METHODS IN ENZYMOLOGY Editors-in-Chief
JOHN N. ABELSON AND MELVIN I. SIMON Division of Biology California Institute of ...
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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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Ludmila Chistoserdova Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Dong W. Choi Department of Biological and Environmental Science, Texas A&M UniversityCommerce, Commerce, Texas, USA Andrew Crombie School of Life Sciences, University of Warwick, Coventry, United Kingdom Svetlana N. Dedysh Winogradsky Institute of Microbiology, Russian Academy of Sciences, Moscow, Russia Alan A. DiSpirito Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA Peter F. Dunfield Department of Biological Sciences, University of Calgary, Calgary, Alberta, Canada Anne Fjellbirkeland Department of Molecular Biology, and Department of Biology, Centre for Geobiology, University of Bergen, Bergen, Norway Warren H. Gallagher Department of Chemistry, University of Wisconsin-Eau Claire, Eau Claire, Wisconsin, USA Valerie S. Gilles Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University, Ames, Iowa, USA David W. Graham School of Civil Engineering and Geosciences, Newcastle University, Newcastle Upon Tyne, United Kingdom Scott C. Hartsel Department of Chemistry, University of Wisconsin-Eau Claire, Eau Claire, Wisconsin, USA Harald B. Jensen Department of Molecular Biology, University of Bergen, Bergen, Norway Marina G. Kalyuzhnaya Department of Microbiology, University of Washington, Seattle, Washington, USA
Contributors
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Odd A. Karlsen Department of Molecular Biology, University of Bergen, Bergen, Norway Valentina N. Khmelenina Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Ralf Kiese Institute for Meteorology and Climate Research, Atmospheric Environmental Research (IMK-IFU), Karlsruhe Institute of Technology (KIT), GarmischPartenkirchen, Germany Hyung J. Kim Departments of Medicine and Biochemistry, University of Utah Health Sciences Center, Salt Lake City, Utah, USA Michael C. Konopka Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Stephan M. Kraemer Department of Environmental Geosciences, University of Vienna, Althanstrasse, Vienna, Austria Øivind Larsen Department of Molecular Biology, and Uni Environment, University of Bergen, Bergen, Norway Mary E. Lidstrom Department of Chemical Engineering, and Department of Microbiology, University of Washington, Seattle, Washington, USA Chunyan Liu Institute of Atmospheric Physics, Chinese Academy of Sciences (IAP-CAS), Beijing, China Sarah McQuaide Department of Electrical Engineering, University of Washington, Seattle, Washington, USA Akimitsu Miyaji Department of Environmental Chemistry and Engineering, Tokyo Institute of Technology, Nagatsuta-cho, Midori-ku, Yokohama, Japan J. Colin Murrell School of Life Sciences, University of Warwick, Coventry, United Kingdom Ildar I. Mustakhimov Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia
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H.-Hoa T. Nguyen Transmembrane BioSciences, Pasadena, California, USA David S. Ojala Department of Chemical Engineering, University of Washington, Seattle, Washington, USA Alexander S. Reshetnikov Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Amy C. Rosenzweig Department of Molecular Biosciences, and Department of Chemistry, Northwestern University, Evanston, Illinois, USA Olga N. Rozova Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Jeremy D. Semrau Department of Civil and Environmental Engineering, The University of Michigan, Ann Arbor, Michigan, USA Stephen M. Smith Department of Molecular Biosciences, and Department of Chemistry, Northwestern University, Evanston, Illinois, USA Thomas J. Smith Biomedical Research Centre, Sheffield Hallam University, Sheffield, United Kingdom Yuri A. Trotsenko Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Sukhwan Yoon Department of Civil and Environmental Engineering, The University of Michigan, Ann Arbor, Michigan, USA, and Department of Biogeochemistry, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany Steve S.-F. Yu Institute of Chemistry, Academia Sinica, Taipei, Taiwan
PREFACE
The production and consumption of methane by microorganisms is central to the global carbon cycle. It has been 2 decades since a Methods in Enzymology volume focused explicitly on this field (Volume 188, Hydrocarbons and Methylotrophy). During that time, interest in methane metabolism has steadily increased in the context of dwindling petroleum reserves, increased greenhouse gas emissions, and environmental hydrocarbon pollution. A field previously dominated by microbiology and protein biochemistry has exploded in multiple directions. In particular, the advent of genomic and proteomic techniques has transformed the way methane metabolic pathways are studied. In these volumes, we cover both the generation (Part A) and utilization (Part B) of methane. Part A describes recent developments that enable a wide variety of experiments with methanogenic Archaea, which seemed intractable two decades ago to all but those initiates who “grew up” studying anaerobes. Methods are presented to readily culture and to perform genetic experiments on these oxygen-sensitive microbes, as well as to characterize the remarkable enzymes and respiratory proteins that allow methanogens to generate energy for growth and produce as a byproduct a very important fuel that may be adopted as the “fuel of the future.” Included is the state of the art in “biomethanation,” the biotechnological use of methanogens to produce this important energy source. Finally, approaches are described for the deployment of “omics” technologies to understand how methanogens regulate metabolism. The first methanotroph genome sequence was completed in 2004, and research has become progressively more “omic” in the past few years. Part B combines traditional approaches to methanotroph isolation and enzyme chemistry with state-of-the-art genomic and proteomic techniques. Novel methods have been developed and used to address challenging problems such as linking metagenomic data with environmental function or generating mutant forms of the methane monooxygenase enzymes. Moreover, whole new areas of research, such as the study of the copper chelator methanobactin, have been discovered. Taken together, we hope that the two volumes capture the excitement that pervades this rapidly developing field. We thank an outstanding group of colleagues with diverse points of view for providing ideas on content and ultimately contributing a series of excellent chapters. The high level of enthusiasm in the community resulted in the xv
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unexpected final production of two, rather than one, volumes. The methods and novel approaches described here should inspire and guide future research in the field as well as provide a central resource for researchers interested in methane metabolism. AMY C. ROSENZWEIG AND STEPHEN W. RAGSDALE
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
Methods in Enzymology
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
Methods in Enzymology
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
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VOLUME 381. Oxygen Sensing Edited by CHANDAN K. SEN AND GREGG L. SEMENZA VOLUME 382. Quinones and Quinone Enzymes (Part B) Edited by HELMUT SIES AND LESTER PACKER VOLUME 383. Numerical Computer Methods (Part D) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 384. Numerical Computer Methods (Part E) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 385. Imaging in Biological Research (Part A) Edited by P. MICHAEL CONN VOLUME 386. Imaging in Biological Research (Part B) Edited by P. MICHAEL CONN VOLUME 387. Liposomes (Part D) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 388. Protein Engineering Edited by DAN E. ROBERTSON AND JOSEPH P. NOEL VOLUME 389. Regulators of G-Protein Signaling (Part A) Edited by DAVID P. SIDEROVSKI VOLUME 390. Regulators of G-Protein Signaling (Part B) Edited by DAVID P. SIDEROVSKI VOLUME 391. Liposomes (Part E) Edited by NEJAT DU¨ZGU¨NES¸ VOLUME 392. RNA Interference Edited by ENGELKE ROSSI VOLUME 393. Circadian Rhythms Edited by MICHAEL W. YOUNG VOLUME 394. Nuclear Magnetic Resonance of Biological Macromolecules (Part C) Edited by THOMAS L. JAMES VOLUME 395. Producing the Biochemical Data (Part B) Edited by ELIZABETH A. ZIMMER AND ERIC H. ROALSON VOLUME 396. Nitric Oxide (Part E) Edited by LESTER PACKER AND ENRIQUE CADENAS VOLUME 397. Environmental Microbiology Edited by JARED R. LEADBETTER VOLUME 398. Ubiquitin and Protein Degradation (Part A) Edited by RAYMOND J. DESHAIES
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VOLUME 399. Ubiquitin and Protein Degradation (Part B) Edited by RAYMOND J. DESHAIES VOLUME 400. Phase II Conjugation Enzymes and Transport Systems Edited by HELMUT SIES AND LESTER PACKER VOLUME 401. Glutathione Transferases and Gamma Glutamyl Transpeptidases Edited by HELMUT SIES AND LESTER PACKER VOLUME 402. Biological Mass Spectrometry Edited by A. L. BURLINGAME VOLUME 403. GTPases Regulating Membrane Targeting and Fusion Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 404. GTPases Regulating Membrane Dynamics Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 405. Mass Spectrometry: Modified Proteins and Glycoconjugates Edited by A. L. BURLINGAME VOLUME 406. Regulators and Effectors of Small GTPases: Rho Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 407. Regulators and Effectors of Small GTPases: Ras Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 408. DNA Repair (Part A) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 409. DNA Repair (Part B) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 410. DNA Microarrays (Part A: Array Platforms and Web-Bench Protocols) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 411. DNA Microarrays (Part B: Databases and Statistics) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 412. Amyloid, Prions, and Other Protein Aggregates (Part B) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 413. Amyloid, Prions, and Other Protein Aggregates (Part C) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 414. Measuring Biological Responses with Automated Microscopy Edited by JAMES INGLESE VOLUME 415. Glycobiology Edited by MINORU FUKUDA VOLUME 416. Glycomics Edited by MINORU FUKUDA
Methods in Enzymology
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Characterization of the Recombinant PyrophosphateDependent 6-Phosphofructokinases from Methylomicrobium alcaliphilum 20Z and Methylococcus capsulatus Bath Valentina N. Khmelenina, Olga N. Rozova, and Yuri A. Trotsenko Contents 1. Introduction 2. Methods 2.1. Culture conditions 2.2. Routine genetic manipulations 2.3. PPi-PFKs cloning, expression, and purification 2.4. Assay of pyrophosphate-dependent 6-phosphofructokinase activity 2.5. Assay of fructosebisphosphate aldolase activity by using the recombinant PPi-PFK 3. Brief Overview of Current Knowledge of PPi-PFK in Microorganisms 4. Properties of PPi-PFK from Methanotrophs 4.1. PPi-PFK from Mc. capsulatus Bath 4.2. PPi-PFK from Mm. alcaliphilum 20Z 5. Conclusion Acknowledgments References
2 4 4 4 4 4 6 7 8 8 10 10 11 11
Abstract The Embden–Meyerhof–Parnas (EMP) glycolysis is the starting point of the core carbon metabolism. Aerobic methanotrophs possessing activity of the pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK) instead of the classical glycolytic enzyme ATP-dependent 6-phosphofructokinase (ATP-PFK) are Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00001-2
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2011 Elsevier Inc. All rights reserved.
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promising model bacteria for elucidation of the role of inorganic pyrophosphate (PPi) and PPi-dependent glycolysis in microorganisms. Characterization of the His6-tagged PPi-PFKs from two methanotrophs, halotolerant alkaliphilic Methylomicrobium alcaliphilum 20Z and thermotolerant Methylococcus capsulatus Bath, showed differential capabilities of PPi-PFKs to phosphorylate sedoheptulose-7phosphate and this property correlated well with the metabolic patterns of these bacteria assimilating C1 substrate either via the ribulosemonophosphate (RuMP) pathway (Mm. alcaliphilum 20Z) or simultaneously via the RuMP and serine pathways and the Calvin cycle (Mc. capsulatus Bath). Analysis of the genomic draft of Mm. alcaliphilum 20Z (https://www.genoscope.cns.fr/agc/mage) has provided in silico evidence for the existence of a PPi-dependent pyruvate-phosphate dikinase (PPDK). Expression of the ppdk gene at oxygen limitation along with the presence of PPi-PFK in Mm. alcaliphilum 20Z implied functioning of PPi-dependent glycolysis and PPi recycling under conditions when oxidative phosphorylation is hampered.
1. Introduction Energy metabolism of many organisms is based on glycolysis. In aerobic microorganisms, glycolysis is a minor source of energy, since ca. 95% of ATP production comes from subsequent tricarboxylic acid cycle reactions and oxidative phosphorylation. In contrast, anaerobic organisms rely almost exclusively on glycolysis and fermentation for ATP production (Mertens, 1991; Mertens et al., 1993). One significant variation of the standard glycolytic pathway is pyrophosphate-dependent glycolysis, which uses PPi instead of ATP as a phosphoryl donor (Mertens, 1991; Saavedra et al., 2005). In this version of glycolysis, two key glycolytic enzymes, ATP-PFK and ADP-pyruvate kinase (PK), are replaced by pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK) and pyruvate phosphate dikinase (PPDK) the latter catalyzing the reversible reaction of AMP- and PPi-dependent conversion of phosphoenolpyruvate (PEP) into pyruvate accompanied by ATP synthesis (Mertens, 1991; Mertens et al., 1993). PPi-dependent glycolysis can in theory yield five ATP molecules instead of the two yielded by standard glycolysis. This increase of 2.5-fold can be crucial under severe energy-limiting conditions. This modified glycolysis has been well studied in anaerobic bacteria and protists (Saavedra et al., 2005). Apart from the widely distributed ATP-PFK catalyzing an irreversible catabolic reaction, the phosphorylation of fructose-6-phosphate (Fru 6-P) to fructose-1,6-bisphosphate (Fru 1,6-P2), PPi-PFK (EC 2.7.1.90) catalyzes the same reaction in reversible way and can thus function both in glycolysis and gluconeogenesis (Reeves et al., 1974, 1976).
Pyrophosphate-Dependent 6-Phosphofructokinase from Methanotrophs
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Mg2þ
Fructose-6-phosphate þ PPi > Fructose-1; 6-bisphosphate þ Pi
PPi-PFK activity was detected in several protozoans, in higher plants and in prokaryotes (Bruchhaus et al., 1996; Carnal and Black, 1983; Mertens et al., 1993; O’Brien et al., 1975; Petzel et al., 1989). In many organisms, PPi-PFK replaces allosteric ATP-PFK which catalyses an essentially irreversible catabolic reaction of classical glycolysis (Slamovits and Keeling, 2006; Wood et al., 1977). In some cases, PPi-PFK is present alongside ATP-PFK in the same organism, suggesting that PPi-PFK may perform an unknown specific function (Alves et al., 1994, 2001; Van Praag, 1997). Activity of PPi-PFK was found in 10 strains of types I, II, and X aerobic methanotrophic bacteria (Beschastny et al., 1992, 2008; Trotsenko and Shishkina, 1990). Also, intracellular PPi concentrations 20 times higher than in Escherichia coli cells and only low ATP concentration were found in the methanotrophs studied and such metabolic features correlated with extremely low inorganic pyrophosphatase activity (Beschastny et al., 2008; Chetina and Trotsenko, 1987; Trotsenko and Shishkina, 1990). The role of PPi and PPi-dependent enzymes in methanotrophs is not obvious. It is generally accepted that aerobic methanotrophs oxidize methane to carbon dioxide and water via the intermediates of methanol, formaldehyde, and formate, thereby providing energy for biosynthesis of cell components (Trotsenko and Murrell, 2008). ATP production is thought to come from oxidative phosphorylation but with low efficiency in methanotrophs studied (Chetina and Trotsenko, 1987). ATP production relying on substratelevel phosphorylation may play a crucial role in these bacteria. Methanotrophs are different in their constructive metabolic pattern. Type I methanotrophic bacteria assimilate C1 compounds via the ribulosemonophosphate (RuMP) cycle of formaldehyde fixation, in which the first intermediates are C6 phosphosugars. Phosphotrioses are formed by cleavage of phosphohexoses through both the Entner–Doudoroff and Embden– Meyerhof–Parnas (EMP) pathways (Shishkina and Trotsenko, 1982; Stro¨m et al., 1974). In contrast, type II methanotrophs use the serine pathway of C1 assimilation, where C3 compounds are firstly formed after condensation of formaldehyde and glycine (Stro¨m et al., 1974; Trotsenko and Murrell, 2008); therefore, C6 phosphosugar synthesis via gluconeogenesis reactions is needed. Notably, simultaneous functioning of three pathways: the RuMP cycle as the major pathway, the serine pathway and the Calvin cycle was detected for the type X methanotrophs Methylococcus capsulatus Bath and Methylocaldum szegediense (Medvedkova et al., 2009; Trotsenko and Murrell, 2008). Operation of the Calvin cycle may be related to the thermophilic nature of these methanotrophic species (Trotsenko et al., 2009). Thus, in type I and type X methanotrophs, both C3 phosphotrioses and hexosephosphates can be formed bypassing the classical EMP
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pathway. This suggests that glycolysis and the PPi-dependent PFK may perform unknown specific function(s). Availability of the genome sequences of several methanotrophs, including halotolerant Mm. alcaliphilum 20Z (https:// www.genoscope.cns.fr/agc/mage) and thermotolerant Mc. capsulatus Bath (Ward et al., 2004), has generated the required template to investigate in detail their metabolic pattern at both genetic and enzymatic levels. In this study, we describe properties of purified PPi-PFKs from Mm. alcaliphilum 20Z and Mc. capsulatus Bath.
2. Methods 2.1. Culture conditions Cells of Mm. alcaliphilum were grown under methane-air atmosphere (1:1) in liquid mineral medium containing 3% NaCl (Khmelenina et al., 1999). The standard NMS medium (Whittenbury et al., 1970) was used for Mc. capsulatus Bath cultivation at 43 C.
2.2. Routine genetic manipulations DNA from Mm. alcaliphilum 20Z and Mc. capsulatus Bath was isolated by a standard phenol:chloroform method (Sambrook and Russell, 2001). For RNA isolation from 10 ml of exponentially grown culture (OD600 ¼ 0.8), the RNA extraction method of Chomczynski and Sacchi (1987) was used as described in Reshetnikov et al. (2006).
2.3. PPi-PFKs cloning, expression, and purification The standard procedures may be used for cloning of the pfp genes into the bacterial expression vector pET22b designed to express a C-terminal His6tagged fusion protein under the control of the T7 promoter. A single step of immobilized metal ion affinity chromatography on a Ni2þ-NTA column was performed for purification of His6-tagged PPi-PFK fusion protein from E. coli extracts (Reshetnikov et al., 2008).
2.4. Assay of pyrophosphate-dependent 6-phosphofructokinase activity 2.4.1. Assay of PPi-PFK with Fru 6-P as phosphoryl acceptor 1 ml of reaction mixture contained: 50 mM HEPES–HCl, pH 7.0; 20 mM Fru 6-P;
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2 mM PPi; 5 mM MgCl2; 0.25 mM NADH; 0.2 U of aldolase (from rabbit muscle); 0.3 U of a-glycerophosphate dehydrogenase; 0.3 U of triosephosphate isomerase (Sigma). After preincubation at 30 C for 3 min, 1–2 mg of PPi-PFK was added and the NADH oxidation rate was measured by the decrease in absorbance at 340 nm. 2.4.2. Assay of PPi-PFK with sedoheptulose-7-phosphate as phosphoryl acceptor 1 ml of reaction mixture contained: 50 mM HEPES–HCl, pH 7.0 5 mM sedoheptulose-7-phosphate; 2 mM PPi; 5 mM MgCl2; 0.25 mM NADH; 0.2 U of aldolase (from rabbit muscle); 0.3 U of a-glycerophosphate dehydrogenase; After preincubation at 30 C for 3 min, 1–2 mg of PPi-PFK was added and the NADH oxidation rate was measured by the decrease in absorbance at 340 nm. 2.4.3. Assay of PPi-PFK with Fru 1,6-P2 1 ml of reaction mixture contained: 50 mM HEPES–HCl, pH 7.0; 2 mM Fru 1,6-P2; 15 mM NaH2PO4; 5 mM MgCl2; 0.3 mM NADPþ; 0.12 U of glucose-6-phosphate dehydrogenase; 0.24 U of glucose phosphate isomerase. Incubate at 30 C for 3 min. Add 1 mg of PPi-PFK. Monitor NADPþ reduction at 340 nm. 2.4.4. Assay of PPi-PFK with sedoheptulose-1,7-bisphosphate or ribulose-1,5-bisphosphate 0.4 ml reaction mixture contained: 50 mM HEPES–HCl, pH 7.0;
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mM sedoheptulose-1,7-bisphosphate (or 3 mM ribulose-1,5bisphosphate); 15 mM NaH2PO4; 5 mM MgCl2. The mixture was incubated for 3 min at 30 C and reaction was stopped by addition of 0.4 ml of 1 M H2SO4. Estimation of PPi (Heinonen et al., 1981): 0.4 ml of reagent consisting of 4 ml of 40 mM ammonium heptamolybdate, 1 ml of 5 M H2SO4, 50 mM of triethylamine is added to the mixture and let stand for at least 15 min and then centrifuged 5 min at 2000g. Then 0.2 ml of 5 M H2SO4 was added to the supernatant and centrifuged at the same condition. To the second supernatant 0.08 ml of 1 M 2-mercaptoethanol was added by mixing and the color developed for 15 min. The absorbance at a wavelength of 700 nm was measured. PPi concentration was calculated from a calibration line made with Na4P2O7 standard solutions (5–200 nmol in 0.4 ml of H2O).
2.5. Assay of fructosebisphosphate aldolase activity by using the recombinant PPi-PFK Activity of the fructosebisphosphate aldolase (FBA) toward the Fru 1,6-P2 synthesis was rarely studied due to the complexity of Fru 1,6-P2 detection. This problem may be resolved by using the highly active preparation of the recombinant PPi-PFK as the coupling enzyme (Rozova et al., 2010a). 1 ml of reaction mixture contained: 50 mM Tris–HCl, pH 7.5; 2 mM dihydroxyacetone-3-phosphate; 2 mM glyceraldehyde-3-phosphate; 1 mM NADPþ; 1 mM NaH2PO4; 4 mM MgCl2; 0.5 U of glucose phosphate isomerase (Sigma); 0.5 U of glucose-6-phosphate dehydrogenase (from rabbit muscle, Sigma); 0.5 U of recombinant PPi-PFK from Mm. alcaliphilum 20Z or Methylomonas methanica (Reshetnikov et al., 2005). After preincubation at 30 C for 3 min, 15 mU of FBA was added and the NADPþ reduction rate at 340 nm was measured.
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3. Brief Overview of Current Knowledge of PPi-PFK in Microorganisms PPi-PFK was first found in the anaerobic amitochondrial protozoan parasite Entamoeba histolytica H200 (Reeves et al., 1974, 1976). Subsequently, PPi-PFK activity was detected in other protozoans, in higher plants and in prokaryotes (Bruchhaus et al., 1996; Carnal and Black, 1983; Mertens et al., 1993; Petzel et al., 1989). Two types of PPi-PFK are known. Activity of type I PPi-PFK, which is a homopolymer with a subunit molecular mass of 40–50 kDa, is independent on fructose-2,6-bisphosphate (Fru 2,6-P2) and this enzyme has been found in protists and prokaryotes. Type II PPiPFK is a heterodimer, stimulated by Fru 2,6-P2 and found in photosynthetic organisms, higher plants and Euglena gracilis (Bruchhaus et al., 1996; Miyatake et al., 1984). The first report of the presence of PPi-PFK in the prokaryotic organism Propionibacterium freudenreichii appeared in 1975 (O’Brien et al., 1975). The enzyme (2 48 kDa) functions in either glycolysis or gluconeogenesis depending on growth substrate (glucose vs. glycerol or lactate). Synthesis of lipids, carbohydrates, proteins, nucleic acids, or NADH-dependent reduction of fumarate into succinate connected with the electron transport chain were proposed as sources of PPi for this enzyme (O’Brien et al., 1975). PPi-PFK has no allosteric effectors with the exception of the enzyme from Rhodospirillum rubrum, which was inhibited by AMP, ADP, and ATP (Pfleiderer and Klemme, 1980). In many cases, the physiological role of PPi-PFK is not obvious (Siebers et al., 1998). More precisely, the simultaneous presence of PPi-PFK and ATP-PFK (i.e., of a reversible and an irreversible enzyme) in a single organism, even if rarely reported, has suggested that PPi-PFK may perform an alternative unknown function (Alves et al., 2001; Van Praag, 1997). ATP-PFK and PPi-PFK share a common ancestry, but phylogenies show a very complex evolutionary history (Bapteste et al., 2003; Siebers et al., 1998). Several amino acids are essential for ATP-PFK and PPi-PFK functions (Moore et al., 2002). PFK working with ATP (with some exception) harbors Gly at positions 104 and 124 (according to the numbering of E. coli ATP-PFK), while PPi-PFK has an Asp104 and a Lys124. A singlepoint mutation can induce a change of the phospho-donor (Chi and Kemp, 2000). In addition to possible convergent adaptive mutations, horizontal gene transfer (HGT) events, including those of mutated sequences, further complicate the distribution of the enzymes. Hence, based on the PFK phylogenies it is not possible to conclude which phosphoryl donor, ATP or PPi, was ancestrally used by PFK (Bapteste et al., 2003).
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4. Properties of PPi-PFK from Methanotrophs PPi-PFK from Mc. capsulatus Bath is homodimeric (2 45), similar to the most known eubacterial enzymes, and that from Mm. alcaliphilum 20Z is homotetrameric (4 45). PPi-PFKs from both methanotrophs are not subjected to allosteric activation by organic compounds (ATP, ADP, AMP, NADP, NAD, and NADPH) and are thus probably regulated exclusively at the level of substrate and product concentrations (Reshetnikov et al., 2008; Rozova et al., 2010b). ATP, ADP, AMP, CTP, CDP, CMP, UTP, UDP, UMP, GTP, GDP, GMP at 1 mM as well as polyphosphates (n ¼ 3, n ¼ 9, n ¼ 15, and n ¼ 45) at 0.1 mg ml 1 could not substitute for PPi.
4.1. PPi-PFK from Mc. capsulatus Bath The enzyme from Mc. capsulatus Bath has the lowest affinity for both Fru 6-P and Fru 1,6-P2 in comparison to all known bacterial PPi-PFKs (Reshetnikov et al., 2008). It also reversibly phosphorylates Se 7-P with much higher activity and affinity and also displays activity with ribulose-5-P and ribulose-1,5-P2 (Table 1.1). Based on these properties of the enzyme, Table 1.1 Properties of PP-PFK from methanotrophs
Parameters
Vmax (U/mg) F-6-P S-7-P FBP Km (mM) PPi F-6-P S-7-P FBP Pi Mg2þ (forward reaction) Mg2þ (reverse reaction) Mr (kDa) (subunit number) Optimum pH Optimum temperature ( C) Effectors
Methylomicrobium alcaliphilum 20Z (Rozova et al., 2010b)
Methylomonas methanica 12 (Beschastny et al., 1992)
Methylococcus capsulatus Bath (Reshetnikov et al., 2008)
577 0.18 805
840 n.d. 850
7.6 31 9.0
0.118 0.64 1.01 0.095 3.4 0.22 0.33 180 (4 45) 7.5 30 No
0.051 0.39 n.d. 0.1 1.7 0.038 0.35 92 (2 45) 8.0 40 No
0.027 2.27 0.030 0.328 8.69 0.028 2 90 (2 45) 7.0 30 No
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its involvement in the nonoxidative pentose phosphate segment of the Calvin cycle, in which sedoheptulose and ribulose mono- and bisphosphates are the obligatory intermediates, may be proposed. The use of Se 7-P as a substrate was earlier shown for the E. histolytica PPi-PFK (Susskind et al., 1982). However, E. histolytica PPi-PFK has a lower affinity for Se 7-P (Km ¼ 64 mM) than for Fru 6-P (Km ¼ 38 mM). 4.1.1. Transcriptional organization of pfp and hpp genes in Mc. capsulatus Bath and their phylogeny The gene pfp encoding PPi-PFK and the gene hpp (MCA1252) encoding a putative V-type Hþ-pyrophosphatase (Hþ-PPi-ase, EC 3.6.1.1) are colocated in chromosome of Mc. capsulatus Bath (Ward et al., 2004). The closest homologues of the Hþ-PPi-ase is a protein from Geobacter sulfurreducens (TIGR_35554j2947) and R. rubrum (AAC38615) (62% and 58% identities). Cotranscription of the pfp and hpp genes was demonstrated by a standard RT-PCR approach (Reshetnikov et al., 2008). Putative promoter sequences (10 region TAAGTT; 35 region TTGTAA) were revealed 30 bp upstream of the start codon of pfp. According the accepted classification (Bapteste et al., 2003; Mu¨ller et al., 2001), PPi-PFK from Mc. capsulatus is clustered in clade B2 of the type II PFK enzymes and this clade is represented by eubacterial PPi-PFKs only. The most closely related neighbors of the Mc. capsulatus PPi-PFK are the enzymes from lithoautotrophic ammonia oxidizers bacteria Nitrosomonas europaea (Klotz et al., 2006) and Nitrosospira multiformis (Norton et al., 2008) and the facultative b-proteobacterial methylotroph Methylibium petroleiphilum PM1 (Kane et al., 2007). Similarly to Mc. capsulatus, the genomes of these bacteria also contain genes for ribulose bisphosphate carboxylase. Since no genes encoding ATP-PFK or fructose bisphosphatase (FBPase, EC 3.1.3.11) were revealed in the genome of Mc. capsulatus Bath (Ward et al., 2004), the single reversible PPi-PFK may fulfill the functions of these enzymes. Importantly, the bacteria containing PPi-PFK highly homologous to Mc. capsulatus Bath enzyme exhibit common biochemical features. They use monooxygenases requiring NADH or reduced cytochromes for energy substrate oxidation: methane monooxygenase (in methanotrophs), ammonia monooxygenase (in Nitrosospira and Nitrosomonas species), or toluene/ benzene monooxygenase (in M. petroleiphilum PM1) (Kane et al., 2007). Thus, oxidative phosphorylation may be hampered due to limiting by NADH or reduced cytochromes. Therefore, the use of PPi instead of ATP in some metabolic reactions may economize ATP and provide some advantages. Notably, in the lithoautotrophic bacteria, the pfp and hpp genes are also colocated in chromosomes. Coexpression of PPi-PFK and the putative Hþ-PPase suggests the hypothesis that the source of PPi for Fru 6-P phosphorylation may be energy-dependent PPi synthesis on the cell membrane. Conversely, PPi-PFK functioning in the reverse direction
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produces PPi, which may be utilized for generation of a Hþ gradient across the membrane, supporting ATP synthesis (Baltschevsky et al., 1999). The Mc. capsulatus PPi-PFK is clearly separated from a phylogenetic group “P” of PPi-PFKs from P. freudenreichii (O’Brien et al., 1975), Mm. alcaliphilum 20Z and M. methanica 12 (Reshetnikov et al., 2005) (12.6% and 16.5% identity, respectively).
4.2. PPi-PFK from Mm. alcaliphilum 20Z PPi-PFK from Mm. alcaliphilum 20Z showed very high activities and affinities for the forward and reverse reaction substrates, and could be involved in both glycolysis and gluconeogenesis (Rozova et al., 2010b). It appears to transform Fru 1,6-P2 more efficiently than Fru 6-P suggesting that a role of the enzyme in gluconeogenesis may be physiologically more significant. It also phosphorylates Se 7-P but with 103 times less activity than with Fru 6-P. In Mm. alcaliphilum 20Z, C3 compounds for biosynthetic needs (pyruvate and phosphotrioses) can be generated from C6 phosphosugars (the first products of the RuMP cycle) by cleavage via the Entner–Doudoroff pathway (Khmelenina et al., 1997, 2010). Thus, both C3 and C6 compounds can be formed bypassing the EMP pathway. PPi-PFK, together with FBA, may participate in regulating the ratio of C6 and C3 compounds in the cell. Since a candidate gene for the fructose-1,6-bisphosphatase (FBPase) is also present in the genome, some redundancy of the gluconeogenic activity occurs in Mm. alcaliphilum 20Z. Simultaneous functioning of PPi-PFK and FBPase may comprise a futile cycle for removing excess PPi which is the byproduct of many anabolic reactions including synthesis of sucrose accumulated by this halotolerant methanotroph at high levels in high salinity growth conditions (Khmelenina et al., 1999). The differential metabolic properties and functions of PPi-PFKs from Mm. alcaliphilum 20Z and Mc. capsulatus Bath correlate with their large sequence differences (16.5% identity), suggesting the possibility of an independent origin of these enzymes, along the lines of the hypothesis of “rampant” horizontal transfer of genes coding for microbial phosphofructokinases (Bapteste et al., 2003; Mu¨ller et al., 2001).
5. Conclusion Inorganic pyrophosphate created during biosynthetic polymerization reactions in most organisms is hydrolyzed by inorganic pyrophosphatase in order to thermodynamically favor the anabolic processes. Studied methanotrophs are characterized by negligible activity of inorganic
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pyrophosphatase and use PPi as the phosphoryl donor at least in the key glycolytic reaction phosphorylating Fru 6-P. Based on differential properties of PPi-PFKs from two methanotrophs and analyses of the recent genomic data, several important functions may be proposed for the enzyme: involvement in PPi recycling to economize ATP synthesis, regulation of hexosephosphates and triosephosphates interconversion as well as PPi levels in cells (in type I methanotrophic bacteria) and additionally in the Calvin cycle operation (in the type X methanotroph Mc. capsulatus Bath). Moreover, a gene encoding for PPDK catalyzing the PPi- and AMP-dependent conversion of PEP to pyruvate accompanied by ATP formation is present in the genomes of Mm. alcaliphilum 20Z and Mc. capsulatus Bath. Transcriptome analysis of Mm. alcaliphilum 20Z culture indicated that expression of PPDK increased under conditions of oxygen limitation (Kalyuzhnaya, personal communication). Thus, PPi-dependent glycolysis involving the two PPi-dependent enzymes, PPi-PFK and PPDK, may be proposed for this methanotroph. These two enzymes make glycolysis fully reversible. Importantly, PK and PEP synthetase encoding genes can also be expressed in this Mm. alcaliphilum 20Z. High flexibility at the key glycolytic steps may provide some advantages for methanotrophs, allowing metabolism to adapt to different energy states of cell. However, these hypotheses must be verified experimentally, particularly by characterizing the properties of the respective proteins. Further studies for unraveling the evolutionary history of PPi-dependent enzymes of glycolysis and the physiological role of PPi in aerobic methanotrophic bacteria are needed.
ACKNOWLEDGMENTS The work was supported by grants RFBR (08-04-01484-a and 10-04-01224-a), CRDF (Rub1-2946-PU-09), and joint PICS CNRS grant No3380. We thank the team at Ge´noscope for high-throughput sequencing and annotation of the M. alcaliphilum 20Z genome (IbiSA-Ge´noscope 2008 campaign) and M. G. Kalyuzhnaya for kindly providing microarray analysis information.
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Genes and Enzymes of Ectoine Biosynthesis in Halotolerant Methanotrophs Alexander S. Reshetnikov, Valentina N. Khmelenina, Ildar I. Mustakhimov, and Yuri A. Trotsenko Contents 1. Introduction 2. PCR-Based Approach for Identification of the Ectoine Biosynthesis Genes in Methanotrophs 3. Transcriptional Regulation of the Ectoine Biosynthesis Genes 4. Key Enzymes of Ectoine Biosynthesis 4.1. Diaminobutyric acid aminotransferase 4.2. DABA acetyltransferase 4.3. Ectoine synthase 4.4. Aspartate kinase 5. Conclusion Acknowledgments References
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Abstract Ectoine (1,4,5,6-tetrahydro-2-methyl-4-pyrimidine carboxylic acid) is a widely distributed compatible solute accumulated by halophilic and halotolerant microorganisms to prevent osmotic stress in highly saline environments. Ectoine as a highly water keeping compound stabilizing biomolecules and whole cells can be used in scientific work, cosmetics, and medicine. Detailed understanding of the organization/regulation of the ectoine biosynthetic pathway in various producers is an active area of research. Here we review current knowledge on some genetic and enzymatic aspects of ectoine biosynthesis in halophilic and halotolerant methanotrophs. By using PCR methodology, the genes coding for the specific enzymes of ectoine biosynthesis, diaminobutyric acid (DABA) aminotransferase (EctB), DABA acetyltransferase (EctA), and ectoine synthase (EctC), were identified in several methanotrophic species. Skryabin Institute of Biochemistry and Physiology of Microorganisms, RAS, Pushchino, Moscow Region, Russia Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00002-4
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Organization of these genes in either ectABC or ectABC-ask operons, the latter additionally encoding aspartate kinase isozyme (Ask), correlated well with methanotroph halotolerance and intracellular ectoine level. A new gene, ectR1 encoding the MarR-like transcriptional regulatory protein EctR1, negatively controlling transcription of ectoine biosynthetic genes was found upstream of ectABC-ask operon in Methylomicrobium alcaliphilum 20Z. The ectR-like genes were also found in halotolerant methanol utilizers Methylophaga alcalica and Methylophaga thalassica as well as in several genomes of nonmethylotrophic species. The His6-tagged DABA acetyltransferases from Mm. alcaliphilum, M. alcalica, and M. thalassica were purified and the enzyme properties were found to correlate with the ecophysiologies of these bacteria. All these discoveries should be very helpful for better understanding the biosynthetic mechanism of this important natural compound, and for the targeted metabolic engineering of its producers.
1. Introduction Most halophilic and halotolerant microorganisms adapt to high environmental salinity by accumulation of low-molecular weight organic compounds called compatible solutes that equilibrate external osmotic pressure and support intracellular turgor that is higher than in the surroundings. The cyclic imino acid ectoine was originally discovered as an osmoprotectant in anoxygenic phototrophs of the Ectothiorhodospira group (Galinski et al., 1985) and subsequently found in many other Gram-negative and Gram-positive bacteria (Galinski, 1995; Kempf and Bremer, 1998; Severin et al., 1992), including the salt-dependent methanotrophs Methylomicrobium alcaliphilum, Mm. buryatense, Mm. kenyense, Methylobacter marinus (Kalyuzhnaya et al., 2001, 2008; Khmelenina et al., 1999, 2010; Trotsenko and Khmelenina, 2002) as well as methanol- and methylamine-utilizing bacteria of genera Methylophaga and Methylarcula (Doronina et al., 1998, 2000, 2003a,b). Ectoine is a compound compatible with cell metabolism even if accumulated to high intracellular concentrations. It has also been shown to act as a chemical chaperone increasing the stability of proteins. The mechanism of such stabilization can provide insights into protein folding. Moreover, ectoine and its hydroxylated derivative, hydroxyectoine, have attracted commercial interest for use in medicine and cosmetics as stabilizers of biomolecules and whole cells against different damaging factors such as heating, freezing, desiccation, and UV radiation (Buenger and Driller, 2004; Buenger and Driller, 2004; Graf et al., 2008; Jebbar et al., 1992; Sauer and Galinski, 1998). Since ectoines are produced only biotechnologically, understanding their biosynthetic pathways and regulation has been active area of research. The necessity of elucidation of the genes and enzymes responsible for ectoine biosynthesis in halophilic and halotolerant
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methylotrophs is conditioned by practical demands for new technologies of ectoine production from methane and methanol. The pathway of ectoine biosynthesis in the bacteria studied represents a biochemical sequence that is a branch of aspartate family amino acids synthesis (Peters et al., 1990). Three special enzymes are involved in this pathway: diaminobutyric acid (DABA) aminotransferase (EctB, EC 2.6.1.76), catalyzing transamination of aspartate semialdehyde into DABA, DABA acetyltransferase (EctA, EC 2.3.1.178), acetylating DABA into Ngacetyl-DABA, and ectoine synthase (EctC, EC 4.2.1.108), which forms ectoine by cycling of Ng-acetyl-DABA. In most halophilic bacteria studied, the genes encoding these enzymes are combined in chromosomes in the cluster ectABC (Canovas et al., 1999; Go¨ller et al., 1998; Kuhlmann and Bremer, 2002; Louis and Galinski, 1997). Database searches reveal the highly homologous ectoine biosynthetic genes in 200 finished genomes of bacteria belonging to phyla Proteobacteria, Actinobacteria, and one archaeon Nitrosopumilus maritimus. Sometimes, an additional gene ectD coding for ectoine hydroxylase is present and this enzyme converts ectoine to hydroxyectoine (Bursy et al., 2007; Garcı´a-Estepa et al., 2006; Prabhu et al., 2004). High homology of the ectoine biosynthesis genes in various microorganisms of different taxonomic position and physiological properties is a reliable indication of the evolutionary conservation of this biochemical pathway (Kuhlmann and Bremer, 2002). We present below a brief overview of current knowledge on the peculiarities of the ectoine biosynthetic pathway in aerobic halophilic and halotolerant methanotrophs, focusing on the main genetic and biochemical techniques that have been used to unravel mechanisms regulating this important natural product biosynthesis. In the first section, we highlight the strategies that are used for identification of ectoine biosynthetic genes in methanotrophs. In the second section, we describe initial results of transcriptional analysis of ect-genes. The third section briefly describes some properties of the ectoine biosynthesis enzymes. Despite the focus on the ectoine biosynthetic pathway in methanotrophs, many of the techniques could be readily adapted to the study of other ectoine producers.
2. PCR-Based Approach for Identification of the Ectoine Biosynthesis Genes in Methanotrophs Since the genes coding for the ectoine biosynthetic enzymes contain highly conserved regions and are generally organized in an ectABC operon, PCR was very helpful methodology to identify them in Mm. alcaliphilum 20Z (Reshetnikov et al., 2006). Assuming that ect-genes in other
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methanotrophs are also combined in the chromosome as an ectABC operon, we have been exploring the use of better-focused probes more specific for the ectoine biosynthetic gene cluster (Reshetnikov et al., 2006). Based on the alignment of the conserved regions of DABA aminotransferase, DABA acetyltransferase, and ectoine synthase, several degenerate primers Tra1, Tra4, Tra3, CR, and Atf for PCR have been designed and tested (Table 2.1, Fig. 2.1A). Primers Tra1 and Tra4 gave a single specific product about 750 bp in size with Mm. alcaliphilum genomic DNA as a template and this product was shown to match the predicted sequence. Degenerate primer pair Tra3/CR was used to amplify a 1000-bp fragment containing the 30 end of ectB and 50 end of ectC genes. By using another degenerate primer, Atf, corresponding to a segment of ectA and the primer Tra2.1 homologous to 50 region of ectB, a DNA fragment of 700 bp was amplified. All these PCR products were sequenced and analyzed using Vector NTI v.9 (Invitrogen). The assembled sequence of 2100 bp contained three orfs (one complete and two incomplete) oriented in the same direction. These orfs were revealed by database searches using BLAST
Table 2.1 Oligonucleotide primer sequences used for studying of ectoine biosynthesis genes in methanotrophs
Primers
Gene targeted
Sequences (50 –30 )
Tra1 Tra2 Tra2.1 Tra3 Tra4 CR C20 EF AskF AskR AskIR AskIF AtrI Atf ATZ AMOFor AMOA AMOR AMOZ AMOF 7CF 7CR
ectB ectB ectB ectB ectB ectC “vectorette” ectC ask ectB ask ask ectA ectA ectA ectA ectA ectB ectB ectC ectC ectB
CCCTIAA(T/C)TA(T/C)GGICA(T/C)AA(C/T) CC(G/A)TG(A/G)AAI(G/C)C(G/A)TTIGT(G/A)AA ATCTTCAGTGCCGCTTCAACC ACCGG(T/C)ACITT(C/T)TT(C/T)AGITT(C/T)GA CG(G/A)AAIGTGCC(A/G)TT(G/A)TG(T/C)TCNCC GGIGG(A/G)TT(A/G)AANAC(A/G)CA CTCTCCCTTCTCGAATCGTAA TGCTGCTGAAGGATGACGGTATGGGCT AGCATCTTTGTCGGCACGGTCATT CCCGCTTTGCTGACCTCTTTACTGA GCCTCATCCGCCTTGGTCAGTA TGCGTAGAGATTATTGCCGGGTGT ACAAAACCAACCAACTCATCGCCA CTIGA(T/C)(C/T)(T/C)IAA(T/C)TCI(T/G/A)(C/T)ITA TT(C/T)GT(G/T/C)TGGCA(A/G)GTIGCNGT CGAAATGCTGGTTAAGATTGGTCCGT ATTGGGCGGATTGATCGTAGTTTC ATTTGATCGGCGAAACGGTTAT CTTGAGGCTTGCCTCGGCTATC CGACGATTTATCAAGGTGCGGA CGATCAAGGCGGGCACGGTCTAT TCGACAATGGCCTGCAGACTT
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Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs
O. B. B. M. S. V. C.
iheyensis pasteurii halodurans halophilus coelicolor cholerae salexigens
:GAGALNYGHNPSEM: :GAGSLNYGHNNEKM: :GAGALNYGHNDEKM: :GAGALNYGHNDENM: :GAGSLNYGHNNAVL: :GAGALNYGHNNAVL: :GAGTLNYGHNHPKL:
O. B. B. M. S. V. C.
iheyensis pasteurii halodurans halophilus coelicolor cholerae salexigens
Tra1 (forvard) CCCTIAA(T/C)TA(T/C)GGICA(T/C)AA(C/T)
O. B. B. M. S. V. C.
iheyensis pasteurii halodurans halophilus coelicolor cholerae salexigens
:PGEHNGTFRGNN LA: :PGEHNGTFRGNN MA: :PGEHNGTFRGNN HA: :PGEHNGTFRGNN FA: :PGEHNGTFRGNN PA: :PGEHNGTFRGNN HA: :PGQYNGTFRGFNLA:
Tra4 (reversed) CG(A/G)AAIGTGCC(G/T)TT(A/G)TG(C/T)TCNCC
: : : : : : :
GRTGTFFSFEEAGINPD GRTGTFFSFEPAGIQPD GRTGTFFSFEDAGITPD GRTGTFFSFEPAGIKPD GRTGAFFSFEEAGITPD GRTGTFFSFEPSGIEPD GRTGKFFSFEHAGITPD
Tra3 (forvard) ACCGG(T/C)ACITT(C/T)TT(C/T)AGITT(C/T)GA
M. B. B. V. C. S. O.
halophilus : halodurans : pasteurii : cholerae : salexigens : coelicolor : iheyensis :
MVCVFNPPL MVCVFNPPI LICTFNPPL MACVFNPPL VACVFNPAL CICVFNPPV MVCVFNPAL
CR (reversed)
GGIGG(G/A)TT(G/A)AANAC(G/A)CA
Figure 2.1 Conserved regions of DABA aminotransferase (EctB) and ectoine synthase (EctC) homologues and degenerate primers designed for ectB (Tra1, Tra3, and Tra4) and ectC (CR) genes.
to encode proteins with high amino acid sequence identities to the ectoine biosynthesis enzymes: DABA acetyltransferase (EctA), DABA aminotransferase (EctB), and ectoine synthase (EctC). Then, the newly obtained sequences were used to design primers AtrI (homologous to ectA) and EF (homologous to ectC) which were used in inverse PCR (IPCR) to clone the missing 50 end of ectA and 30 end of ectC. In the PCR product of 1.8 kb, a new incomplete orf downstream of ectC was found which showed a significant degree of derived polypeptide (50% amino acid sequence identity) similar to ectoine-associated aspartokinase (Ask) from Vibrio cholerae. To identify the 30 region of the putative ask gene, IPCR was performed with the self-ligated PvuI-fragment and primers AskR homologous to ect and AskF homologous to ask (Fig. 2.2A). For the second IPCR, 500 ng of chromosomal DNA was digested with MunI in a 50 ml reaction volume. DNA fragments were then ligated at 16 C overnight in a 100 ml reaction volume containing 200 ng of digested DNA and 2U of T4 DNA ligase in 1 ligase buffer as recommended by the manufacturer. Circular DNA molecules were used as a template for PCR with primers AskIR and AskIF homologous to the two ask segments. In the sequenced and assembled resultant products, another orf of 948 bp was found 360 bp downstream of ectABC-ask (Fig. 2.2A). Its closest homologue was a gene encoding ectoine hydroxylase (EctD) from Streptomyces avermitilis MA-4680 (GenBank BAC 74106) and S. chrysomallus (GenBank AAS02097) (40% and 42% identities), which is a member of the nonheme
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A PvuI
MunI Atf
ectR
Tra2.1 Tra2 Tra3
ectA AtrI
B
ectB Tra1
Tra4
AMOA
phyH
ask
EF
Tra4
Tra1
MunI
ectC CR
AclI
Eco RI Tra3
ectA C20
MunI
AskF
ectB
ectA
C
PvuI
AskIR AskIF
MunI AMOR AMOZ AMOF Tra 3
AMOFor
ATZ
CR
ectC
AclI
AclI
MunI
AskR
CR
ectB
ectC 7CR
7CF
Na-transporter
C20
Figure 2.2 Sequencing strategies used for identification of ectoine biosynthetic genes in Methylomicrobium alcaliphilum 20Z (A), Methylomicrobium kenyense AMO1 (B), and Methylobacter marinus 7C (C). The nucleotide sequences of oligonucleotide primers are given in Table 2.1.
iron(II)- and 2-oxoglutarate-dependent dioxygenase family (Reuter et al., 2010). Although no hydroxyectoine accumulation was detected by 1H NMR in Mm. alcaliphilum 20Z cells grown at salinities of 3% or 9% NaCl, it is still possible that the organism can make this compound under some growth conditions. From the assembled five-gene cluster, only four genes encoding DAB acetyltransferase (EctA, 18.8 kDa), DAB transaminase (EctB, 47.8 kDa), ectoine synthase (EctC, 15.2 kDa), and L-aspartokinase (Ask, 53.3 kDa) are transcribed as a single polycistronic mRNA as was shown by RT-PCR (Reshetnikov et al., 2006). The ectABC genes could be functional in Escherichia coli XL1-Blue since recombinant cells carrying them in the low-copy-number vector pHSG575 grew in the presence of 4% NaCl and synthesized ectoine (Reshetnikov et al., 2006). The analogous PCR strategy was used for deciphering the nucleotide sequences of ect-genes in another alkaliphilic methanotroph, Mm. kenyense AMO1 (Fig. 2.2B). Only three orfs corresponding to ectA, ectB, and ectC were found in a 3.5 kb genomic fragment of Mm. kenyense AMO1 with intergenic regions of 47 bp between ectA and ectB and 170 bp between ectB and ectC. No additional orf was found within 600 bp downstream of the ectC gene. PCR with degenerate primers Tra3/CR as the first step was also used to identify ect-genes in Mb. marinus 7C. On the basis of the sequence obtained ( 1000 bp), the homologous primers 7CF and 7CR were synthesized and
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used in “Vectorette” PCR performed as described by Ko et al. (2003). For this purpose, samples of genomic DNA were digested by EcoRI. After annealing and ligation of double strand “vectorette” unit (vect 57TTAA which complemented vect 53) to DNA digested by EcoRI, a “vectorette” PCR amplification with primer 7CF was performed (Table 2.1, Fig. 2.2C). For the second “vectorette” PCR, genomic DNA was digested by AclI. After annealing and ligation of “vectorette” unit (vect 57CG which also complemented vect 53), primer 7CR was used in the second round of PCR amplification. Generated fragments were sequenced and assembled. The 4.25 kb DNA fragment from M. marinus 7C included tightly linked ectABC genes without intergenic regions. An additional orf4 (1371 bp) was detected in the DNA locus 81 bp downstream of ectC. The product of this Orf4 (49 kDa) was similar to transport proteins belonging to Naþ/solute symporter family (SSF) from Mariprofundus ferrooxydans PV-1 ZP_01451115 (56% identity of amino acid sequences), Nitrosococcus oceani ATCC 19707 YP_343605 (46%), and Hyphomonas neptunium ATCC 15444 YP_760353 (43%). Two main types of ectoine gene organization were found in the methanotrophs which correlated with their salt tolerance and intracellular ectoine contents. Thus, Mb. marinus 7C and Mm. kenyense AMO1 possessing ectABC genes accumulated ectoine up to 70 mg per g of DCW and these methanotrophs were capable of growth at salinity 4–5% NaCl (Khmelenina et al., 2010). Conversely, Mm. alcaliphilum 20Z carrying the ectABC-ask operon was able to grow at higher salinity (up to 10% NaCl) and accumulate more ectoine (>120 mg per g of DCW). This implied an important role of a specific aspartate kinase in ectoine synthesis. Conversion of aspartate into b-aspartyl phosphate by aspartate kinase represents the starting point of the pathway for biosynthesis of both aspartate family amino acids and ectoine. Being osmotically controlled, aspartate kinase could make ectoine accumulation more independent from complex machinery regulating the divergent pathways leading from aspartate semialdehyde. Specialized ectoine-associated Ask enzymes can also be found in other halophilic Proteobacteria as judged from homology analysis of aspartate kinases (Lo et al., 2009). Remarkably, no Gram-positive bacteria contain an ectABC-ask gene cluster. The absence of the ask homologs in ectoine gene clusters of Firmicutes is explainable since carbon flow must also be directed to biosynthesis of the osmoprotectants belonging to the glutamate family amino acids, proline and/or glutamine, contributing to osmotic balance (Kuhlmann and Bremer, 2002; Saum and Muller, 2008). The second Ask encoded by the separate gene may occur in the Mm. alcaliphilum 20Z genome (https://www.genoscope.cns.fr/agc/mage) and it shares 17% identity with the osmotically controlled Ask. Notably, Ect proteins from the gammaproteobacterial alkaliphilic methanotrophs Mm. alcaliphilum 20Z and Mm. kenyense AMO1 are only
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distantly related (37–51% identities) to the enzymes from neutrophilic Mb. marinus 7C (also belonging to the Gammaproteobacteria) being most closely related to Ect proteins from zetaproteobacterial M. ferrooxydans (55– 80% identities). So far, Mb. marinus is the single species of the Methylobacter genus that is salt-resistant. It could be proposed that the ect operon ubiquity in ancient prokaryotic world that was largely marine was followed by loss in lineages that became adapted to a terrestrial environment (Lo et al., 2009).
3. Transcriptional Regulation of the Ectoine Biosynthesis Genes Routine genetic manipulations (DNA and RNA isolation from Mm. alcaliphilum 20Z, construction of mutant strains, Northern hybridization, and mapping of the transcriptional start sites) are described previously (Mustakhimov et al., 2010a) and elsewhere in this volume (Ojala et al., 2010). Analysis of the promoter region sequence showed that the ectABC-ask operon in Mm. alcaliphilum 20Z is initiated from two s70-like promoters ectAp1 and ectAp2. ectAp1 has a high level of identity with the consensus sequence of the E. coli s70-recognized promoter while the 10 and 35 sequences of ectAp2 differ from the respective regions of the E. coli s70promoter. Therefore, expression from ectAp2 could be less effective than from ectAp1. Northern hybridization also indicated that the ectC and ask genes might be cotranscribed as an additional transcriptional unit from the promoter region located upstream of the ectC gene. Transcription of ectR1 was carried out from a single s70-like promoter ectR1p located between ectAp1 and ectAp2. This implies that transcription from ectR1p may be controlled by its own product (Mustakhimov et al., 2010a). Upstream of ectA an additional gene transcribed in the reverse orientation was revealed (Mustakhimov et al., 2010a). This gene was named ectR1 (ectoine biosynthesis regulator) and it encoded a protein (20.6 kDa) homologous to the MarR-type transcriptional regulators (Wilkinson and Grove, 2006). A regulatory function of EctR1 was confirmed by studies of the ectR1 knockout mutant. Like most MarR-family regulators, EctR1 was shown to negatively control transcription of the ect operon by binding the promoter region (Mustakhimov et al., 2010a,b). It protects from DNAse I action an asymmetrical DNA locus that includes the 10 sequence of the ectA1p. The EctR1 binding site contains a pseudopalindromic sequence composed of 8-bp half-sites separated by 2 bp (TATTTAGT-GT-ACTATATA) suggesting dimeric association of the EctR1 with the DNA in which each subunit binds an inverted repeat. Gel-filtration data showed that EctR1 is
Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs
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a dimer both in free solution (44–45 kDa) and in the DNA-binding state (50–55 kDa). The ectR1-like genes were identified in the methanol- and methylamine-utilizing bacteria Methylophaga alcalica, Methylophaga thalassica, and Methylarcula marina (76%, 66%, and 54% identities of translated amino acid sequences) as well as in 22 genomes of heterotrophic halophiles (35.5–55.1% identities) (Mustakhimov et al., 2010b; Reshetnikov et al., 2010). All the EctR-like proteins showed relatively low identity (20%) with the characterized MarR-family transcriptional regulators. MarRfamily proteins include a diverse group of regulators controlling various physiological functions including response to environmental stresses, virulence factors, and aromatic catabolic pathways (Wilkinson and Grove, 2006). Although EctR1 and the MarR-family regulators have only low sequence identities, they likely have analogous protein structures due to the presence of helix-turn-helix DNA-binding motifs and flanking “wing 1” regions (Mustakhimov et al., 2010a).
4. Key Enzymes of Ectoine Biosynthesis As described above, the genes for ectoine biosynthesis in Mm. alcaliphilum 20Z are organized in one transcription unit ectABC-ask and have high similarity to the genes occurring in all groups of bacteria synthesizing ectoine. The encoded proteins have the following enzyme activities: EctB is diaminobutyric acid aminotransferase, EctA is diaminobutyric acid acetyltransferase, EctC is ectoine synthase, and Ask is aspartate kinase.
4.1. Diaminobutyric acid aminotransferase This enzyme was tested in a 1-ml assay volume containing 50 mM Tris– HCl buffer, pH 8.5, 10 mM 2-oxoglutarate, 20 mM diaminobutyric acid, 10 mM pyridoxal-50 -phosphate, 100 mM KCl and an enzyme preparation with activity in the range of 100 mU. The mixture was incubated at 25 C for 30 min. The reaction was stopped by boiling the mixture for 5 min and insoluble material was separated by centrifugation. In all, 20 ml samples were withdrawn and mixed with 5 ml of o-phthalaldehyde reagent (50 mg o-phthalaldehyde was dissolved in 1.25 ml of methanol, 50 ml of mercaptoethanol, and 11.2 ml 0.4 M borate buffer, pH 9.5). After 1 min exposure, 45 ml Na-acetate, pH 7.0, was added and immediately loaded onto a reversed-phase SEPARON SIX C-18 column (150 3.3 mm, 5 mm, Czech Republic). Degassed solutions of 0.1 M Na-acetate, pH 7.2, and methanol (0–40%) were used for gradient elution with a flow of 1 ml/min. Measurements were carried out on a fluorimeter “Gilson 121” (France) in
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9 ml cuvette with filters for excitation 305–395 and emission at 420–650 nm. Glutamic acid was used as inner internal standard. DABA aminotransferase from Mm. alcaliphilum 20Z was shown to be a 300 kDa hexameric enzyme with specific activity of 0.027 U/mg of protein and a calculated pI 5.49. DABA aminotransferase from Halomonas elongata purified by classical biochemical methods (Ono et al., 1999) was also a homohexameric (250 kDa) pyridoxal-50 -phosphate-dependent enzyme. It required Kþ for activity and stability and was more active in the presence of 0.01–0.5 M KCl than at the same concentrations of NaCl, specific to L-glutamate as aminodonor (Km for L-glutamate 9.1 mM) and to D,L-aspartyl semialdehyde (Km 4.5 mM). The reaction catalyzed by DABA aminotransferase is thought to be a limiting step of the ectoine biosynthetic pathway thus explaining the absence of DABA in H. elongata KS3 cells (Ono et al., 1999). Importantly, the second putative DABA aminotransferase gene located outside of ectABC-ask operon clustering with genes coding for ectoine degradation enzymes were found in the draft genome of Mm. alcaliphilum 20Z.
4.2. DABA acetyltransferase DABA acetyltransferase (EctA) acetylates diaminobutyric acid with acetyl coenzyme A forming g-N-acetyl-a,g-diaminobutyric acid. The enzyme activity was tested at pH 9.0 and 25 C in a 1-ml assay volume containing 50 mM Tris/HCl buffer, 0.1 mM 5,5-dithio-bis-(2-nitrobenzoic acid) (DTNB), 10 mM diaminobutyric acid, 1 mM acetyl-CoA, 200 mM KCl, and enzyme preparations with activity in the range of 10 mU. Formation of 2-thio-5-nitrobenzoic acid as a result of the interaction between 5,5-dithiobis-(nitrobenzoic acid) and sulfhydryl groups of coenzyme A released in the course of the reaction was monitored at l ¼ 412 (e412 ¼ 13.6 mM 1 cm 1) (Reshetnikov et al., 2006). DABA acetyltransferase from Mm. alcaliphilum 20Z was obtained by cloning of ectA into a pET22bþvector (Novagen, USA) and expression in E. coli BL21 (DE3). The His-tagged enzyme was purified on a Ni-NTAagarose column (Qiagen, Germany) in one stage (Reshetnikov et al., 2005). The kinetic properties of the Mm. alcaliphilum DABA acetyltransferase are presented in Table 2.2. Similar methods have been used for obtaining the purified recombinant EctA enzymes from two moderately halophilic methylotrophic bacteria M. thalassica and M. alcalica (Mustakhimov et al., 2008). Comparison of the enzyme properties revealed their correlation with ecophysiologies of host bacteria. Thus, the DABA acetyltransferases from the neutrophilic bacteria H. elongata (Ono et al., 1999) and M. thalassica were more active at lower pH (pH 8.2 or 9.0), than those from the alkaliphilic species Mm. alcaliphilum 20Z and M. alcalica (pH optima 9.5). DABA acetyltransferase from M. thalassica isolated from marine water was considerably inhibited by carbonates while the enzymes from the soda lake isolates
25
Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs
Table 2.2 Some properties of diaminobutyric acid acetyltransferases from methylotrophic bacteria
Properties
a
Mm. alcaliphilum M. alcalica M. thalassica 20Z (Reshetnikov (Mustakhimov (Mustakhimov et al., 2005) et al., 2008) et al., 2008)
pHopt Topt ( C) Molecular mass (SDS-PAGE) (kDa) Mr (gel-filtration) (kDa) Km (DABA) (mM) Km (acetyl-CoA) (mM) Inhibitors (1 mM)
9.5 20 20
9.5 30–35 20
8.5 30–35 20
40 0.465 36.7 Zn2þ, Cd2þ
Optimum KCl Optimum NaCl Stability
0.25 M 0.1–0.2 M Stablea
40 0.375 30 Zn2þ, Cd2þ, Cu2þ 0 0 Stablea
40 0.365 76 Zn2þ, Cd2þ, Cu2þ 0 0 Stablea
The enzyme was stable in 50 mM Tris–HCl, pH 8.5, at least for 1 month at 4–70 C at protein concentration 0.5 mg/ml.
M. alcalica and Mm. alcaliphilum 20Z were not affected, thus corresponding to in situ physiology of these species. Interestingly, Cu2þ at 1 mM completely inhibited the enzyme activity from M. alcalica and 47% of that of M. thalassica. In contrast, no inhibitory effect of Cu2þ was found for the Mm. alcaliphilum 20Z enzyme thus confirming an important role of copper in methane oxidation by the particulate methane monooxygenase of the methanotroph (Murrell et al., 2000). The differential dependence on ionic strength is another intriguing feature of the DABA acetyltransferases. The enzymes from H. elongata and Mm. alcaliphilum 20Z were activated by 0.4 or 0.2 M NaCl whereas the enzymes from M. thalassica and M. alcalica were inhibited by these salts. It might be speculated that the methanol-utilizing bacteria have more effective ion extrusion mechanisms in comparison to methane- or glucose-assimilating bacteria. We hypothesize that low monovalent inorganic ion concentrations are maintained in cytoplasm of the methanol utilizers. Different behavior of the enzyme at high or low ionic strength may be a result of its long-term adaptation in the host species.
4.3. Ectoine synthase The enzyme activity was tested at pH 9.0 and 25 C in a 1-ml assay volume containing 50 mM sodium-phosphate buffer, 20 mM N-a,g-diaminobutyric acid, and 300 mM NaCl and 10 mU of the recombinant EctC. The reaction
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was stopped by boiling the mixture for 5 min and insoluble material was separated by centrifugation for 20 min at 12,000g. The soluble fraction was dried under vacuum and ectoine content was measured by HPLC (Eshinimaev et al., 2007). A preparation of the recombinant ectoine synthase with specific activity of 64 U/mg of protein was obtained from Mm. alcaliphilum 20Z. The molecular mass of the purified ectoine synthase (35 kDa) suggests that the enzyme is a homodimer. The pI is 4.99. The native homogenous ectoine synthase purified from H. elongata with activity of 16 U/mg of protein was purified in the presence of 1 mM N-acetyl-DABA and 2 M NaCl as stabilizing compounds (Ono et al., 1999). The molecular mass of the native enzyme remained unclear because no activity was found after gel-filtration at 0.5 M NaCl. The enzyme was specific to Ng-acetyl-DABA and N-acetyl group in a-position could not be involved in cycling. The protein contained enhanced levels of aspartate and glutamate.
4.4. Aspartate kinase Halotolerant bacteria may use a specialized Ask paralog and the specialization can be achieved by differential regulation accomplished via allosteric control of enzyme activity and/or by transcriptional/translational control of enzyme synthesis (Lo et al., 2009). However, the ectoine-associated Ask enzyme of halophilic/halotolerant bacteria has never been studied. We made several attempts to obtain the recombinant enzyme by cloning of the gene ask either into vector pET30 (Novagen) designed to express a C-terminal His-tagged fusion protein or pET28 (Novagen) designed to express a N-terminal His-tagged fusion protein. However, E. coli BL21 (DE3) transformed with the pET30/ask or pET28/ask plasmids synthesized a non-soluble protein. Co-expression of aspartate kinase with chaperones (GroES/EL) was also accompanied by insoluble protein synthesis. Aspartate kinase in the soluble form (53.3 kDa) was obtained by expression in Methylobacterium extorquens AM1. Gene ask was amplified from pET28/ask or pET30/ask and DNA fragments were cloned into vector pCM160 carrying constitutive promoter of methanol dehydrogenase (Pmax) from M. extorquens AM1 (Marx and Lidstrom, 2001). The resulting plasmid pCMaskN-his (or pCMaskC-his) was transferred into M. extorquens AM1 by conjugation and cells were grown on mineral medium supplemented by 0.5% (v/v) methanol. A single step of immobilized metal ion affinity chromatography on a Ni2þ-NTA column was performed for purification of His6-tagged Ask fusion protein from M. extorquens extract (Mustakhimov et al., 2008). The preparation of the recombinant enzyme with activity 25 mU/mg of protein was obtained. A protein band corresponding to
Genes and Enzymes of Ectoine Biosynthesis in Methanotrophs
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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2011 Elsevier Inc. All rights reserved.
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facultative methanotrophy in new isolates requires a great degree of caution since methanotrophic cultures are frequently contaminated by heterotrophic bacteria that survive on metabolic by-products of methanotrophs. The presence of only a few satellite cells in a culture may lead to false conclusions regarding substrate utilization, and several early reports of facultative methanotrophy are likely attributable to impure cultures. Another recurring mistake is the misidentification of nonmethanotrophic facultative methylotrophs as facultative methanotrophs. This chapter was prepared as an aid to avoid both kinds of confusion when examining methanotrophic isolates.
1. Introduction At present, aerobic methanotrophic capabilities are recognized in members of two bacterial phyla, the Proteobacteria (the classes Alpha- and Gammaproteobacteria), and the Verrucomicrobia (reviewed in Op den Camp et al., 2009). Bacteria from the as yet unnamed candidate phylum NC10 also perform aerobic methanotrophy after forming free dioxygen from NO, but as yet little is known about these (Ettwig et al., 2010). Most aerobic methanotrophs grow only on methane, plus in some instances on methanol, formate, formaldehyde, and methylamines, and are therefore termed “obligate methanotrophs.” By contrast, facultative methanotrophs are able to use either methane or some multicarbon compound(s) as their sole carbon and energy source. The occurrence of facultative methanotrophy was recently demonstrated in several alphaproteobacterial methanotrophs of the genera Methylocella, Methylocystis, and Methylocapsa (Belova et al., 2011; Dedysh et al., 2005; Dunfield et al., 2010; Im et al., 2011). Members of these genera differ with regard to their physiology and substrate preferences. Methylocella species were the first to be conclusively shown to have a facultative capability. Among the known methanotrophs, these bacteria are very unique. They possess only a soluble methane monooxygenase (sMMO) and lack an extensive intracytoplasmic membrane (ICM) system common to all other methanotrophs. Perhaps not surprisingly given their unique status among methanotrophs, they can also utilize a number of multicarbon compounds (acetate, pyruvate, succinate, malate, and ethanol). These substrates are in fact preferred, and sMMO in Methylocella is repressed if an alternative multicarbon growth substrate is present (Theisen et al., 2005). Later, several facultatively methanotrophic Methylocystis spp. and Methylocapsa aurea were described. Unlike the oddball Methylocella, these are more typical aerobic methanotrophs, possessing a particulate methane monooxygenase enzyme (pMMO) and a well-developed ICM system in which pMMO is bound. The preferred growth substrate of these organisms is methane, but growth
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can also occur on acetate and/or ethanol in the absence of methane. Notably, it has been shown for a Methylocystis that pMMO is expressed in the presence of these alternate substrates, which can be utilized alone or together with methane. Thus, a facultative lifestyle may occur in both pMMO- and sMMOpossessing methanotrophs. It should be noted, however, that the existence of methanotrophs capable of growth on multicarbon compounds was a controversial topic for a long time prior to 2005. Periodic reports of facultative methanotrophs could not be independently verified (reviewed in Dedysh and Dunfield, 2010; Theisen and Murrell, 2005). The most common problems with these studies were: (1) lack of evidence for the genetic and enzymatic machinery for methane oxidation in a target organism and (2) questionable culture purity. In other words, either a nonmethanotrophic facultative methylotroph was erroneously identified as a facultative methanotroph or the study was conducted using a tight syntrophic association between a methanotroph and a facultative methylotroph. In this chapter, we describe our standard procedures for identifying methanotrophic capabilities in a novel isolate, for testing its ability to grow on multicarbon substrates, and for evaluating methanotroph culture purity.
2. Identification of Methanotrophic Capabilities in Novel Isolates 2.1. Culture conditions Both obligate and facultative methanotrophs are cultivated using liquid or solid mineral media with methane as a growth substrate. The use of multicarbon compounds for isolation or laboratory maintenance of facultative methanotrophs is not recommended since these substrates give a selective advantage to heterotrophic bacteria, which overgrow and contaminate methanotrophic cultures. Media that can be used for methanotroph cultivation include: A. Nitrate mineral salts (NMS) medium (Bowman, 2000; Whittenbury et al., 1970) is most widely used for methanotroph cultivation. It contains (g/l distilled water): KNO3, 1; 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 (<1 in 100). A pyrosequencing procedure could be used to obtain a larger number of sequences. Alternatively, to rule out this possibility, a simple dilution growth experiment can be made as follows: 1. Serial (10-fold) dilutions for most-probable-number (MPN) counts are prepared using quadruplicate 20-ml test tubes containing 5 ml of medium. The source culture for the MPN should be grown through multiple transfers on acetate (or other desired substrate) alone. One set of MPN tubes contains acetate for the enumeration of acetate-utilizing bacteria, while another set of tubes is incubated in closed glass desiccators under a headspace containing 10% (v/v) CH4 for the enumeration of methane-utilizing bacteria. 2. Tubes are incubated for 1 month and MPNs are calculated using an Excel-based program (Briones and Reichardt, 1999). If pure, the examined culture should contain equal numbers of methaneutilizing and acetate-utilizing cells, even after multiple transfers of this culture on acetate alone. The same experiment can be made with a source culture grown through multiple transfers on methane.
4.6. Quantification of methane monooxygenase-coding genes during growth on an alternate substrate The increase of methanotroph-specific mmoX or pmoA genes during growth on an alternate substrate such as acetate clearly demonstrates that methanotrophic cells and their genomes are multiplying on a substrate other than methane. 1. Substrate utilization tests should be performed as in Section 3. Periodically samples should be taken for measurement of optical density or direct cell counts. 2. Genomic DNA is extracted from each sample using a mechanical cell disruption procedure. 3. qPCR is performed using a Sybr-Green based method (A TaqMan approach is also feasible). A series of pmoA and mmoX primers targeting
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various groups of methanotrophs have been designed for use in qPCR assays by Kolb et al. (2003). 4. qPCR should be performed to target the 16S rRNA gene and the respective pmoA or mmoX gene of the isolate (Kolb et al., 2003). Variations in extraction efficiency and copy numbers may mean that the resulting gene counts are not equal, however, the growth curves measured via OD or direct cell counting should closely parallel the increase in copy numbers of the 16S rRNA genes and methanotrophspecific functional genes.
REFERENCES Auman, A. J., Stolyar, S., Costello, A. M., and Lidstrom, M. E. (2000). Molecular characterization of methanotrophic isolates from freshwater lake sediment. Appl. Environ. Microbiol. 66, 5259–5266. Belova, S. E., Baani, M., Suzina, N. E., Bodelier, P. L. E., Liesack, W., Dunfield, P. F., and Dedysh, S. N. (2011). Acetate utilization as a survival strategy of peat-inhabiting Methylocystis spp. Environ. Microbiol. Reports 3, 36–46. Bourne, D. G., McDonald, I. R., and Murrell, J. C. (2001). Comparison of pmoA PCR primer sets as tools for investigating methanotroph diversity in three Danish soils. Appl. Environ. Microbiol. 67, 3802–3809. Bowman, J. (2000). The methanotrophs—The families Methylococcaceae and Methylocystaceae. release 3.1. In “The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community,” (M. Dworkin, et al., eds.), 3rd edn. http://link.springer-ny.com/ link/service/books/10125/. Briones, J. A. M., and Reichardt, W. (1999). Estimating microbial population counts by ‘most probable number’ using Microsoft Excel. J. Microbiol. Meth. 35, 157–161. Costello, A., and Lidstrom, M. E. (1999). Molecular characterization of functional and phylogenetic genes from natural populations of methanotrophs in lake sediments. Appl. Environ. Microbiol. 65, 5066–5074. Dedysh, S. N., and Dunfield, P. F. (2010). Facultative methane oxidizers. In “Handbook of Hydrocarbon and Lipid Microbiology,” (K. N. Timmis, ed.), pp. 1967–1976. SpringerVerlag, Berlin. Dedysh, S. N., Liesack, W., Khmelenina, V. N., Suzina, N. E., Trotsenko, Y. A., Semrau, J. D., Abing, A. M., Panikov, N. S., and Tiedje, J. M. (2000). Methylocella palustris gen. nov., sp. nov., a new methane-oxidizing acidophilic bacterium from peat bogs, representing a novel subtype of serine-pathway methanotrophs. Int. J. Syst. Evol. Microbiol. 50, 955–969. Dedysh, S. N., Knief, C., and Dunfield, P. F. (2005). Methylocella species are facultatively methanotrophic. J. Bacteriol. 187, 4665–4670. Dunfield, P. F., Khmelenina, V. N., Suzina, N. E., Trotsenko, Y. A., and Dedysh, S. N. (2003). Methylocella silvestris sp. nov. a novel methanotrophic bacterium isolated from an acidic forest cambisol. Int. J. Syst. Evol. Microbiol. 53, 1231–1239. Dunfield, P. F., Belova, S. E., Vorob’ev, A. V., Cornish, S. L., and Dedysh, S. N. (2010). Methylocapsa aurea sp. nov., a facultatively methanotrophic bacterium possessing a particulate methane monooxygenase. Int. J. Syst. Evol. Microbiol. 60, 2659–2664.
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Ettwig, K. F., Butler, M. K., Le Paslier, D., Pelletier, E., Mangenot, S., Kuypers, M. M. M., Schreiber, F., Dutilh, B. E., Zedelius, J., de Beer, D., Gloerich, J., Wessels, H. J. C. T., et al. (2010). Nitrite-driven anaerobic methane oxidation by oxygenic bacteria. Nature 464, 543–549. Holmes, A. J., Costello, A., Lidstrom, M. E., and Murrell, J. C. (1995). Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132, 203–208. Im, J., Lee, S.-W., Yoon, S., Dispirito, A. A., and Semrau, J. D. (2011). Characterization of a novel facultative Methylocystis species capable of growth on methane, acetate and ethanol. Environ. Microbiol. Reports 10.1111/j.1758-2229.2010.00204.x (in press). Kolb, S., Knief, C., Stubner, S., and Conrad, R. (2003). Quantitative detection of methanotrophs in soil by novel pmoA-targeted real-time PCR Assays. Appl. Environ. Microbiol. 69, 2423–2429. McDonald, I. R., Kenna, E. M., and Murrell, J. C. (1995). Detection of methanotrophic bacteria in environmental samples with the PCR. Appl. Environ. Microbiol. 61, 116–121. McDonald, I. R., Bodrossy, L., Chen, Y., and Murrell, J. C. (2008). Molecular ecology techniques for the study of aerobic methanotrophs. Appl. Environ. Microbiol. 74, 1305–1315. Miquez, C. B., Bourque, D., Sealy, J. A., Greer, C. W., and Groleau, D. (1997). Detection and isolation of methanotrophic bacteria possessing soluble methane monooxygenase (sMMO) genes using the polymerase chain reaction (PCR). Microbial Ecol. 33, 21–31. Op den Camp, H. J. M., Islam, T., Stott, M. B., Harhangi, H. R., Hynes, A., Schouten, S., Jetten, M. S. M., Birkeland, N. K., Pol, A., and Dunfield, P. F. (2009). Minireview: Environmental, genomic, and taxonomic perspectives on methanotrophic Verrucomicrobia. Environ. Microbiol. Reports 1, 293–306. Shigematsu, T., Hanada, S., Eguchi, M., Kamagata, Y., Kanagawa, T., and Kurane, R. (1999). Soluble methane monooxygenase gene clusters fron trichloroethylene-degrading Methylomonas sp. strains and detection of methanotrophs during in situ bioremediation. Appl. Environ. Microbiol. 65, 5198–5206. Stahl, D. A., and Amann, R. (1991). Development and application of nucleic acid probes. In “Nucleic Acid Techniques in Bacterial Systematics,” (E. Stackebrandt and M. Goodfellow, eds.), pp. 205–248. Wiley, New York, NY. Theisen, A. R., and Murrell, J. C. (2005). Facultative methanotrophs revisited. J. Bacteriol. 187, 4303–4305. Theisen, A. R., Ali, M. H., Radajewski, S., Dumont, M. G., Dunfield, P. F., McDonald, I. R., Dedysh, S. N., Miguez, C. B., and Murrell, J. C. (2005). Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol. Microbiol. 58, 682–692. Weisburg, W. G., Barns, S. M., Pelletier, D. A., and Lane, D. J. (1991). 16S ribosomal DNA amplification for phylogenetic study. J. Bacteriol. 173, 697–703. Whittenbury, R., Phillips, K. C., and Wilkinson, J. F. (1970). Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61, 205–218.
C H A P T E R
F O U R
Approaches for the Characterization and Description of Novel Methanotrophic Bacteria John P. Bowman Contents 1. Introduction to Aerobic Methanotroph Characterization for Systematic Research and Other Purposes 2. Broad Level Systematic Diversity 2.1. Environmental relevance and knowledge limitations 2.2. Requirements related to the Bacteriological Code 3. Methodological Approaches 3.1. Enrichment and isolation 3.2. Minimal standards for characterization 3.3. Chemotaxonomy 3.4. Informative sequence-based analyses and genomic comparisons 3.5. Culture collections 4. Concluding Remarks References
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Abstract Methanotrophs are a key group of environmental microorganisms that play an integral role in the global cycling of methane. Modern culture-independent techniques, traditional painstaking culture methods, and genomics-related approaches continue to reveal new information about methanotrophs, including their ecosystem associations, biochemistry, and their systematic and evolutionary nature. The increasing interest in methanotrophic bacteria especially in the context of climate change will likely lead to an increase in cultures available for in-depth studies. This chapter details the rigorous approach needed for successful appraisal and confirmation of novel aerobic methanotrophic bacterial species that can then be made accessible for in vitro research purposes. This includes a guide in which descriptions maintain adherence to the Bacteriological Code; that Tasmanian Institute of Agricultural Research, University of Tasmania, Hobart, Tasmania, Australia Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00004-8
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2011 Elsevier Inc. All rights reserved.
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meet minimal standards; and the mandatory deposition of representative strains in approved culture collections.
1. Introduction to Aerobic Methanotroph Characterization for Systematic Research and Other Purposes Aerobic methanotrophs represents a metabolically allied collection of bacteria that possess the ability to grow on methane as a sole carbon and energy source. To this current point in time, methanotrophy is carried out by specialist bacteria, which have largely dispensed with multicarbon compound catabolism. All known methanotrophs can utilize the metabolic products generated from methane oxidation (methanol, formaldehyde) and sometimes other C1 compounds. Claims of facultative methanotrophy, in which strains have been shown to be able to utilize methane as well as multicarbon compounds, has been controversial (Dedysh et al., 2004) but far from being unprecedented (Dunfield et al., 2010; Stoecker et al., 2006). Methanotrophy that occurs anaerobically is carried out by consortia of specific methanogenic archaea and sulfate-reducing bacteria (Knittel and Boetius, 2009). Since this process is ecologically and biochemically distinct from aerobic methanotrophy it will not be dealt with in this chapter. Furthermore, the syntrophic nature of anaerobic methanotrophy hinders the systematic understanding of the associated microorganisms performing the process and at this stage much research is still being focused on better understanding the biochemical and ecological nature of the process itself.
2. Broad Level Systematic Diversity The aerobic methanotrophic process on the other hand is much better understood and is based on decades of research involving pure cultures, including model strains, such as Methylococcus capsulatus BATH (Ward et al., 2004), coupled to environmental in situ studies. Aerobic methanotrophy is possible due to possession of methane monooxygenase (MMO; EC 1.14.13.25) an oxidoreductase enzyme that comes in two major varieties (Baik et al., 2003). Most cultured bacteria that possess MMO belong to the phylum Proteobacteria and are thus Gram-negative. Most are motile by flagella. No cultured strains have been shown, conclusively at least, to be capable of autotrophic growth using CO2 for carbon or able to use an alternative energy sources such as H2 or CO; however, systematic studies to date tend to follow a well-trodden path in terms of test methods (see Section 3). Recently, extremely acidophilic and mildly thermophilic
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methanotrophs were identified that belong to the phylum Verrucomicrobia (Dunfield et al., 2007; Islam et al., 2008; Pol et al., 2007). As of late 2010, none of these verrucomicrobial isolates were validly described though they have been provisionally proposed as belonging to a genus designated “Methylacidiphilium” (Op den Camp et al., 2009). Methanotrophs belonging to the phylum Proteobacteria are grouped into families that are generally monophyletic (clustered together) or represent closely related paraphyletic (distinct but adjacent) lineages. These distinct groupings suggest methanotrophy arose multiple times following the major divergence of bacteria from archaea (Zerkle et al., 2005) and possibly coevolved with ammonia oxidizers owing to the similarity in ammonia monooxygenase and the copper-containing particulate MMO (pMMO). The members of each of the proteobacterial families possess distinct phenotypic differences and thus the methanotrophs are often termed in generalist subgroupings as type I, type II (and occasionally in the case of Methylococcus and its immediate relatives, “type X”) methanotrophs. Type I methanotrophs include those belonging to the class Gammaproteobacteria and include psychrophilic to mildly thermophilic species and are grouped together in the family “Methylococcaceae” represented by genus Methylomonas and its relatives. Methanotrophs with a preference for mildly thermal environments form a cluster deeper in class Gammaproteobacteria and include genus Methylococcus and Methylocaldum and represent effectively family Methylococcaceae sensu stricto, since Methylococcus is the type genus. The description of a multitude of new species has resulted in the originally described family Methylococcaceae (Bowman et al., 1995) becoming increasingly less monophyletic; however, it is now well recognized that cultured methanotroph diversity is only a fraction of what can be observed using PCR-based and array-based culture-independent techniques (Kip et al., 2010). Type II methanotrophs occur in the closely adjacent families Methylocystaceae and Beijerinckaceae, which belong to the class Alphaproteobacteria. The family Beijerinckiaceae incorporate non-methanotrophic genera (i.e., Beijerinckia, Rhodoblastus); however, recent evidence suggests most of these probably are methylotrophic (Dedysh et al., 2005; Kulichevskaya et al., 2006). The type II methanotrophs are also closely related to the genus Methylobacterium commonly isolated pink-pigmented, methylotrophs able to utilize C1 and multicarbon compounds. It is thus recognized the trait of methylotrophy, especially methanol utilization, is highly relevant to methanotrophs. All methanotrophic genera have traditionally been named with the genus prefix “Methylo-”; however, an exception is Crenothrix polyspora. This microbe is very much a distinction since it suggests that methanotrophs have evolved not only surprising complexity in terms of life cycle and growth strategies but also unsuspected metabolic flexibility, including
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utilization of alternative energy sources. As perhaps the best example of a microorganism that does not follow the standard “methanotroph mold,” C. polyspora is a sheathed, filamentous, iron-accumulating bacterium that has been shown to have a rather distinctive pMMO; forms unusual hexagonal and fibrillar-like inclusion bodies of unknown function; a life cycle and morphology not unlike various cyanobacteria, that is, formation of gonidia (Vo¨lker et al., 1977); and based on radio-labeling studies is able to take up multicarbon substrates (Stoecker et al., 2006). Further (systematic) studies may yet reveal that methanotrophs are diverse, sophisticated, and highly networked microorganisms within the Earth’s biosphere.
2.1. Environmental relevance and knowledge limitations Ecologically aerobic methanotrophs exist at positions in which oxygen and methane is available. Methane is mainly derived from methanogenesis but also occurs due to abiotic processes (i.e., volcanic, extraterrestrial). The densest populations of aerobic methanotrophs overly anoxic zones in which methane is emitted and enters aerobic zones. Such locations include mainly freshwater aquatic environments and soil (Chen and Murrell, 2010). In marine zones, aerobic methanotrophy may occur if the environment is sulfate depleted or where active natural gas seeps or accumulated methane clathrates occur, in general, anaerobic methane oxidation may be otherwise more important (Treude and Ziebis, 2010). Little information is available on marine aerobic methanotrophs and is thus fertile ground for study. In soil ecosystems, it is also noted that indigenous methanotrophs may have high affinity MMO and thus are able to utilize atmospheric methane (currently at about 1.8 ppm). No examples of these “high affinity” methanotrophs are available in pure culture; indeed, it is uncertain whether they exist at all and rather cultured methanotrophs behave differently under conditions of low methane relying on other forms of exogenous carbon (Degelmann et al., 2010). Indeed, the general practice is to always grow methanotrophs under highly methane excess conditions; thus understanding of the full metabolic potential of methanotrophs is currently poorly developed.
2.2. Requirements related to the Bacteriological Code Microbial taxonomy and the nomenclatural framework it rests upon in recent years have increasingly become better defined. As a primer, it is strongly suggested the reader consult the paper “Notes on the characterization of prokaryote strains for taxonomic purposes” (Tindall et al., 2010), which spells out in detail the process and information content expected for the quality expected in novel species descriptions. An additional critical criterion that must be emphasized is strain availability. For a species type strain to be validly described it must be housed in at least two internationally
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recognized culture collections (Ka¨mpfer, 2010). Culture collection recognition is conducted by the Word Federation of Culture Collections ( www. wfcc.info); however, the extant nature of the collection is not verified and thus methanotroph deposition should be performed using only collections that have a reliable history and the associated expertise (see Section 3.5 for more information). Other strains can be described for research purposes; however, only by fulfilling the strict criteria adopted by the International Committee on Systematic Bacteriology (www.the-icsp.org) and the Bacteriological Code (Lapage et al., 1992; freely available at http://www.ncbi.nlm.nih. gov/bookshelf/br.fcgi?book¼icnb) can they be recognized as taxonomically official. A full list of official names is maintained at the LPSN (List of Prokaryotic names with Standing in Nomenclature) website at www.bacterio. cict.fr. The LPSN web resource is an excellent information portal for taxonomic names and their status, type strain designations, culture collections where type strains are found; and verified 16S rRNA sequence data derived from type strains (Pruesse et al., 2007; www.arb-silva.de). The actual process through which the appropriate and exacting isolation and characterization of methanotrophs occurs by is very important, owing to special problems that are a feature of this particular type of bacteria. For successful taxonomic delineation of methanotroph isolation a set of minimal requirements is clearly required.
3. Methodological Approaches The processes by which the taxonomy and thus the essential biological and biochemical features of methanotrophs are obtained are described below in a series of distinct stages: (3.1) enrichment and isolation; (3.2) essential identification and baseline characterization; (3.3) chemotaxonomy; (3.4) sequence analysis; and (3.5) long-term preservation.
3.1. Enrichment and isolation Enrichment is usually required to isolate methanotrophs though direct plating of diluted samples onto agar can also be used where they occur in abundance. The latter applies typically to freshwater sediment and some soils. Primary selection medium is traditionally nitrate mineral salts (NMS) formulated originally by Roger Whittenbury and colleagues (Bowman, 2006; Whittenbury et al., 1970). Ammonia salts are not used as they may competitively inhibit MMO. Methane must be of high purity since as even traces of acetylene will permanently inactivate MMO. The NMS medium can be modified to suit the environments being sampled. For isolation of methanotrophs from saline systems the NMS salt content should be adjusted
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to match the salinity profile of the source samples (Bowman, 2006). Methanotrophs from freshwater and soil samples are usually rather salt-sensitive. For acidic soils the medium has been modified by diluting the medium 1:5 and pH adjusted to approximately pH 4.5–6.0 using a phosphate buffer (Dunfield et al., 2007). Typically for acidic soils soil particles are placed directly on agar plates (Dunfield et al., 2003). Obviously empirical media modification could reveal novel methanotrophs and fundamentally the initial conditions should match the environment being sampled. For enrichments bottles or tubes that are crimped sealed with butyl rubber stoppers are ideal. Agar plates can be incubated in desiccators or anaerobic jars. The atmospheres of the vessels are adjusted by removal of 5–50% of the headspace and replacing with methane. For routine cultivation, the standard level of methane most often used is a 1:4 CH4:air ratio. Since all known pure cultures have a relatively low affinity MMO (be it pMMO or soluble MMO) the actual concentration of the methane atmosphere is not critical except perhaps from a safety point of view. Incubation temperatures should reflect the source environment and can potentially range from 4 C for tundra and Antarctic samples to ca. 60 C for thermal springs and aquifers. The given optimal growth temperature of isolates can be tested empirically. An ideal means to do biokinetic temperature data accurately is described by Ratkowsky et al. (1983). Most polar species described to date have temperature growth ranges typical of “psychrotrophs” (4 to 30–37 C) or psychrophiles (0–22 C) (Wartiainen et al., 2006). The latter are often notoriously slow growing (doubling times 1 day) (Bowman et al., 1997). Most known methanotrophs are mesophiles growing from 10 to 40 C (optimum ca. 30 C) while the species of genus Methylococcus and Methylocaldum grow from 20 to 55 C (optimum ca. 45 C). Species evolved for acidic or alkaline environments do occur (Kalyuzhnaya et al., 2008). Type I methanotrophs include several examples of halophilic strains including salt and soda lakes, coastal and estuarine species (e.g., Methylomicrobium pelagica, Methylohalobius crimeensis; Heyer et al., 2005; Kalyuzhnaya et al., 2008). The general lack of descriptions from more saline water bodies suggests large untapped methanotrophic diversity in many different aquatic ecosystems that could be readily accessible through standard cultivation approaches. The analysis of geochemical signatures could be a very useful first step in exploring sites for the presence of active aerobic methanotrophy. For example the presence of signature fatty acids (see Section 3.3) and aminobacteriohopanepolyols (Zhu et al., 2010) may be useful as a guide for sample selection for subsequent isolation and characterization studies. Agar media can be prepared with normal agar. Agarose or noble agar as a solidifying agent may be ideal during the purification phases though not a requirement. It should be noted some methanotrophs do not grow or do not maintain stable growth on agar-solidified media possibly due to intolerance to the agar (Bowman et al., 1997; Rahalkar et al., 2007). In these cases, liquid
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culture is required or silica gel-solidified media (Bazylinski and Rosenberg, 1980); however, the latter is potentially technically difficult to prepare and has been used infrequently for methanotrophs. Purification of agarophobic methanotrophs requires repeated dilution to extinction in liquid media (Bowman et al., 1997) and so far has rarely been done due to technical difficulties and some element of luck in obtaining pure cultures. The utilization of highthroughput culturing methods (Connon and Giovannoni, 2002; Toledo et al., 2006) has yet to be exploited significantly for methanotroph cultivation and could be ideal approaches for isolation of novel methanotrophs. Colonies that subsequently appear on primary isolation agar plates are without exception contaminated with other non-methanotrophic bacteria. The main culprits are non-methanotrophic methylotrophs and oligotrophic, heterotrophic bacteria such as Brevundimonas, Caulobacter, etc. To obtain pure cultures the most effective method is to add a small amount of readily useable carbon to the medium (typically 0.1% glucose or 0.05% yeast extract) and then to dilute cell suspensions to extinction. The extra carbon stimulates the growth of the contaminants to form distinct colonies otherwise they grow by cross-feeding on metabolites from the methanotrophs and thus cannot be separated from primary growth. Plates need to be incubated for extended periods of time and thus care must be made against fungal contamination. Addition of cycloheximide, nystatin, or natamycin is advisable. Containers holding plates need to be cleaned regularly with ethanol and disinfectant to avoid fungal contamination. Final checks of purity require microscopic examination of cells and colonies for morphological consistency. This process may require repetition before pure cultures can be obtained, if at all. Validation of purity must be performed by plating cultures onto rich media such as R2A agar or 0.1 strength trypticase soy agar (both available from Oxoid and Difco Laboratories). Very rich media such as nutrient agar or full-strength trypticase soy agar are not ideal since oligotrophic contaminants may not grow on these media. The final confirmation of purity requires extra rigor for claims of facultative methanotrophy to be proven owing to past issues with culture purity. This can be achieved by denaturing gradient gel electrophoresis of 16S rRNA PCR products in which case only one PCR product should be observable; and sequence analysis of 16S rRNA, which should reveal a chromatogram with only a low background signal. Confirmation of purity of verrucomicrobial methanotrophs required direct genome sequencing (Pol et al., 2007). In some cases, methanotroph growth is stimulated by the presence of exogenous multicarbon compounds (such as small amounts of tryptone or yeast extract); however, there is a perpetual threat of culture contamination thus it is ideal for cultures to be maintained under methane only conditions. Finally, vitamin requirements are not unprecedented for methanotrophs and should be considered in any enrichment and isolation process though they may potentially complicate purification of strains.
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3.2. Minimal standards for characterization Standard tests for methanotrophs are summarized in Table 4.1 and are linked to special features possessed by certain methanotrophs and thus have known discriminating power. A species description of high quality would involve thorough investigation of all properties listed in Table 4.1 and would ideally involve comparison of multiple strains against suitable reference cultures; however, the requirement to have multiple strains is not obligatory. Popular commercial test strips such as API (Biome´rieux-Vitek) and Biolog are generally not utilized in testing methanotrophs. However, certain salient properties such as enzymes, and responses to nitrogen and phosphate sources as well as response to exogenous carbon sources have not to this point in time been given much attention. Opportunities thus exist to utilize relatively new testing platforms such as the Phenotype MicroarrayTM system from Biolog (http://www.biolog.com/pmMicrobialCells.html) to test for novel properties of both new and old methanotrophic strains. These would advance the taxonomy as well as general physiological understanding of methanotrophs. Fundamentally, the essential basics for validation of methanotrophs, especially if a claim for facultative use of methane is argued includes (a) determination of cellular morphology and importantly intracytoplasmic membrane ultrastructure; (b) determination of the predominant pathway for formaldehyde fixation; (c) means of methane oxidation (i.e., pMMO and/or sMMO); (d) basic cellular and colonial nutritional and physiological properties; (e) chemotaxonomy (see Section 3.3); and (f) 16S rRNA, pmoA, and nifH gene sequence analysis. Many descriptions to date are based on single isolates thus the inherent variation in traits at the species level can generally only be assessed at the genus level.
3.3. Chemotaxonomy Guanosine–cytosine (GþC) ratios of DNA and fatty acid profile are mandatory for novel descriptions of methanotrophs as these data are considered a useful means of differentiation not only of the broad types of methanotrophs but also of individual genera. GþC ratio analysis is now popularly analyzed by HPLC analysis and is considered a rapid, routine analysis procedure (Mesbah et al., 1989). The presence of distinct fatty acids by aerobic methanotrophs are taxonomically useful in distinguishing genera and/or species and has also been adopted as signatures of their presence in environmental samples (Bodelier et al., 2009; Bowman et al., 1991) though limitations to this technique are now clear due to some methanotrophs apparently lacking any designated signature lipids (e.g., Methylovulum miyakonense; Iguchi et al., 2010). Also it has been found that some type II methanotrophs may possess signature fatty acids typically found in type I methanotrophs (Dedysh et al., 2007). Most studies utilize the Sherlock Microbial Identification System
Table 4.1 A list of characteristics designated as minimal standards for characterization of novel aerobic methanotrophs Characteristicsa
Morphology and ultrastructure Cell shape, cell division, capsules Mode of motility Resting cells—type and relation to survival of desiccating conditions and heating Cell wall stability based on lysis in 2% sodium dodecyl sulfate solution Intracytoplasmic membrane (ICM) presence and morphology Spinae (extensions of the S-layer) Intracellular granules—polyhydroxyalkanoate (PHA) and polyphosphate Pigmentation Physiology and biochemistry pH range; NaCl tolerance; temperature range for growth Naphthalene oxidation assay for sMMO in the presence and absence of copper Formaldehyde fixation pathway
Special features that occur in some methanotrophs
Rosettes and budding cell division (Methylosinus spp., some Methylocella spp.); sarcinal packets (Methylosarcina spp.) Polar tufts (Methylosinus spp.) or polar flagella otherwise Cysts (most type I methanotrophs; Methylocystsis spp.); exospores (Methylosinus spp.) Type II methanotrophs are more stable and resist lysis in 2% SDS Absent in Methylocella spp. have a distinct ICM arrangement compared to other methanotrophs Formed by some Methylocystis spp. (can be up to 150 nm) Large PHA granules may occur depending on the strain (possibly can be confused with cysts) some strains may have distinct pigments though it is uncommon Methylocella and Methylocapsa are acidophilic; various type I methanotrophs show preferences for different pH, salinity, and/or temperature Methylocella spp. only possess sMMO
(continued)
Table 4.1 (continued) Characteristicsa
Special features that occur in some methanotrophs
Ribulose monophosphate pathway (type I methanotrophs); Serine pathway (type II methanotrophs) Catalase, cytochrome c oxidase Completeness of the TCA cycle; presence of glyoxylate cycle, glycolysis, and pentose phosphate pathway key enzymes
2-Oxoglutarate dehydrogenase may be absent in some type II methanotrophs
Other enzymatic activity—phosphatases, nitrate reduction, urease Carbon/nitrogen source tests Many strains can use a range of amino acids as nitrogen sources Nitrogen fixation – resting cell acetylene reduction assay; nifH gene sequencing Sequence analysis pmoA, 16S rRNA sequencing a
May vary between strains of the same species
Specific methods for characteristics can be found in the recent methanotroph taxonomic literature: Dedysh et al. (2007), Dunfield et al. (2003, 2010), Heyer et al. (2005), Kalayuzhnaya et al. (2008), and Rahalkar et al. (2007). More general guidelines for taxonomic descriptions and the standard and quality needed for such data is described in Tindall et al. (2010).
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(MIDI Inc., http://www.midi-inc.com) and several laboratories offering paid MIDI analysis services (e.g., DSMZ—German Collection of Microorganisms; http://www.dsmz.de/identification/main.php). Since the transesterified fatty acids are identified by GC in comparison to retention time libraries, the more unusual fatty acids typical of several methanotrophs (such as 16:1o8c, 18:1o8c) may not be accurately identified. To confirm their presence requires specific double-bond directed derivatization (Nichols et al., 1986) and subsequent GC/MS analysis. Certain fatty acid components are also produced at highly variable levels (e.g., affected by O2 availability), especially cyclopropane fatty acids ( Jahnke and Nichols, 1986) and thus there is a need for standardization in culture conditions. Quinone analysis may also have value owing to some methanotrophs possessing quinones that have methylenated isoprenoid chains (Watzinger et al., 2008). Analysis of quinone compounds requires HPLC and MS for confirmation (Tindall et al., 2010). Polar lipid analysis is becoming more popular in bacterial taxonomic characterization and has potential value for methanotrophs as some species produce distinctive patterns of polar lipids (Andreev and Galchenko, 1983). It is required that this type of analysis be performed using two-dimensional thin layer chromatography with appropriate use of indicator spray reagents. Tindall et al. (2010) describes appropriate standards needed for description of fatty acid, quinone, and polar lipid data. Methanotrophs also contain unusual sterol and hopanoid compounds (Cvejic et al., 2000; Elvert and Niemann, 2008). No systematic study has been completed to determine if these lipids vary between species or groups and whether they have taxonomic value. Such information could be valuable as these more unusual lipids could also be useful as environmental biomarkers.
3.4. Informative sequence-based analyses and genomic comparisons DNA:DNA hybridization data is an obligatory element for novel taxa descriptions unless they are distinct in terms of 16S rRNA sequence similarity. The generally accepted threshold is 97%; however, a description must also be supported by other evidence including phenotypic, chemotaxonomic, and other genetic data (Tindall et al., 2010). DNA:DNA hybridization can be performed by a variety of means including the S1 nuclease technique (Christensen et al., 2000), renaturation kinetics (De Ley et al., 1970) or by using photobiotin labeled DNA (Hirayama et al., 1996). The DSMZ offers a paid service to perform DNA:DNA hybridization analyses. It has become the practice to also compare novel strains on the basis of similarity of the pmoA gene though the comparisons may not correlate with 16S rRNA gene sequence data for closely related taxa. The analysis also further validates the nature of methanotrophy in a given strain. Though sMMO genes can be also identified by PCR normally the naphthalene oxidation assay (Brusseau et al., 1990) is performed for cells grown in the
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presence and absence of copper ions. Oxidation leading to the formation of naphthol in the absence of copper and abolishment of this activity in the presence of copper is usually sufficient evidence for sMMO activity. It should be noted some type II methanotrophs, primarily Methylocella spp. have been described that lack pMMO and only form sMMO. Evidence for nitrogen fixation can also be established by PCR detection of nitrogenase nifH genes; however, a resting acetylene reduction assay must be also used to confirm nitrogenase activity and supported by evidence of growth in a medium lacking a combined nitrogen source (Bowman, 2006).
3.5. Culture collections Though methanotrophs have been shown to be stably maintained under a methane atmosphere for several months (Bowman, 2006), many methanotrophic species (especially type I methanotrophs) tend to survive long-term preservation (lyophilized, deep freezing in glycerol suspensions, etc.) sometimes poorly or not at all. Green and Woodford (1992) suggest methanol and polyvinylpyrrolidone could be useful cryoprotectants; however, these chemicals need to be tested with species isolated in recent years. Thus, for long-term maintenance of type strains or other important strains for perpetuity and also to make them publically accessible the strains need to be housed in culture collections that have already extensive methanotroph collections, for example, NCIMB, DSM, and VKM (Table 4.2). As listed in Table 4.2, the available type strains are housed in a disparate range of collections. It would be ideal thus that methanotrophs are distributed as much as possible thus to reduce the chance of being lost, for example, a situation made possible by culture collection extinction brought about potentially by lack of funding. All species that are in the “lost” category are type I methanotrophs thus extra attention is obviously required for novel taxa described in this group due to their apparent sensitivity to freezing.
4. Concluding Remarks The role of methane as a greenhouse gas and current suspected trends of global climate change (e.g., Shakhova et al., 2010) has resulted in much scientific interest and research investigating the significance aerobic methanotrophy plays as a biological sink for methane. One aspect of this research is the discovery and characterization of novel methanotrophic bacteria. The current trend suggests that we still have an undeveloped concept of the diversity of methanotrophs (Kip et al., 2010; Op den Camp et al., 2009; Stoecker et al., 2006) and as mentioned above their inherent physiological capacities are still unplumbed. Thus, cultivation studies coupled to the
Table 4.2
Aerobic methanotroph pure culture location and environmental locations of isolation
Genus name
Extant species and type strain location
Type I methanotrophs Methylomonas M. methanica ATCC, NCIMB, VKM Methylobacter M. luteus ATCC, NCIMB; M. psychrophilus VKM, M. tundripaludum ATCC, DSM, M. whittenburyi ATCC, NCIMB Methylomicrobium M. agile ATCC NCIMB; M. albus ATCC, NCIMB, VKM; M. pelagicus NCIMB; M. alcaliphilum NCIMB, VKM; M. japanense NBRC, FERM-P, VKM; M. kenyense NCCB, NCIMB, VKM Methylosoma M. difficile DSM, JCM Methylovulum M. miyakonense NBRC, DSM Methylosarcina M. fibrata ATCC, DSM; M. lacus ATCC, JCM; M. quisquiliarum ATCC, DSM Methylohalobius M. crimeensis ATCC, DSM Methylothermus M. thermalis VKM Methylococcus group Methylococcus M. capsulatus ATCC; M. thermophilus NCIMB Methylocaldum
M. gracile NCIMB, VKM; M. szegediense NCIMB
Known environmental associations
Terrestrial (soil, water), groundwater Temperate and polar terrestrial (soil, water) Terrestrial (soil, water), freshwater and soda lakes, estuarine, seawater
Freshwater lakes Forest soil
Hypersaline lake (Crimea, Russia) Hot spring (Japan) Terrestrial (soil, water); warm spring, ground water Warm springs, groundwater (continued)
Table 4.2
(continued)
Genus name
Extant species and type strain location
Type II methanotrophs Methylosinus M. sporium ATCC, NCIMB; M. trichosporium ATCC, NCIMB Methylocystis M. echinoides IMET, VKM; M. heyeri DSM, VKM; M. hirsuta ATCC, DSM; M. parvus ATCC, DSM, NCIMB, VKM; M. rosea ATCC, DSM Methylocapsa M. acidiphila DSM, NCIMB Methylocella M. palustris ATCC; M. silvestris DSM, NCIMB; M. tundra DSM, NCIMB
Known environmental associations
Terrestrial (soil, water), groundwater Terrestrial (soil, water), groundwater, tundra soil Acidic peat/sphagnum moss soils Acidic peat/sphagnum moss soils, tundra soils
ATCC, American Type Culture Collection, Manassas, VA, USA (www.atcc.org); DSM, German Culture Collection, Braunschweig, Germany (www.dsmz.de); NCIMB, National Collection of Industrial and Marine Bacteria, Aberdeen, Scotland, UK (www.ncimb.com); VKM, All-Russian Collection of Microorganisms, Russian Academy of Sciences, Institute of Biochemistry and Physiology of Microorganisms, Pushchino, Moscow Region, Russia (www.vkm.ru); JCM, Japan Collection of Microorganisms, Institute of Physical and Chemical Research (RIKEN), Wako, Saitama, Japan (www.jcm.riken.go.jp); NCCB, Netherlands Culture Collection of Bacteria (formerly Phabagen and LMD), Utrecht University, Utrecht, the Netherlands (www.cbs.knaw.nl/nccb); FERM-P, Fermentation Research Institute, Agency of Industrial Science and Technology, Chiba, Japan; NBRC, NITE Biological Resource Center, Department of Biotechnology, National Institute of Technology and Evaluation, Chiba, Japan (www.nbrc.nite.go.jp). These species no longer appear to be available due to loss of the cultures or due to extinction of the culture collections in which they were housed: Methylobacter marinus; Methylococcus chroococcus; Methylococcus mobilis; Methylomonas aurantiaca; Methylomonas fodinarum; Methylomonas scandinavica; Methylocaldum tepidum; Methylosphaera hansonii. The species Crenothrix polyspora is not available as a pure culture (www.bacterio.cict.fr).
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increasing knowledge capture possible with ‘omics-based technologies, will likely result in methanotroph systematic biological knowledge having periodic and potentially exciting updates. This is especially possible given the very recent appearance of high-throughput proteomics technologies (Yates et al., 2009) that moves us beyond the inert nature of genome sequence data to the dynamic mapping of physiological information in whole-genome contexts. To galvanize this and maximize the utility of novel strains to scientific research, an excellent platform requires to be maintained for the systematic of methanotrophs and stable and reliable repositories for strains.
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C H A P T E R
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Methylococcus capsulatus (Bath): From Genome to Protein Function, and Vice Versa Odd A. Karlsen, Frode S. Berven,1 June I. Bagstevold, Øivind Larsen,2 and Harald B. Jensen Contents 1. Introduction 2. Mapping of the Outer Membrane Proteome of M. capsulatus 2.1. Prediction of OMPs from the predicted M. capsulatus proteome 2.2. Experimental identification of (predicted) OMPs. Subfractionation and 2DE analysis of the M. capsulatus proteome 2.3. Identifying unannotated proteins in the M. capsulatus genome 3. Inner Membrane Protein Complexes Studied with Blue-Native Polyacrylamide Gel Electrophoresis (BN-PAGE) 3.1. Detergent extraction of the M. capsulatus inner membrane protein complexes 3.2. BN-PAGE of Triton X-100 solubilized membranes 3.3. Large-scale 2D BN-/SDS-PAGE 4. Visualization and Identification of M. capsulatus c-type Heme Proteins 4.1. Heme staining of M. capsulatus c-type heme proteins Acknowledgments References
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Abstract The genome sequence of Methylococcus capsulatus (Bath), considered a model methylotroph, was published in 2004 [Ward, N., et al. (2004). Genomic insights into methanotrophy: the complete genome sequence of Methylococcus Department of Molecular Biology, University of Bergen, Bergen, Norway Present address: Proteomic Unit (PROBE), Department of Biomedicine, University of Bergen, Bergen, Norway 2 Present address: Uni Environment, Bergen, Norway 1
Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00005-X
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2011 Elsevier Inc. All rights reserved.
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capsulatus (Bath). PLoS Biol. 2, e303]. In the postgenomic era, the challenge is to determine the gene function, and to this end, genomics must be complemented with proteomic approaches. This chapter describes some experimental and computational approaches we have used and developed for the exploration of the genome and proteome of M. capsulatus (Bath).
1. Introduction As the first complete genome sequence from an obligate methanotroph, the genome sequence of Methylococcus capsulatus (Bath) was published in 2004 (Ward et al., 2004). Analysis of the 3.3-Mb genome allowed the annotation of 3113 open-reading frames (ORFs), with about 1766 proteins similar to proteins of known function and role category, 514 conserved hypothetical proteins, and 504 hypothetical proteins. The genome sequence confirmed previous studies on carbon assimilation (Strom et al., 1974; Taylor et al., 1981), nitrogen fixation (Murrell and Dalton, 1983), hydrogenase (Hanczar et al., 2002), and the presence of particular and soluble methane monooxygenase (Stainthorpe et al., 1989, 1990; Stolyar et al., 1999). More surprisingly was the presence of two ORFs with high sequence similarity to 2-oxyglutarate dehydrogenase, one of the key enzyme activities of the tricarboxylic acid cycle not found in type I methanotrophs (Davey et al., 1972). The genome is continuously updated (NCBI GenBank Accession number AE017282). The genome sequencing uncovered redundancy in many pathways: some 200 linage-specific gene duplications were found covering both methane oxidation, carbon assimilation, amino acid biosynthesis, energy metabolism, transport, regulation, and environmental sensing (Ward et al., 2004). Thus, in addition to the previously known redundancy for methane monooxygenase (Stanley et al., 1983), all of the other steps involved in methane oxidation to CO2 are covered by redundant enzyme systems (Ward et al., 2004). The high content of duplicated ORFs, the relatively large number of c-type cytochromes (Karlsen et al., 2008), and membranemodifying components, including sterols (Bird et al., 1971), hopanoids ( Jahnke et al., 1992), and trans fatty acids (Loffler et al., 2010), are consistent with an organism able to rapidly adapt to varying growth conditions. It has become clear that the large number of ORFs predicted from the increasing number of sequenced genomes alone has limited value in relation to a complete understanding of gene function and cellular physiology. The genome sequence is the blueprint of the organism and its theoretical (metabolic) capabilities. When entering the postgenomic era, the challenge is to determine the gene function, and genomics must therefore be complemented with proteomic approaches. Thus, it is necessary to examine the
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expression levels of the genes and protein responses to biological and environmental stimuli, posttranslational modifications (PTMs), and the interactions of these molecules in the cell. Computational algorithms that have been developed for identifying functional genes in bacterial genomes have recently been reviewed (Gao and Chen, 2010; Poptsova and Gogarten, 2010). The use of computational analysis for exploring the genome has proved very useful for the prediction of genes and the functions, location, and structure of their protein products. Such analysis includes sequence alignment, chromosomal location, structural prediction (Gao and Chen, 2010), and comparative genome analysis (Poptsova and Gogarten, 2010). These analyses can be the fundament for either generating hypotheses that must be experimentally verified or for supporting the conclusions and findings of the experimental work. In the following sections, we describe the experimental and computational approaches we have used and developed for the exploration of the M. capsulatus genome and proteome. Computational tools, including some developed in-house, for the prediction of outer membrane proteins (OMPs) were combined with subproteome fractionation and proteomics methods for the identification of transmembrane b-barrel proteins and lipoproteins. We show how proteomics, in combination with an ORF library, can be used in order to identify c-type cytochromes, also those located to the M. capsulatus cellular surface. We also demonstrate the use of Blue-Native PAGE to reveal protein complexes in the inner membrane of M. capsulatus (Bath), which can aid in decoding the functional proteome.
2. Mapping of the Outer Membrane Proteome of M. capsulatus Integral b-barrel proteins constitute a major protein type in the OM of Gram-negative bacteria. These types of proteins have diverse functions such as passive nutrient intake, active ion transport, membrane anchors, and enzymes (Schulz, 2000). Another group of OMPs is the lipoproteins that are attached to the OM through lipid anchors. Peripheral proteins may be associated to the outer and inner leaflet of the OM, or they can be associated with OMPs through protein–protein interactions, without themselves being in direct contact with the OM. In the following sections, we focus on the prediction of integral b-barrel proteins and lipoproteins from the genome sequence, and experimental identification of such proteins. Most of the tools and approaches we present here are generic and are applicable to most Gram-negative bacteria. The strategies for subproteome fractionation may, however, vary from one bacterial species to another.
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2.1. Prediction of OMPs from the predicted M. capsulatus proteome 2.1.1. Prediction of b-barrel proteins A large number of software tools that predict integral b-barrel OMPs have been developed, and an overview of most of these tools is given elsewhere (Remmert et al., 2009). Some of them can only process one sequence at the time, while others can process thousands of sequences simultaneously. The latter category includes the b-barrel Outer Membrane protein Predictor (BOMP) and HHomp (Berven et al., 2004; Remmert et al., 2009) and is preferred for the prediction of the entire b-barrel subproteome. In addition to b-barrel prediction, the positive predictions are analyzed for the presence of signal sequences, and this is an indication of transport of the protein to the membranes or the periplasm, or secretion. BOMP is based on two separate components to recognize integral b-barrel proteins: one component calculates an integral b-barrel score of the sequence based on the extent to which the sequence contains stretches of amino acids typical of transmembrane b-strands, while the other component searches for a C-terminal pattern typical of many integral b-barrel proteins. The following steps describe the use of BOMP to predict b-barrel OMPs in M. capsulatus, and additional experimental analysis that may support the predictions. 1. Go to the BOMP webpage: http://services.cbu.uib.no/tools/bomp and upload a file with the protein sequences in FASTA format. 2. If BLASTp information is desired for the dataset, check this option and set the E-value (typically, 1e10). This will increase the run time to some extent and will also provide additional information to the predictions, and therefore, we recommend that this option is used. 3. The predictions will be given as categories 1–5 with increasing probability of being a b-barrel protein with higher category; Table 2 in Berven et al. (2004) gives information about the meaning of the categories. 4. Generally, sequences ending up in categories 4 and 5 are considered to be b-barrel OMPs with very high probability. When submitting all the predicted protein encoding sequences from 10 genomes of Gram-negative bacteria, none of the sequences predicted in categories 4 and 5 was confirmed to have localization other than the OM. 5. The output from BOMP is a list of predicted b-barrel OMPs and the category to which they belong. In addition, a FASTA file of these sequences is created, and this list can be submitted to other prediction programs to further increase the reliability of the predictions. 6. We have typically used SignalP (http://www.cbs.dtu.dk/services/SignalP/) and secondary structure prediction programs like PSIPRED (http://bioinf. cs.ucl.ac.uk/psipred/) to obtain more information about the predicted b-barrel OMPs, but other b-barrel OMPs predictors can also be used, as for example, HHomp (Berven et al., 2004; Remmert et al., 2009).
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2.1.2. Prediction of lipoproteins Lipoproteins belong to another important category of proteins of the OM. A couple of software tools have been developed to predict such proteins from larger genome data, including Lipo (http://services.cbu.uib.no/tools/ lipo) developed by us (Berven et al., 2006), and LipoP (http://www.cbs.dtu. dk/services/LipoP/; Juncker et al., 2003). Lipo recognizes lipoproteins based on a lipo-box pattern that was extracted from 183 Gram-negative lipoproteins collected from Swiss-Prot. The predicted lipoproteins were divided into three categories: high, medium, and low, based on how frequent each amino acid appears in each position in the lipo-box in the 183 training set proteins. In the following sections, we will describe our approach for the prediction of lipoproteins in M. capsulatus using Lipo. 1. Go to the Lipo webpage: http://services.cbu.uib.no/tools/lipo and upload a file with the protein sequences in FASTA format. 2. The predictions will be given as categories high, medium, low, indicating the likelihood of a correct prediction. 3. The output from Lipo is a list of predicted lipoproteins and the category to which they belong, and whether the lipoprotein is likely to be located in the IM or OM. In addition, a FASTA file of these sequences is created, and this list can be submitted to other prediction programs to further increase the reliability of the predictions. 4. The predictions in the high-scoring category had a precision of 92%, medium 73.7%, whereas the precision of the low-scoring category was only 13.3%. The number of lipoproteins found in the low category was only 5%, so if high confidence is more important than the possibility of losing some lipoproteins in the prediction, the low-category predictions can be excluded. 5. We have typically used SignalP (http://www.cbs.dtu.dk/services/SignalP/) to obtain more information about the predicted lipoproteins, but other lipoprotein predictors can also be used if increased confidence is desired, as for example, LipoP ( Juncker et al., 2003), DOLOP (http://www.mrc-lmb. cam.ac.uk/genomes/dolop/; Babu et al., 2006; Madan Babu and Sankaran, 2002), and PROSITE (http://au.expasy.org/prosite/).
2.2. Experimental identification of (predicted) OMPs. Subfractionation and 2DE analysis of the M. capsulatus proteome 2.2.1. Subfractionation M. capsulatus cells are fractionated according to the protocols outlined in a separate chapter in this issue (chapter eleven). The resulting protein fractions are enriched in soluble proteins, inner membrane proteins, OMPs, membrane-associated proteins, and surface-associated proteins, respectively.
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2.2.2. 2DE analysis The different protein fractions residing from the fractionation procedure were analyzed by 2DE. By this approach, it is relatively easy to get an overview of the presence and relative amounts of each protein in the different fractions. Most proteins are rarely present exclusively in one fraction, but when looking at the spot abundances from the 2DE gels, it is possible to determine in which fraction the protein is most abundant. This comparison reveals information about the localization of the proteins in the cell and gives indications on which type of OMP that has been identified, for example, surface associated, integral b-barrel protein, membrane associated, etc. The 2DE separation of M. capsulatus OMPs is described in detail in Berven et al. (2003). Protein spots enriched in the OM fractions were analyzed by mass spectrometry and/or N-terminal sequencing revealing their identity, as described elsewhere (Berven et al., 2003, 2006). A representative combined 2DE gel of the enriched OMP fraction is shown in Fig. 5.1. The numbered protein spots have been identified and are given in Table 5 in Berven et al. (2006). 2.2.3. Multiple spots When analyzing the OMPs of M. capsulatus (and in some instances other protein fractions), we observed that many spots in the gel corresponded to the same protein species. The multiple spot patterns could be horizontal as well as vertical. It is not always easy to distinguish between PTMs and pI
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Figure 5.1 2DE gel separation of M. capsulatus OMPs. A 2DE gel showing the OMP fraction of M. capsulatus separated using six overlapping narrow range IPG strips with overlapping pH intervals, followed by separation of each strip using 12.5% SDS-PAGE. The six resulting gel images were combined to give one large 2DE gel. All strips were loaded with equal protein amounts. Spots were detected by silver staining. Spot numbers refer to Table 5 by Berven et al. (2006). The figure is reproduced from Fig. 1 in the same article with kind permission from Springer ScienceþBusiness Media.
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conformational equilibria of proteins (Berven et al., 2003). The origin of horizontal spot patterns could be caused by PTMs giving charge alterations, as well as conformational equilibria arising in vitro during the 2DE procedure. The 2DE-based protocol outlined in Section 2.2.3.1 can be followed in order to reveal if the latter is the case. Figure 5.2 shows the result of using this procedure to analyze a train of polypeptide spots from the OMP fraction, spots 1–3 in Fig. 5.1, all representing the MopG protein, see Table 5 in Berven et al. (2006). It is clear from the figure that the entire original spot pattern can be regenerated from (any) one of the excised spots. Due to the great stability of b-barrel proteins (Schulz, 2000), oligomers, folded monomers, and denatured monomers can be found in varying amounts in 2DE, due to incomplete denaturation (Berven et al., 2003; Exner et al., 1995; Fjellbirkeland et al., 1997). An example can be seen in Fig. 5.1, spots 11–17, which all represent the OMP MopB (see Table 5 in Berven et al., 2006). See the 1DE procedure outlined in Section 2.2.3.2 in order to investigate such a case. 2.2.3.1. Analyzing (in vitro-generated) horizontal multiple spot patterns in 2DE gels
1. Analyze the protein sample using 2DE (Berven et al., 2003), and load as much protein as possible. 2. Excise one spot from the horizontal train of spots observed in the Coomassie Brilliant Blue R-250 stained 2-DE gels. 3. Elute the excised polypeptide e.g. by using the model 422 Electro-Eluter from Bio-Rad, utilizing a volatile electrode buffer, according to the manufacturer’s protocol. pH 4.3 Original spot pattern Re-run spot 1 Re-run spot 2
Train 3 1
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Figure 5.2 2DE reapplication of 2DE separated polypeptides. Polypeptide spots of MopG, labeled as train 3 in the figure, were excised from Coomassie Brilliant Blue-stained 2-DE gels and resubjected to a new 2DE separation. The polypeptide pattern observed after the first 2DE separation is shown in the upper panel. The spot numbers indicate the polypeptide spot that was resubjected to 2DE. Narrow-range IPG-strips (18 cm) pH 4.0–5.0 were used in the first-dimensional separation and SDS-polyacrylamide gels (12.5%) in the second-dimension electrophoresis. The gels were stained with silver nitrate. The figure is reproduced from Fig. 1 in Berven et al. (2003) with kind permission from Wiley-VCH verlag Gmbh & Co. KGaA.
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4. Precipitate the eluted polypeptide using 80% (v/v) acetone, allow at least 1 h on ice, resuspend in standard IEF buffer and resubject the polypeptide to 2DE separation, but without the use of an alkylating agent. 5. After the second 2DE separation, the gel is typically silver-stained in order to account for the inevitable loss of material during sample preparation and the second 2DE run, and since one original spot can split up to many isoforms. 6. If a similar spot pattern as the original one is generated from one single excised spot, then it is likely that the original spot pattern was not generated by in vivo PTMs, but rather generated during the 2DE analysis procedure. 2.2.3.2. Analyzing structural isoforms of OMPs resulting in spots with identical pI but different apparent MW in 2DE gels Analyze the protein sample on 2DE, excise and elute relevant polypeptide spots as above (Section 2.2.3.1, 1–4), but resuspend in SDS sample buffer and heat each polypeptide for at least 5 min at 95 C before 1D SDS-PAGE. Most proteins, also the very stable outer membrane b-barrel proteins, are now found as denatured monomers.
2.3. Identifying unannotated proteins in the M. capsulatus genome 2.3.1. Generation and use of an ORF library A database is generated from the total genome sequence using GLIMMER (http://cbcb.umd.edu/software/glimmer/). All the theoretically possible ORFs with a length of more than 100 bases are generated, resulting in a database of 31,490 sequences. Thus, we are sure that all putative ORFs, also the unannotated ones, are included in the database. A refined database based on proteins containing the c-type heme motif, CxxCH, can then be constructed from the ORF library. All mass spectra obtained from excised polypeptide spots/bands, where no identification was obtained, were searched against the ORF library. This gave us the possibility to identify potentially important proteins that would not have been discovered if the original annotated database had been used (Berven et al., 2006; Karlsen et al., 2008).
3. Inner Membrane Protein Complexes Studied with Blue-Native Polyacrylamide Gel Electrophoresis (BN-PAGE) As the number of sequenced genomes increases rapidly, the challenge is to assign a function to each protein and elucidate its interactions with other proteins and macromolecules in the cell. BN-PAGE was initially
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developed to study protein–protein interactions of membrane proteins, which due to their hydrophobic character, have been difficult to study with other methods (Schagger and von Jagow, 1991). BN-PAGE has later also been applied for studying soluble protein complexes (CamachoCarvajal et al., 2004), and has proved to be an efficient method for a highresolution separation of multiprotein complexes, maintaining their native conformation and providing information of their relative abundance and exact composition. In the following sections, we describe (unpublished results) how to empirically find the best-suited detergent concentrations for efficient extraction of inner membrane protein complexes, and we present the combined BN/SDS-PAGE procedure to resolve protein complexes present in inner-membranes of M. capsulatus. Proteins of interest can be identified with MS, and these findings can be compared to genomic structure to provide information about gene clusters and operons, and composition of the protein complexes and possibly, on their biological role.
3.1. Detergent extraction of the M. capsulatus inner membrane protein complexes To study membrane proteins and their complexes by BN-PAGE, it is crucial to extract them from their biological surroundings. Mild detergents are used in order to substitute the hydrophobic environment of the lipid bilayer and transfer the membrane proteins into the aquatic phase. It is important to notice that protein complexes differ substantially in stability. Detergent solubilization must therefore be tested empirically with different detergents at different concentrations and incubation times to ensure that protein–protein interactions are maintained (Reisinger and Eichacker, 2006). Triton X-100 has proved to efficiently solubilize the inner membrane of M. capsulatus, as described by (Fjellbirkeland et al., 1997) and elsewhere in this issue. Total membranes of M. capsulatus are incubated with different concentrations of Triton X-100 and subsequently analyzed by BN-PAGE to find a suitable concentration of detergent for efficient extraction of the inner membrane protein complexes. The same general procedure can be applied to membranes isolated from other bacterial species, and the type of detergent can be changed (e.g., BOG, DDM, Digitonin; Reisinger and Eichacker, 2006). 1. A total membrane fraction from M. capsulatus is obtained, as described in chapter eleven in this issue. 2. Samples containing 100 mg of total membranes are centrifuged at 100,000g for 1 h at 4 C. The pellets are resuspended in 17.5 ml BNPAGE sample buffer (750 mM e-amino caproic acid, 50 mM Bis–Tris, pH 7.0, and 0.5 mM EDTA–Na2) and incubated with Triton X-100 in concentrations ranging from 0.25% to 2.5% (w/v). 3. The samples are solubilized for 30 min at 4 C and centrifuged as above for separation of the insoluble material from the solubilized protein complexes.
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4. The supernatants are gently mixed with 1 ml BN-PAGE loading buffer (750 mM e-amino caproic acid and 5% Coomassie G-250) and BN-PAGE is performed, as described in Section 3.2. Coomassie G-250 renders the proteins with negative charges prior to the electrophoresis.
3.2. BN-PAGE of Triton X-100 solubilized membranes 1. A NativePAGETM gel cassette (4–16%, Invitrogen) is prepared as described by the manufacturer. The BN-PAGE gel can also be cast in the lab, as described elsewhere (Reisinger and Eichacker, 2006). 2. Cathode buffer (50 mM Tricine, 15 mM Bis–Tris, pH 7.0, 0.02% Coomassie G-250) is added to the wells prior to protein loading of the Triton X-100 extracted samples. A molecular mass marker used for native gels is applied to one of the lanes (e.g., NativeMark Unstained Protein Standard (Invitrogen)). 3. Cathode buffer and anode buffer (50 mM Bis–Tris, pH 7.0) are added to the inner and outer buffer chamber, respectively. Both buffers are prechilled to 4 C. 4. The electrophoresis is carried out at 4 C with a constant voltage of 150 V for 1 h, which subsequently is increased to 250 V until the blue dye-front reaches the bottom of the gel. The blue cathode buffer is replaced with a colorless cathode buffer (lacking Coomassie) when the Dye-front has migrated approximately one-third of the gel. Proteins are visualized by staining with Coomassie R-250 (Fig. 5.3).
3.3. Large-scale 2D BN-/SDS-PAGE BN-PAGE is often combined with SDS-PAGE in a second dimension, a variant of the common two-dimensional isoelectric focusing/SDS-PAGE. In order to identify the subunits of the protein complexes that were maintained during the BN-PAGE, a lane from the BN-gel is treated with a denaturing solution containing SDS and reducing agents, and placed on top of the second-dimension stacking gel. As a result, subunits comprising the same complex will dissociate and be separated based on their individual molecular mass in the second-dimension electrophoresis. Due to comigration in the first dimension, subunits will appear as spots along a straight vertical line (Reisinger and Eichacker, 2006). 3.3.1. The first-dimension BN-PAGE 1. An M. capsulatus total membrane faction, corresponding to 400 mg of proteins, is centrifuged for 100,000g for 1 h at 4 C.
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% Triton X-100 kDa 0.25 0.5 1.0 1.5 2.0 1048
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Figure 5.3 Empirical testing of Triton X-100 extraction of inner membrane protein complexes. Approximately 100 mg of proteins from the enriched total membrane fraction were solubilized using Triton X-100 concentrations ranging from 0.25% to 2.5% (w/v), and subsequently analyzed with BN-PAGE. An increased efficiency of protein extraction and electrophoretic mobility shift toward protein complexes of higher molecular weight are observed when a 1% Triton X-100 concentration is used for protein extraction. This is especially evident for the protein band migrating with a molecular mass between 242 and 480 kDa, as indicated in the figure, suggesting that a Triton X-100 concentration of approximately 1% is suitable for the extraction of the inner membrane proteins. The Native Mark Unstained Protein Standard (Invitrogen) is shown to the left. The gel is stained with Coomassie R-250.
2. Resuspend the pellet in 70 ml BN sample buffer (as described above) and add Triton X-100 to a final concentration of 1.25%. 3. Solubilize the sample for 30 min at 4 C. 4. The sample is centrifuged at 100,000g for 1 h at 4 C to remove the membranes from the soluble protein complexes. The supernatant is gently mixed with 5 ml of BN loading buffer (as described above). The BN-PAGE electrophoresis is performed as described (Section 3.2) in a large polyacrylamide gel electrophoresis system (typically, a gel size of 18–16 cm) with a BN gradient gel of desired concentration. The power program is divided into two phases: Phase I 1000 V, 6 mA, 24 W, 45 min; Phase II: 1000 V, 12 mA, 24 W, 2 h. The electrophoresis is carried out at 4 C. 3.3.2. The second-dimension SDS-PAGE 1. The lane from the BN-PAGE electrophoresis containing the separated protein complexes is excised from the BN polyacrylamide gel.
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2. The gel is transferred to a small compartment and incubated for 20 min with 2% SDS and 2% b-mercaptoethanol while rotating. The equilibrated lane is placed horizontally on top of the second-dimension polyacrylamide gel (uniform polyacrylamide concentration of 12.5% with a 4% stacking gel). 3. A molecular mass marker (denatured) is added onto a small piece of Whatman 3MM paper and placed beside the first-dimension lane. 0.5% agarose in Tris-glycine electrophoresis buffer is pored over the lane and the molecular mass marker to seal their position. Care is taken to avoid trapping air between the lane and the stacking gel. The electrophoresis is carried out using the following power program: Phase I: 1000 V, 30 mA, 30 W, 30 min; Phase II: 1000 V, 8 mA, 30 W, 14 h. The electrophoresis unit is cooled to 15 C during the electrophoresis. The resulting gel can be stained with e.g. Colloidal Coomassie or electrotransferred to a membrane for protein immunoblot analyses (Fig. 5.4). Subunits of a protein complex are observed as spots aligned in a vertical line. These spots can be excised from the polyacrylamide gel and identified by mass spectrometry. Reverse genetics toward the M. capsulatus genome provides information about genomic organization (operon), other putative interaction partners, and possibly biological function (e.g., see Fig. 5.4).
4. Visualization and Identification of M. capsulatus c-type Heme Proteins The genome sequencing of M. capsulatus revealed an unexpected large complement of c-type cytochromes; 57 proteins containing a c-type hemebinding motif (CxxCH) were identified during the genome annotation (Ward et al., 2004). This is in clear contrast to Escherichia coli where only seven c-type cytochrome-encoding genes are present in the genome (Methe et al., 2003). The large number of proteins with c-type heme-binding motifs makes M. capsulatus resemble some facultative or strictly anaerobic bacteria that contain numerous c-type cytochromes, such as the dissimilatory metal reducing bacteria Shewanella oneidensis MR1 and Geobacter sulfurreducens, with 42 and 111 predicted c-type cytochromes, respectively (Heidelberg et al., 2004; Methe et al., 2003). Recently, several of the multi c-type heme proteins of M. capsulatus were found located on the cellular surface (Karlsen et al., 2008). This is not commonly observed in bacteria, but a feature of dissimilatory metal-reducing bacteria that utilize an extracellular electron acceptor (Myers and Myers, 2003). These findings imply that redox reactions involving c-type cytochromes can also take place at this cellular localization in methanotrophic bacteria. In the following sections, we outline a protocol for visualizing the
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Figure 5.4 Two-dimensional BN/SDS-PAGE of M. capsulatus inner membrane proteins. (A) Approximately 400 mg of enriched total membrane fraction was solubilized with 1.25% Triton X-100, and separated in a 5–15% BN gradient gel, prior to the second-dimension electrophoresis, using a uniform 12.5% SDS-polyacrylamide gel. Subunits comprising heteromeric protein complexes during the first-dimension BNPAGE appeared along vertical lines in the 2D-gel. Spots 1–3 were identified by MS as the ATP synthase F1a-, b-, and g-subunit, respectively, originating from a protein complex migrating with a molecular mass corresponding to 500–650 kDa. The other polypeptide spots in the same vertical line has not yet been identified, but they correspond well in molecular masses to the ATP synthase subunits F1d (19 kDa), F1e (15 kDa), F0a (29 kDa), F0b (17 kDa), and F0c (8 kDa), as predicted from their encoding genes in the M. capsulatus (Bath) genome. Thus, the ATP synthase complex migrates in the BN-PAGE with an apparent molecular weight that is in line with the expected molecular mass predicted for the M. capsulatus ATP synthase complex, with a possible stoichiometric arrangement of F1a3:b3:g:d:e and F0a:b2:c10–14 as demonstrated for Escherichia coli (Pedersen and Amzel, 1993). Depending on the number of c subunits in the complex, which may vary among different bacteria, the complex in M. capsulatus (Bath) should be about 531–565 kDa.
complement of c-type cytochromes in cellular fractions of M. capsulatus, and how they can be identified by mass spectrometry combined with a c-type heme database constructed from the genome.
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4.1. Heme staining of M. capsulatus c-type heme proteins These methods take advantage of the intrinsic peroxidase activity of the covalently bound heme group of denaturated c-type cytochromes. The procedures described below are based on Dorward (1993), Thomas et al. (1976), and Vargas et al. (1993). 4.1.1. c-type heme staining on protein transfer membranes 1. SDS-PAGE is carried out without the addition of reducing agents. 2. Proteins are electrotransferred to a nitrocellulose/PVDF membrane at 100 V for 1 h. 3. The membrane is washed twice in Tris-buffered saline for 5 min. 4. The membrane is subsequently incubated with ECL as described by the manufacturer (GE Healthcare) and exposed to a Hyperfilm ECL (GE Healthcare) in the dark for 5 to 30 min, depending on the abundance of c-type heme proteins present in the sample. The membranes can be stained for total protein content with Amidoschwarz (nitrocellulose membrane) or Coomassie Blue (PVDF membrane) (Dunn, 1999). 4.1.2. In-gel c-type heme staining 1. SDS-PAGE is carried out without the addition of reducing agents. 2. Tetramethylbenzidin (TMBZ) is solubilized in methanol to a final concentration of 6.3 mM. 3. The polyacrylamide gel is covered with a solution consisting of freshly prepared TMBZ solution and 0.25 M sodium acetate (pH 5.0) in a 3:7 ratio, and incubated for 1–2 h in the dark with a gentle stirring every 10–15 min. 4. H2O2 is added to a final concentration of 30 mM. The staining of the c-type heme proteins should be visible within a few minutes and increases in intensity over the next 30 min. 5. The gel is placed in isopropanol and 0.25 M sodium acetate (pH 5.0) at a ratio of 3:7 in order to remove any precipitated TMBZ, to clear the gel background, and to increase the staining intensity. The acetate-buffered 30% isopropanol solution is replaced twice. 4.1.3. Identification of c-type heme proteins Figure 5.5 shows protein samples extracted from the M. capsulatus surface under different copper concentrations in the growth medium following staining for c-type heme with the protocols outlined above. A parallel Coomassie Blue stained gel of the same samples is used to locate the c-type heme-positive bands. These proteins can be excised from the gel, trypsinated and identified with mass spectrometry analyses. A refined database based on
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A kDa 181.8 115.5 82.2 64.2 48.8
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Figure 5.5 c-type heme staining of the M. capsulatus surfaceome obtained from cultures grown at different copper concentrations. Cultures were grown with copper concentrations corresponding to 0 (lane 1), 0.8 (lane 2), 1.6 (lane 3), 5 (lane 4), and 10 mM (lane 5) copper. (A) A 12.5 % polyacrylamide gel was used and stained with CBB. (B, C) c-type heme staining of (A) using ECL on nitrocellulose membranes (B) and in-gel staining (C). Arrowheads indicate gel bands analyzed by MS, and the numbers correspond to the numbers given in Table 2 in Karlsen et al. (2008). The figure was adopted from Karlsen et al. (2008) with kind permission from John Wiley & Sons, Inc.
proteins containing the c-type heme motif, CxxCH, is constructed from the ORF library, as described in Section 2.3.1. The CxxCH consensus sequence contains the two cysteines that covalently attach the heme group with thioether bonds, and the histidine that is one of the two axial ligands coordinating the heme iron (Barker and Ferguson, 1999). This arrangement is shared by all known c-type cytochromes, and this database will therefore comprise the full complement of putative c-type heme proteins.
ACKNOWLEDGMENTS We thank Johan Lillehaug for his participation and continuous support during our work. This work was supported in parts by grants from the Norwegian Research Council, The Meltzer Foundation, SUP140785/420 (GABI), and Norferm DA.
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Berven, F. S., Karlsen, O. A., Murrell, J. C., and Jensen, H. B. (2003). Multiple polypeptide forms observed in two-dimensional gels of Methylococcus capsulatus (Bath) polypeptides are generated during the separation procedure. Electrophoresis 24, 757–761. Berven, F. S., Flikka, K., Jensen, H. B., and Eidhammer, I. (2004). BOMP: A program to predict integral beta-barrel outer membrane proteins encoded within genomes of Gramnegative bacteria. Nucleic Acids Res. 32, W394–W399. Berven, F. S., Karlsen, O. A., Straume, A. H., Flikka, K., Murrell, J. C., Fjellbirkeland, A., Lillehaug, J. R., Eidhammer, I., and Jensen, H. B. (2006). Analysing the outer membrane subproteome of Methylococcus capsulatus (Bath) using proteomics and novel biocomputing tools. Arch. Microbiol. 184, 362–377. Bird, C. W., Lynch, J. M., Pirt, F. J., and Reid, W. W. (1971). Steroids and squalene in Methylococcus capsulatus grown on methane. Nature 230, 473–474. Camacho-Carvajal, M. M., Wollscheid, B., Aebersold, R., Steimle, V., and Schamel, W. W. (2004). Two-dimensional Blue native/SDS gel electrophoresis of multi-protein complexes from whole cellular lysates: A proteomics approach. Mol. Cell. Proteomics 3, 176–182. Davey, J. F., Whittenbury, R., and Wilkinson, J. F. (1972). The distribution in the methylobacteria of some key enzymes concerned with intermediary metabolism. Arch. Mikrobiol. 87, 359–366. Dorward, D. W. (1993). Detection and quantitation of heme-containing proteins by chemiluminescence. Anal. Biochem. 209, 219–223. Dunn, M. J. (1999). 2-D proteome analysis protocols. In “Methods in Molecular Biology, Vol. 112,” (A. J. Link, ed.).Humana Press Inc., Totowa. Exner, M. M., Doig, P., Trust, T. J., and Hancock, R. E. (1995). Isolation and characterization of a family of porin proteins from Helicobacter pylori. Infect. Immun. 63, 1567–1572. Fjellbirkeland, A., Kleivdal, H., Joergensen, C., Thestrup, H., and Jensen, H. B. (1997). Outer membrane proteins of Methylococcus capsulatus (Bath). Arch. Microbiol. 168, 128–135. Gao, J., and Chen, L. L. (2010). Theoretical methods for identifying important functional genes in bacterial genomes. Res. Microbiol. 161, 1–8. Hanczar, T., Csaki, R., Bodrossy, L., Murrell, J. C., and Kovacs, K. L. (2002). Detection and localization of two hydrogenases in Methylococcus capsulatus (Bath) and their potential role in methane metabolism. Arch. Microbiol. 177, 167–172. Heidelberg, J. F., Seshadri, R., Haveman, S. A., Hemme, C. L., Paulsen, I. T., Kolonay, J. F., Eisen, J. A., Ward, N., Methe, B., Brinkac, L. M., Daugherty, S. C., Deboy, R. T., et al. (2004). The genome sequence of the anaerobic, sulfate-reducing bacterium Desulfovibrio vulgaris Hildenborough. Nat. Biotechnol. 22, 554–559. Jahnke, L. L., Stan-Lotter, H., Kato, K., and Hochstein, L. I. (1992). Presence of methyl sterol and bacteriohopanepolyol in an outer-membrane preparation from Methylococcus capsulatus (Bath). J. Gen. Microbiol. 138, 1759–1766. Juncker, A. S., Willenbrock, H., Von Heijne, G., Brunak, S., Nielsen, H., and Krogh, A. (2003). Prediction of lipoprotein signal peptides in Gram-negative bacteria. Protein Sci. 12, 1652–1662. Karlsen, O. A., Lillehaug, J. R., and Jensen, H. B. (2008). The presence of multiple c-type cytochromes at the surface of the methanotrophic bacterium Methylococcus capsulatus (Bath) is regulated by copper. Mol. Microbiol. 70, 15–26. Loffler, C., Eberlein, C., Mausezahl, I., Kappelmeyer, U., and Heipieper, H. J. (2010). Physiological evidence for the presence of a cis-trans isomerase of unsaturated fatty acids in Methylococcus capsulatus Bath to adapt to the presence of toxic organic compounds. FEMS Microbiol. Lett. 308, 68–75.
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Madan Babu, M., and Sankaran, K. (2002). DOLOP—Database of bacterial lipoproteins. Bioinformatics 18, 641–643. Methe, B. A., Nelson, K. E., Eisen, J. A., Paulsen, I. T., Nelson, W., Heidelberg, J. F., Wu, D., Wu, M., Ward, N., Beanan, M. J., Dodson, R. J., Madupu, R., et al. (2003). Genome of Geobacter sulfurreducens: Metal reduction in subsurface environments. Science 302, 1967–1969. Murrell, J. C., and Dalton, H. (1983). Nitrogen fixation in obligate methanotrophs. J. Gen. Microbiol. 129, 3481–3486. Myers, C. R., and Myers, J. M. (2003). Cell surface exposure of the outer membrane cytochromes of Shewanella oneidensis MR-1. Lett. Appl. Microbiol. 37, 254–258. Pedersen, P. L., and Amzel, L. M. (1993). ATP synthases. Structure, reaction center, mechanism, and regulation of one of nature’s most unique machines. J. Biol. Chem. 268, 9937–9940. Poptsova, M. S., and Gogarten, J. P. (2010). Using comparative genome analysis to identify problems in annotated microbial genomes. Microbiology 156, 1909–1917. Reisinger, V., and Eichacker, L. A. (2006). Analysis of membrane protein complexes by blue native PAGE. Proteomics 6(Suppl. 2), 6–15. Remmert, M., Linke, D., Lupas, A. N., and Soding, J. (2009). HHomp—Prediction and classification of outer membrane proteins. Nucleic Acids Res. 37, W446–W451. Schagger, H., and von Jagow, G. (1991). Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal. Biochem. 199, 223–231. Schulz, G. E. (2000). beta-Barrel membrane proteins. Curr. Opin. Struct. Biol. 10, 443–447. Stainthorpe, A. C., Murrell, J. C., Salmond, G. P., Dalton, H., and Lees, V. (1989). Molecular analysis of methane monooxygenase from Methylococcus capsulatus (Bath). Arch. Microbiol. 152, 154–159. Stainthorpe, A. C., Lees, V., Salmond, G. P., Dalton, H., and Murrell, J. C. (1990). The methane monooxygenase gene cluster of Methylococcus capsulatus (Bath). Gene 91, 27–34. Stanley, S. H., Prior, S. D., Leak, D. J., and Dalton, H. (1983). Copper stress underlies the fundamental change in intracellular location of methane mono-oxygenase in methane utilizing organisms: Studies in batch and continuous cultures. Biotechnol. Lett. 5, 487–492. Stolyar, S., Costello, A. M., Peeples, T. L., and Lidstrom, M. E. (1999). Role of multiple gene copies in particulate methane monooxygenase activity in the methane-oxidizing bacterium Methylococcus capsulatus Bath. Microbiology 145(Pt 5), 1235–1244. Strom, T., Ferenci, T., and Quayle, J. R. (1974). The carbon assimilation pathways of Methylococcus capsulatus, Pseudomonas methanica and Methylosinus trichosporium (OB3B) during growth on methane. Biochem. J. 144, 465–476. Taylor, S. C., Dalton, H., and Dow, C. S. (1981). Ribulose-1,5-bisphosphate carboxylase/ oxygenase and carbon assimilation in Methylococcus capsulatus (Bath). J Gen Microbiol. 122, 89–94. Thomas, P. E., Ryan, D., and Levin, W. (1976). An improved staining procedure for the detection of the peroxidase activity of cytochrome P-450 on sodium dodecyl sulfate polyacrylamide gels. Anal. Biochem. 75, 168–176. Vargas, C., McEwan, A. G., and Downie, J. A. (1993). Detection of c-type cytochromes using enhanced chemiluminescence. Anal. Biochem. 209, 323–326. Ward, N., Larsen, O., Sakwa, J., Bruseth, L., Khouri, H., Durkin, A. S., Dimitrov, G., Jiang, L., Scanlan, D., Kang, K. H., Lewis, M., Nelson, K. E., et al. (2004). Genomic insights into methanotrophy: The complete genome sequence of Methylococcus capsulatus (Bath). PLoS Biol. 2, e303.
C H A P T E R
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Functional Metagenomics of Methylotrophs Marina G. Kalyuzhnaya,* David A. C. Beck,†,‡ and Ludmila Chistoserdova‡ Contents 1. Introduction 2. Enrichment for Specific Functional Types Using SIP 2.1. Sample collection and cell labeling 2.2. DNA extraction, isopycnic centrifugation, and labeled DNA recovery 3. Creating a Metagenomic Database and Linking It to Functionality 3.1. DNA sequencing and assembly 3.2. Gene-centric analysis 3.3. Organism-centric analysis 4. Ultrashort Read-Based Metatranscriptomics 4.1. Principle and strategy 4.2. RNA isolation 4.3. mRNA enrichment and (optional) cDNA synthesis 4.4. Data processing 4.5. Metatranscriptome coverage and specificity 5. Conclusions and Future Perspectives Acknowledgments References
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Abstract It is widely recognized that most microbes in the biosphere remain uncultured and unknown. In the recent few years, whole genome shotgun (WGS) sequencing of environmental DNA (metagenomics) has revolutionized the field of environmental microbiology by allowing one to tap into the genomic content of microbial communities in specific ecological niches, deducing information on their biochemical potentials. However, ascribing specific functions to specific organisms remains very difficult in most cases, due to low sequence coverage and the lack of * Department of Microbiology, University of Washington, Seattle, Washington, USA eScience Institute, University of Washington, Seattle, Washington, USA Department of Chemical Engineering, University of Washington, Seattle, Washington, USA
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Methods in Enzymology, Volume 495 ISSN 0076-6879, DOI: 10.1016/B978-0-12-386905-0.00006-1
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2011 Elsevier Inc. All rights reserved.
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sequence assembly that result from metagenomics of complex microbial communities. Therefore, methods that link specific biogeochemical processes to specific members of such complex natural communities are urgently needed. We have developed and implemented a functional metagenomics approach that allows such a connection via substrate-specific stable isotope labeling, followed by WGS sequencing of the labeled DNA to describe bacterial populations involved in metabolism of single-carbon compounds in a freshwater lake. We also developed a pipeline for community transcript analysis based on ultrashort read highthroughput sequencing of messenger RNA, matching these to a specific scaffold. The methodologies described in this chapter can be applied in a wide variety of ecosystems for targeting methylotrophs as well as other functional guilds of microbes.
1. Introduction Metagenomics is a fast growing and diverse field within environmental biology directed at obtaining knowledge on genomes of environmental microbes, without prior cultivation, as well as of entire microbial communities. When applied to communities of low complexity, exemplified by the communities of the acid mine drainage biofilm (Tyson et al., 2004) or the symbionts of a marine oligochaete (Woyke et al., 2006), the metagenomics approach, even at a modest sequencing effort, allows for sequence assembly. Thus, analysis of almost complete genomes of the dominant species in these communities can be carried out, including accurate metabolic reconstruction and even detection of strain-specific genomic variants. However, the situation is quite different when metagenomics is applied to communities of high complexity, such as the communities of marine habitats or soils (Rusch et al., 2007; Tringe et al., 2005; Venter et al., 2004). In these cases, significantly larger sequencing efforts resulted in very fragmented assemblies even for the most abundant species, with most of the datasets being represented by singleton sequencing reads. While gene-centric analysis (Tringe et al., 2005) can be conducted on the non-assembled metagenomic data and predictions on the major metabolic pathways can be made, the specific metabolic capabilities are hard or impossible to place into the context of individual species. This approach is especially vulnerable when used with short sequence reads produced by the next-generation sequencing technologies such as 454 Roche (Gomez-Alvarez et al., 2009; Wommack et al., 2008). One approach to directly link a function in the environment to a specific guild performing this function is through the use of a technique known as stable isotope probing (DNA-SIP). This involves feeding the natural population a substrate of interest, labeled by a heavy isotope (e.g., 13C), followed by characterization of the heavy fraction of communal DNA that should be enriched in DNA of microbes that actively metabolize the labeled substrate.
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This technique has been effective in identifying microbes involved in specific biogeochemical transformations such as methylotrophy, phenol degradation, glucose metabolism, etc. (Chen and Murrell, 2010; Friedrich, 2006; Madsen, 2006; Radajewski et al., 2000). Typically, small amounts of DNA are isolated in these experiments, and the DNA is used for phylogenetic profiling and detection of key functional genes, after PCR amplification (Friedrich, 2006). A modification of this approach has been described involving a multiple displacement amplification step, followed by metagenomic library construction and screening for specific marker genes (Chen and Murrell, 2010; Chen et al., 2008). However, we demonstrated that it is possible to scale up the SIP protocol to obtain amounts of DNA sufficient to enable the whole genome shotgun (WGS) sequencing approach, and we applied such an approach to characterize methylotroph communities of Lake Washington (Kalyuzhnaya et al., 2008). Methylotrophy is an important part of the global carbon cycle on this planet (Guenter, 2002; Hanson and Hanson, 1996). Identities of methylotrophs involved in utilization of specific C1 substrates (such as methane, methanol, methylated amines, etc.) in a variety of environments have previously been assessed by both culture-reliant and culture-independent methods (Chistoserdova et al., 2009), the former providing important models for understanding the specific biochemical pathways enabling methylotrophy, and the latter providing insights into species richness within specific functional groups. However, while genomic data for some model methylotrophs are now available (Chistoserdova et al., 2009), these may not represent major players in specific functional guilds. At the same time, current methods for environmental detection provide little insight into the genomic structure of uncultivated methylotrophs. The goal of the functional metagenomics approach is twofold: to reduce the complexity of a community and to directly link specific substrate repertoires of the community to specific functional guilds. The schematic of the enrichment and sequencing flow is shown in Fig. 6.1.
2. Enrichment for Specific Functional Types Using SIP 2.1. Sample collection and cell labeling 1. Samples are collected using an appropriate devise and transported to the laboratory on ice. Samples of Lake Washington sediment used here to demonstrate the method were collected as previously described (Kalyuzhnaya et al., 2004). 2. Microcosms are set up in conditions that mimic the in situ conditions, including substrate concentrations that should approximate the in situ
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12C-DNA 13C-DNA
13C-Methane 13C-Methanol
13C-Methylamine
13C-Formaldehyde 13C-Formate
Analysis Major players
Figure 6.1 General strategy for functional metagenomics exemplified by SIP-based enrichment for methylotrophic functional types.
concentrations. However, they should be high enough to allow for efficient labeling. In our case, each microcosms contained 10 ml of the sediment, 90 ml of Lake Washington water filtered through 0.22 mm filters (Millipore), and one of following 13C-labeled substrates: methane (50% of air), methanol (10 mM), methylamine (10 mM), formaldehyde (1 mM), and formate (10 mM; Kalyuzhnaya et al., 2008). 3. Samples are incubated, preferably at the in situ temperature, for the duration of time that allows some of the DNA to become labeled with 13 C (in Kalyuzhnaya et al., 2008, 3–14 days, dependent on the substrate, at room temperature with shaking were required). Other microcosm incubations were conducted at an in situ temperature (8 C), and lower concentrations of substrates were used, with similar results but requiring longer duration (not shown).
2.2. DNA extraction, isopycnic centrifugation, and labeled DNA recovery 1. Total DNA is extracted using an appropriate protocol for a given sample. PowerSoilR DNA isolation kit (MO Bio Laboratories, Inc., Carlsbad, CA, USA) produces good results with soil and sediment samples. 2. DNA is prepared for CsCl-ethidium bromide density gradient ultracentrifugation as previously described (Radajewski et al., 2000) and centrifuged at 265,000g (Beckman VTi 65 rotor) for 16 h at 20 C. DNA fractions are visualized in UV, and 13C-DNA fractions are collected using 19-gauge needles (Nercessian et al., 2005). DNA preparations may be subjected to a second round of CsCl-ethidium bromide density gradient ultracentrifugation, followed by a standard DNA purification
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procedure (e.g., Sambrook et al., 1989). Optionally, the fractions may be collected and analyzed for comparison.
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C-DNA
3. Creating a Metagenomic Database and Linking It to Functionality 3.1. DNA sequencing and assembly Shotgun libraries are constructed, one per each SIP microcosm. In our original study that utilized the Sanger technology (Kalyuzhnaya et al., 2008), they were constructed in the pUC18 vector (1–3 kb inserts) and vector inserts were sequenced with BigDye Terminators v3.1 and resolved with ABI PRISM 3730 (ABI) sequencers at the Joint Genome InstituteProduction Genomics Facility ( JGI-PGF; Walnut Creek, CA, USA). A total of 344,832 reads comprising 255 Mb of Phred Q20 sequence were generated. Sequences were stringently quality- and vector-trimmed using LUCY (Chou and Holmes, 2001), and the trimmed sequences were both by the individual sample and by en masse (combined assembly) using the PGA assembler. Assembly statistics for the datasets described by Kalyuzhnaya et al. (2008) are shown in Table 6.1. Sequences were automatically annotated and loaded into the JGI’s IMG/M system (http://img.jgi.doe.gov/cgi-bin/m/main.cgi). Sequence coverage and degree of assembly depended on the sequencing effort applied and on the species richness and evenness of the enriched communities. These draft quality assemblies were manually validated and used for all downstream analyses. Currently, the Sanger technology is being replaced by the next-generation sequencing technologies, the longer read length and the higher quality of the read being traded off for the higher throughput and the lower cost (Ansorge, 2009; Lapidus, 2009). With the data generated by these new technologies, new assembly tools and new quality controls are being developed (Lapidus, 2009; Miller et al., 2010). Concurrently, new sequencing technologies are in development, and when they enter the market, new bioinformatics tools will need to be implemented to process the next wave of genomic data types.
3.2. Gene-centric analysis 3.2.1. Phylogenetic markers The community complexity and content can be evaluated via profiling of 16S rRNA genes and other phylogenetic markers. In the case we describe here, community complexity was significantly reduced in microcosms exposed to each of the C1 substrates compared to the complexity of the non-enriched
Table 6.1 Summary of sequencing, assembly and gene prediction statistics
Assembly statistics Number of reads Average read length (bp) Trimmed read length (Mbp) Non-redundant sequence (bp) Percent of reads in contigs Total contigs (> 2 kb) Total singlets Average sequence coverage (x) Highest sequence coverage (x) Average size of contigs (bp) Largest contig (bp) GC content (%) Gene predictions Protein coding genes Genes in COGs Genes in Pfams Predicted enzymes Number of 16S rRNA genes Number of tRNA genes
Methane
Methanol
Methylamine
Formaldehyde
Formate
Combined
Methylotenera
71,808 792 56.85 52.16 10.2 2797 59,417 1.6 7.0 1418 6174 58.9
67,200 797 53.53 50.25 10.0 2871 56,408 1.6 4.8 1288 5913 59.5
83,712 709 59.34 37.23 55.5 7558 29,217 1.9 20.4 2065 20,771 53.0
80,640 712 58.91 57.62 7.3 2583 69,104 1.7 6.4 1166 4714 57.9
41,472 638 26.45 17.57 34.3 3618 18,857 1.9 4.7 1265 6276 65.8
344,832 741 255.08 211.47 27.6 25,877 215,581 1.7 23.1 1593 22,407 58.3
NA NA NA 11.16 100 4078 0 2.1 20.4 2736 15,820 46.2
81,076 43,456 28,090 3089 12 405
77,229 40,773 26,494 3047 12 412
54,340 33,643 23,586 5005 10 376
89,729 46,032 29,375 3065 18 504
28,700 17,112 10,585 1417 5 121
321,503 174,344 115,228 16,780 61 1728
12,719 10,082 8543 3264 3 181
This table is reproduced from the original publication (Kalyuzhnaya et al., 2008). The data currently posted in the JGI’s IMG/M interface differ slightly as a result of a more recent reanalysis by JGI.
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community that we estimate to be over 5000 species (Kalyuzhnaya et al., 2008), and shifted toward specific functional guilds that included both bona fide methylotrophs (Methylobacter tundripaludum, Methylomonas sp., Methylotenera mobilis, Methyloversatilis universalis, Ralstonia eutropha) and organisms only distantly related to any cultivated species, implicating the latter in environmental cycling of C1 compounds. The closest relatives of these included Verrucomicrobia, Nitrospirae, Planctomycetes, Acidobacteria, Cyanobacteria, and Proteobacteria. These data were supported by data on phylogenetic profiling of each metagenomic dataset, based on top BLAST hit distribution patterns (these data can be viewed as part of the IMG/M interface). 3.2.2. Functional genes Enrichment for specific functional genes can be addressed in a similar way, using known genes/proteins as queries in BLAST analyses. We used proteins involved in the reactions of the tetrahydromethanopterin pathway for C1 transfer that is a hallmark pathway in methylotrophs to demonstrate the function-relevant enrichment of the microcosm datasets (Table 6.2). In cases of highly divergent enzymes, multiple queries are required. For example, we used peptide sequences of fae and fae homologs belonging to different phylogenetic groups (Proteobacteria, Planctomycetes, and Archaea) to identify multiple and extremely divergent fae and fae-like sequences in our datasets (Fig. 6.2).
3.3. Organism-centric analysis 3.3.1. Characterizing genomes at high sequence coverage Genomes of individual organisms or populations of closely related strains may be present in a metagenome at high sequence coverage. Estimates of coverage for each organism can be initially gained from the coverage of individual 16S rRNA genes present in a metagenome. For example, Kalyuzhnaya et al. (2008) found that 16S rRNA genes belonging to M. mobilis were covered at up to 20. In such cases, it is reasonable to assume that complete or nearly complete genomes of respective strains may be present in a metagenomic dataset, and these can be extracted using one of the available binning tools (McHardy et al., 2007; Teeling et al., 2004; Tyson et al., 2004). In the metagenomic study of Lake Washington methylotroph populations, a composite genome of M. mobilis totaling slightly over 11 Mb was extracted from the methylamine microcosm metagenome using a compositional binning method PhyloPythia (McHardy et al., 2007; genome statistics are shown in Table 6.1). Genome completeness can be validated by examination of the presence of key metabolic and housekeeping genes (e.g., in Kalyuzhnaya et al., 2008). With satisfactory results, metabolism of an individual organism or a population of closely related strains (which will not necessarily be distinguished between by the
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Table 6.2 Representation of genes involved in tetrahydromethanopterin-linked formaldehyde oxidation in datasets generated in this work, compared to a soil metagenome (Tringe et al., 2005)
Dataset size (Mb)
a
Lake Washington sediment Minnesota farm soil Methane Methanol Methylamine Formaldehyde 100
52
50
37
Gene
Number of copies (coverage score, X)a
16S rRNA fae mtdB/mtdC mch fhcA fhcB fhcC fhcD mptG afp orf5 orf7 orf9 orf17 orf19 orf20 orf21 orf22 orfY
23 (11.5) 3 (1.5) 1 (0.5) 7 (3.5) 5 (2.5) 4 (2.0) 1 (0.5) 2 (1.0) 7 (3.5) 1 (0.5) 4 (2.0) 3 (1.5) 8 (4.0) 3 (6.0) 3 (1.5) 7 (3.5) 6 (3.0) 4 (2.0) 3 (1.5)
12 (13.6) 13 (8.1) 12 (9.7) 2 (2.3) 13 (11.7) 7 (6.3) 5 (2.5) 4 (3.0) 3 (1.5) 3 (1.5) 7 (4.3) 1 (0.5) 7 (5.0) 8 (5.1) 5 (3.8) 10 (7.1) 5 (3.4) 4 (2.0) 5 (3.8)
12 (18.8) 7 (2.9) 6 (3.0) 2 (1.0) 8 (5.1) 6 (5.2) 6 (3.7) 3 (2.2) 7 (4.3) 2 (2.0) 5 (4.2) 3 (1.5) 7 (6.0) 3 (4.1) 0 (0.0) 6 (3.0) 3 (1.5) 2 (1.0) 2 (1.0)
10 (42.5) 27 (45.0) 9 (12.4) 7 (10.1) 16 (18.1) 5 (7.0) 14 (17.6) 16 (21.8) 11 (16.1) 9 (15.7) 8 (13.6) 10 (15.9) 18 (23.5) 10 (13.7) 6 (10.5) 10 (19.1) 7 (20.8) 4 (6.6) 15 (18.9)
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18 (21.4) 11 (5.5) 4 (2.6) 3 (2.4) 6 (5.1) 3 (1.5) 4 (3.2) 4 (2.0) 4 (2.6) 0 (0.0) 2 (1.0) 3 (1.5) 5 (2.5) 2 (1.0) 2 (1.0) 2 (1.7) 2 (1.0) 0 (0.0) 3 (2.4)
Coverage score is calculated as a sum of average contig coverage (X) for each gene. For singleton reads, coverage was arbitrarily counted at 0.5X.
binning methods) can be reconstructed, and genome-wide comparisons may be carried out with other complete or composite genomes. For example, comparing the composite genome of M. mobilis to the complete genome of a close relative M. flagellatus, we were able to uncover examples of highly conserved parts of metabolism, including methylotrophy, as well as of non-conserved parts of metabolism, including non-homologous replacements in common biochemical pathways (Kalyuzhnaya et al., 2008). More recently we obtained the ultimate proof of the precision of the predictions derived from the composite M. mobilis genome (that we dubbed “high-resolution metagenomics”), by completely sequencing a genome of the type strain M. mobilis JLW8 and comparing it to the composite genome extracted from the metagenome (unpublished data).
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Betaproteobacterial Fae
Planctomycetal
Methylamine-specific
Gammaproteobacterial
Fae4
Archaeal
Fae2 Fae3
Figure 6.2 Phylogenetic diversity of Fae peptides identified in metagenomic datasets described in Kalyuzhnaya et al. (2008; only complete or nearly complete sequences were included). Red, methylamine microcosm, green, methane microcosm, blue, methanol microcosm, yellow, formaldehyde microcosm, purple, formate microcosm. Fae, formaldehyde activating enzyme. Fae2-4, homologs of Fae with no demonstrated function. The latter is only found in organisms not possessing H4MPT-linked C1 transfer pathway.
3.3.2. Characterizing genomes at low sequence coverage Binning tools such as PhyloPythia can also be applied to extract less covered genomes. However, in these cases, such approaches will unlikely produce complete genomes of individual organisms or populations. We found it helpful to supplement binning with the so-called protein recruitment technique. In this case a reasonably closely related genome sequence is required, as a reference. A convenient tool for protein recruitment is Phylogenetic Profiler that is part of the IMG/M package. Using the combination of PhyloPythia and Phylogenetic Profiler, we were able to extract an almost complete genome of and reconstruct metabolism of an uncultivated strain of M. tundripaludum and compare it to the genome of Methylococcus capsulatus (Kalyuzhnaya et al., 2008). If a very closely related genome is available as a reference, DNA recruitment can be used in place of protein recruitment (Rusch et al., 2007). Using this technique, we recovered a large portion of a R. eutropha genome (Kalyuzhnaya et al., 2008).
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4. Ultrashort Read-Based Metatranscriptomics 4.1. Principle and strategy For large-scale metatranscriptomics experiments, the next-generation sequencing technologies (Ansorge, 2009; Lapidus, 2009) are especially attractive as assembly is not a prerequisite for transcript analysis. The few metatranscriptomic studies published so far (Frias-Lopez et al., 2008; Gilbert et al., 2008; Urich et al., 2008) employed the 454 sequencing technology, as this technology produces reads of sufficient length to allow for functional predictions based on a single read. These reads are then processed in a genecentric way, overall producing very low resolution data limited to general function assignment for a fraction of reads while many of the reads remain unannotated. A much higher resolution is achieved when the 454 reads can be aligned with a genome of a close relative, presenting a principally different approach, similar to the method that is becoming increasingly popular as applied to single-organism transcriptomics, known as RNA-seq (Sorek and Cossart, 2010; van Vliet, 2009). However, the diversity of most communities is not well covered by sequenced genomes. Our own approach was to match transcripts produced by the Illumina technology (unpaired 75 bp reads) to the existing metagenomic scaffolds derived from the same environment. As the metagenome we employed as a scaffold has been generated from samples specifically enriched for methylotroph functional types (Kalyuzhnaya et al., 2008), as a proof of principle, for the transcriptomics experiments, we carried out enrichments using microcosm incubations, similar to the ones described above. However, only unlabeled substrates were used, and formate was omitted from the transcriptomics experiments. The resulting data were then pooled together. Obviously, the (incomplete) metagenomic scaffolds differ from the (complete, finished) scaffolds representing single genomes, in terms of both sequence coverage and sequence quality. However, we argue that when both a metagenome and a metatranscriptome originate from the same environment/condition, they should have a significant overlap. Most importantly, metatranscriptomes may be repeatedly matched to metagenomes after sequence space expansion (additional sequencing) and/or other iterations, such as an improved assembly.
4.2. RNA isolation The RNA extraction was performed as previously described (Nercessian et al., 2005) with the following modifications. Microcosm samples (0.5 g) were resuspended in 0.75 ml of RNA extraction buffer (0.15 mM NaH2PO4/Na2HPO4 buffer, pH 7.5; 5% CTAB, 1 M NaCl, 2% SDS;
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and 2% N-lauroylsarcosine sodium salt). The DNase I treatment was carried out using the DNAfree kit (BioLabs, Ambion) in accordance with the manufacturer’s instructions. The RNA samples were further purified using the RNeasy columns (Invitrogen, USA). An additional DNase I treatment was carried out directly on the RNeasy columns using the RNase-Free DNase Set (QIAGEN) in accordance with manufacturer’s instructions. The integrity of the RNA preparations was tested on a Bioanalyzer 2100 (Agilent), using Agilent RNA 6000 nano-kit, as suggested by the manufacturer.
4.3. mRNA enrichment and (optional) cDNA synthesis The content of ribosomal RNA was reduced in two steps: first using the MicroExpress Bacterial mRNA Purification Kit (Ambion) and then using the mRNA-ONLYTM Prokaryotic mRNA Isolation Kit (Epicentre Biotechnologies). The resulting mRNA-enriched samples (10 mg) were subjected to first-strand cDNA synthesis using 10 mM random hexamers (Qiagen) and the Omniscript Reverse Transcriptase (Qiagen). The double-stranded DNA was synthesized from single-stranded cDNA using the Exo-Klenow enzyme (Ambion) and standard conditions (Sambrook et al., 1989). The ds-cDNA samples were purified using the QIAquick PCR Purification Kit (Qiagen). The cDNA-synthesis steps can be omitted and the RNA sampled can be submitted to a sequencing facility where a sequencing platform-specific chemistry would be utilized to produce cDNA.
4.4. Data processing The pipeline we used to process and analyze the ultrashort read data is comprised of six steps: (1) the raw reads are subjected to quality assessment and (2) complexity filtering; (3) the reads are then aligned to specific scaffolds in nucleotide space with some allowances for in
