 transcription assay, 242–243 Methane oxidizing bacteria (MOB), methanobactin (mb) isolation chalkophores and siderophores, 260 purification of, 265 sample variability, 268 ESI-TOF mass spectrum of, 266 reverse-phase HPLC chromatography, 267 from spent media concentration of, 263–265 pmoA, RT-PCR of, 262 from whole cells, 261–263 yields in, 261 Methanobactin isolation and purification initial capture and concentration, 233–235 mass spectral analysis, 237–238 purification of, 236 toxicity and growth, assessment of, 238–239 UV-Vis detection of, 235–236 methanotroph growth and production iron effects, 233 Methylosinus trichosporium OB3b, 232–233

306 Methanobactin (cont.) nitrate mineral salts (NMS) media, 231 MOB, from spent media (see Methane oxidizing bacteria (MOB), methanobactin (mb) isolation) in pseudonatural environments copper minerals, solubilization studies with, 239–240 MMO gene expression assays, Cu uptake, 240–243 structures of, 229–230 Methanol production, Methylosinus trichosporium OB3b, 216, 218 Methanotrophic bacteria, characterization and description aerobic methanotroph, for systematic research, 46 methodological approaches agar media, 50–51 chemotaxonomy, 52, 55 culture collections, 56–58 enrichment and isolation, 49–51 guanosine–cytosine (GþC) ratios and fatty acid profile, 52, 55 sequence-based analyses and genomic comparisons, 55–56 standards for, 52–54 systematic diversity bacteriological code, requirements, 48–49 Crenothrix polyspora, 47–48 environmental relevance and knowledge limitations, 48 Proteobacteria, 46–47 type I and type II, 47 Methylocella silvestris BL2 competent cell preparation, 124 description of, 120 electroporation, 127 gene deletion efficiency, 127–128 genome sequence, 120–121 growth of, 122–123 homologus recombination, gene deletion by, 124–126 introduction of DNA, 121–122 isocitrate lyase complementation, 128–129 deletion of, 128 phenotype of, 129–130 linear DNA, electroporation of, 123–124 plasmid DNA, 123 Methylocella species, 32 Methylococcus capsulatus (Bath) computational algorithms, 65 c-type heme proteins identification of, 76–77 in-gel staining, 76 staining on protein transfer membranes, 76 electron microscope of, 170

Subject Index

genome sequence of, 64 inner membrane protein complexes, BN-PAGE detergent extraction of, 71–72 large-scale 2D BN-/SDS-PAGE, 72–74 Triton X-100 solubilized membranes, 72–73 metal reconstitution, of pMMO, 197 outer membrane preparation (see Outer membrane preparation, Methylococcus capsulatus) outer membrane proteome, mapping of b-barrel proteins, 66 experimental identification, 67–70 lipoproteins, 67 unannotated proteins identification, 70 pMMO isolation and purification activities of, 191 chemical composition, 189–190 DEAE-sepharose fast flow column, 188–189 direct transfer and gel filtration, 185 reconstitution, 188 solubilization, 185–187 PPi-PFK, 8–10 redundancy in, 64 SDS-PAGE analysis, 171, 173 Methylomicrobium alcaliphilum 20Z, 101, 108–109 PPi-PFK, 10 Methylomicrobium buryatense 5G and 5B, 101 Methylomicrobium spp., genetic systems, 116 aerobic methanotrophy, 100 broad-host-range vectors in, validation and application of antibiotic resistance markers for, 106 conjugal transformation of, 106, 108 construction of, Methylomicrobium alcaliphilum 20Z mutant strains, 108–109 DNA, RNA purifications, and mRNA enrichment, protocols for, 104–105 genetic manipulations, 101, 106 Methylomicrobium alcaliphilum 20Z and Methylomicrobium buryatense 5G and 5B, 101 pCM66 plasmid-based promoter-probe vector, 109 plasmids in, 107–108 strains and growth conditions, 102–103 gene expression studies microarray design, 112–113 microarray platform, validation of, 113–115 RNA isolation and labeling, 113 pDO4-based, construction of ectAp1p2 promoter regions, 112 plasmid maps, 111 promoter-probe vector, 110 20Z, expression vectors for, 110–112

307

Subject Index

Methylosinus trichosporium OB3b bacteriological growth media and antibiotics, 137–139 components, 136–137 description, 136 expression hosts and conjugation, 141–144 mutagenesis, 140 mutant analysis biomass production, 145–146 characterization, 145 genotype confirmation, 144 mutant design, 139–140 pMMO activity assay of, 215–216 alkanes, alkenes, and halogenated derivatives, 216–218 alkene epoxidation, stereochemistry of, 218 bacterial cells, culture of, 215 culture medium, 214–215 detergent-solubilized, purification of, 221–223 dinuclear copper site, 212 membrane-bound form of, 219–221 methanol production, application for, 216, 218 subcloning of mutants, 141 Microarray, Methylomicrobium spp. design, 112–113 platform for biological replicates, Log2 ratio comparison, 114 dye-swap microarray data, 113–114 high salinity vs. low salinity, 115 Micrometeorological measurements, of CH4 exchange, 284 Microwell array chips, 162–163 MMO. See Methane monooxygenase mmoX genes amplification, PCR primers, 36 mRNA enrichment and cDNA synthesis, ultrashort read-based metatranscriptomics, 91 Mutagenesis, 139–140 N Nitrate mineral salts (NMS) medium, 214, 231 O Obligate methanotrophs, 32. See also Facultative and obligate methanotrophs Organism-centric analysis, functional metagenomics, 87–89 Outer membrane preparation, Methylococcus capsulatus biotin labeling, of surface-exposed OMP, 173–175 integral and tightly associated, 172

isolation of, 170–171 Methylococcus capsulatus, 168 proteins, 168 seperation methods, 168 subcellular fractionation, 169 surface-associated proteins, extraction of, 172–173 Outer membrane proteome (OMP) mapping, Methylococcus capsulatus experimental identification of 2DE analysis, 68 horizontal multiple spot patterns, in 2DE gels, 69–70 multiple spots, 68–69 subfractionation, 67 prediction of b-barrel proteins, 66 lipoproteins, 67 unannotated proteins identification in, 70 P Particulate methane monooxygenase (pMMO), 178–179 isolation and purification, from Methylococcus capsulatus (Bath) activities of, 191 chemical composition, 189–190 DEAE-sepharose fast flow column, 188–189 direct transfer and gel filtration, 185 reconstitution, 188 solubilization, 185–187 metal reconstitution of apo pMMO, preparation of, 197–203 copper role in, 196 crystal structures of, 196–197 pmoB subunit, soluble domain constructs of, 203–208 from Methylosinus trichosporium OB3b activity assay of, 215–216 alkanes, alkenes, and halogenated derivatives, 216–218 alkene epoxidation, stereochemistry of, 218 bacterial cells, culture of, 215 culture medium, 214–215 detergent-solubilized, purification of, 221–223 dinuclear copper site, 212 membrane-bound form of, 219–221 methanol production, application for, 216, 218 overproduction of bacteria, culturing of, 182–183 enriched membranes, isolation of, 183–184 flow reactor for, 181–182 pCM66 plasmid-based promoter-probe vector, Methylomicrobium spp., 109

308 PCR-based approach, ectoine biosynthesis aspartate kinase, 21 conserved regions of, 19 ectABC-ask, 19 ectABC operon, 17–18 oligonucleotide primer sequences, 18 orfs of, 18–19 sequencing strategies, 20 vectorette unit, 21 pDO4-based promoter-probe and expression vectors, for Methylomicrobium spp., genetic systems, 110–112 pfp genes, 9–10 Phase-contrast and electron microscopy, facultative and obligate methanotrophs, 39 pMMO. See Particulate methane monooxygenase pmoA genes amplification, PCR primers, 36 pmoB subunit, soluble domain constructs activity assays of, 207–208 copper assays, 206–207 expression of, 204 inclusion bodies, isolation and purification of, 204–205 inclusion bodies, refolding of, 205–206 refolding assessment, methods for, 207 vectors design spmoB, 204 spmoBd1, 203 spmoBd2, 203 PPi-dependent pyruvate-phosphate dikinase (PPDK)., 2, 11 Propylene epoxidation pMMO activity assays, 198–200 soluble pmoB domains, activity assays of, 207–208 Protein analysis, membrane-bound form of pMMO, 220–221 Pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK) assay of with Fru 1,6-P2, 5 with Fru 6-P, 4–5 with sedoheptulose-1,7-bisphosphate/ ribulose-1,5-bisphosphate, 5–6 with sedoheptulose-7-phosphate, 5 cloning, expression, and purification, 4 culture conditions, 4 FBA activity, recombinant PPi-PFK, 6 in glycolysis and gluconeogenesis, 2–3 metabolic pattern, 3–4 in microorganisms, 7 properties of from Mc. capsulatus Bath, 8–10 from Mm. alcaliphilum 20Z, 10 routine genetic manipulations, 4

Subject Index R Redoxsensor green (RSG) dye, 151–152 Respiration, single cell methods detection system, microobservation chamber description, 161–162 measurements, 164–165 microwell array chips, 162–163 observation chamber preparations, 163–164 sample respiration data, 165 response imaging description of, 158–159 image processing, 161 observation chamber preparations, 159 protocol, 159–160 sample classification data, 161–162 RNA isolation and labeling, in Methylomicrobium spp, 113 ultrashort read-based metatranscriptomics, 90–91 16S rRNA gene clone library analysis, 41–42 S SDS-PAGE inner membrane protein complexes, 73–74 Methylococcus capsulatus surface proteins, analysis of, 171, 173 Siderophores, 228 Single cell methods, for methane oxidation analysis flow cytometry, 151–158 respiration detection system, 161–165 respiration response imaging, 158–161 Soluble methane monooxygenase (sMMO) bacteriological growth media and antibiotics, 137–139 components, 136–137 description, 136 expression hosts and conjugation, 141–144 mutagenesis, 140 mutant analysis biomass production, 145–146 characterization, 145 genotype confirmation, 144 mutant design, 139–140 subcloning of mutants, 141 Spent media, methanobactin (mb) isolation concentration of Dianion HP20, 263–265 Sep-Pak columns, 263 from whole cells centrifugation and filtration, 261–262 tangential flow/hollow fiber filtration, 262–263 Stable isotope probing (SIP), functional metagenomics enrichment for

309

Subject Index

DNA extraction, isopycnic centrifugation, and labeled DNA recovery, 84–85 sample collection and cell labeling, 83–84 strategy for, 83–84 Surface-associated proteins, 172–173 T Terrestrial ecosystems. See Biosphere–atmosphere exchange, methane (CH4) Tetrahydromethanopterin-linked formaldehyde oxidation, 88 Transcriptional regulation, of ectoine biosynthesis genes, 22–23 Triton X-100 solubilized membranes, BN-PAGE, 72–73 U Ultrashort read-based metatranscriptomics data processing alignment outputs, postprocessing of, 92

alignment to scaffold, 92 complexity and quality filtering, 92 database integration, 93 quality assessment, 91–92 statistical analysis and visualization, 93 metatranscriptome coverage and specificity mapping of, 94–95 sequencing, 93 mRNA enrichment and cDNA synthesis, 91 principle and strategy, 90 RNA isolation, 90–91 UV-Vis detection, of methanobactin, 235–236 V Vectors design, pmoB subunit, 203–204 W Whole-cell hybridization, with fluorescent probes, 39–41