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 ...

Author: Amy Rosenzweig | Stephen W. Ragsdale

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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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Contributors

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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Contributors

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

METHODS IN ENZYMOLOGY

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


VOLUME XXXV. Lipids (Part B) Edited by JOHN M. LOWENSTEIN VOLUME XXXVI. Hormone Action (Part A: Steroid Hormones) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XXXVII. Hormone Action (Part B: Peptide Hormones) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XXXVIII. Hormone Action (Part C: Cyclic Nucleotides) Edited by JOEL G. HARDMAN AND BERT W. O’MALLEY VOLUME XXXIX. Hormone Action (Part D: Isolated Cells, Tissues, and Organ Systems) Edited by JOEL G. HARDMAN AND BERT W. O’MALLEY VOLUME XL. Hormone Action (Part E: Nuclear Structure and Function) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XLI. Carbohydrate Metabolism (Part B) Edited by W. A. WOOD VOLUME XLII. Carbohydrate Metabolism (Part C) Edited by W. A. WOOD VOLUME XLIII. Antibiotics Edited by JOHN H. HASH VOLUME XLIV. Immobilized Enzymes Edited by KLAUS MOSBACH VOLUME XLV. Proteolytic Enzymes (Part B) Edited by LASZLO LORAND VOLUME XLVI. Affinity Labeling Edited by WILLIAM B. JAKOBY AND MEIR WILCHEK VOLUME XLVII. Enzyme Structure (Part E) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XLVIII. Enzyme Structure (Part F) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XLIX. Enzyme Structure (Part G) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME L. Complex Carbohydrates (Part C) Edited by VICTOR GINSBURG VOLUME LI. Purine and Pyrimidine Nucleotide Metabolism Edited by PATRICIA A. HOFFEE AND MARY ELLEN JONES VOLUME LII. Biomembranes (Part C: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER

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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


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


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


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VOLUME 448. RNA Turnover in Eukaryotes: Nucleases, Pathways and Analysis of mRNA Decay Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 449. RNA Turnover in Eukaryotes: Analysis of Specialized and Quality Control RNA Decay Pathways Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 450. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 451. Autophagy: Lower Eukaryotes and Non-Mammalian Systems (Part A) Edited by DANIEL J. KLIONSKY VOLUME 452. Autophagy in Mammalian Systems (Part B) Edited by DANIEL J. KLIONSKY VOLUME 453. Autophagy in Disease and Clinical Applications (Part C) Edited by DANIEL J. KLIONSKY VOLUME 454. Computer Methods (Part A) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 455. Biothermodynamics (Part A) Edited by MICHAEL L. JOHNSON, JO M. HOLT, AND GARY K. ACKERS (RETIRED) VOLUME 456. Mitochondrial Function, Part A: Mitochondrial Electron Transport Complexes and Reactive Oxygen Species Edited by WILLIAM S. ALLISON AND IMMO E. SCHEFFLER VOLUME 457. Mitochondrial Function, Part B: Mitochondrial Protein Kinases, Protein Phosphatases and Mitochondrial Diseases Edited by WILLIAM S. ALLISON AND ANNE N. MURPHY VOLUME 458. Complex Enzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles and Peptides Edited by DAVID A. HOPWOOD VOLUME 459. Complex Enzymes in Microbial Natural Product Biosynthesis, Part B: Polyketides, Aminocoumarins and Carbohydrates Edited by DAVID A. HOPWOOD VOLUME 460. Chemokines, Part A Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 461. Chemokines, Part B Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 462. Non-Natural Amino Acids Edited by TOM W. MUIR AND JOHN N. ABELSON VOLUME 463. Guide to Protein Purification, 2nd Edition Edited by RICHARD R. BURGESS AND MURRAY P. DEUTSCHER

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C H A P T E R

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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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Valentina N. Khmelenina et al.

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).

Pyrophosphate-Dependent 6-Phosphofructokinase from Methanotrophs

3

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

4


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

10


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


11

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 Geńoscope for high-throughput sequencing and annotation of the M. alcaliphilum 20Z genome (IbiSA-Geńoscope 2008 campaign) and M. G. Kalyuzhnaya for kindly providing microarray analysis information.

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Siebers, B., Klenk, H. P., and Hensel, R. (1998). PPi-dependent phosphofructokinase from Thermoproteus tenax, an archaeal descendant of an ancient line in phosphofructokinase evolution. J. Bacteriol. 180, 2137–2143. Slamovits, C. H., and Keeling, P. J. (2006). Pyruvate-phosphate dikinase of oxymonads and parabasalia and the evolution of pyrophosphate-dependent glycolysis in anaerobic eukaryotes. Eukaryot. Cell 5, 148–154. Stro¨m, 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, 465–472. Susskind, B. M., Warren, L. G., and Reeves, R. E. (1982). A pathway for the interconversion of hexose and pentose in the parasitic amoeba Entamoeba histolytica. Biochem. J. 204, 191–196. Trotsenko, Y. A., and Murrell, J. C. (2008). Metabolic aspects of obligate aerobic methanotrophy. Adv. Appl. Microbiol. 63, 183–229. Trotsenko, Y. A., and Shishkina, V. N. (1990). Studies on phosphate metabolism in obligate methylotrophs. FEMS Microbiol. Lett. 87, 267–271. Trotsenko, Y. A., Medvedkova, K. A., Khmelenina, V. N., and Eshinimaev, B. Ts. (2009). Thermophilic and thermotolerant aerobic methanotrophs. Microbiology (Moscow) 78, 387–401. Van Praag, E. (1997). Use of 3-D computer modeling and kinetic studies to analyses grapefruit pyrophosphate-dependent phosphofructokinase. Int. J. Biol. Macromol. 21, 307–317. Ward, N., Larsen, Q., Sakwa, J., Bruseth, L., et al. (2004). Genomic insights into methanotrophy: The complete genome sequence of Methylococcus capsulatus (Bath). PLoS Biol. 2, 1616–1628. 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. Wood, H. G., O’Brien, W. E., and Michaels, G. (1977). Properties of carboxytransphosphorylase: Pyruvate phosphate dikinase; pyrophosphate-phosphofructokinase and pyrophosphate-acetate kinase and their roles in the metabolism of inorganic pyrophosphate. Adv. Enzymol. 45, 85–155.

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

16 17 22 23 23 24 25 26 27 27 28

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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Alexander S. Reshetnikov et al.

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

Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs

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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; Garcıá-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

18


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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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.


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

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


: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

20


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


21

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

22


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


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

24


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


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

26


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


27

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

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

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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

Facultative and Obligate Methanotrophs: How to Identify and Differentiate Them

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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; MgSO47H2O, 1; Na2HPO4 12H2O, 0.717; KH2PO4, 0.272; CaCl26H2O, 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; MgSO47H2O, 0.05; CaCl22H2O, 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; ZnSO47H2O, 0.01; MnCl24H2O, 0.003; CoCl26H2O, 0.02; CuSO45H2O, 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 ( 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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C H A P T E R

S E V E N

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

100 101 101 110 112 116 116 116

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

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99

100

David S. Ojala et al.

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.

Genetic Tools for Methylomicrobium spp.

101

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

107


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


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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.

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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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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, 50; streptomycin, 2; gentamicin, 1; spectinomycin, 5; tetracycline, 0.5; neomycin, 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

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EDTA

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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

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40

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80 100 120 Time (min)

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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

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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.

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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

212 213 214 215 215 216 216 219 219 220 221 221 221 222 223 223 224 224 224

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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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

pMMO from M. trichosporium OB3b

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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 FeSO47H2O 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 CuSO46H2O 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

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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

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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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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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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

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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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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 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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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

15 Days

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 Napoleaõ, 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 { {


#


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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

30 0.2

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0 0

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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

Culture

P2

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Filtrate

HP20

P1

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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).


A

60% Acetonitrile

B

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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

1084.06 1215.09

0.6

Unfractionated 541.53 607.04

TIC

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0.4 0.2 0.0 500

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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

0

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10 Time (min)

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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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Sample A (94%:6%)

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B N ple ) Sam :25% % Sample B (75 (48%:52%)

80 Species 1(%)

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40

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0 250

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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., Fernańdez, 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

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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 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 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

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