Methods in Enzymology 459 - Complex enzymes in microbial natural product biosynthesis: Polyketides, aminocoumarins and carbohydrates

METHODS IN ENZYMOLOGY Editors-in-Chief JOHN N. ABELSON AND MELVIN I. SIMON Division of Biology California Institute of ...

Author: DAVID A. HOPWOOD

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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, USA 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 2009 Copyright # 2009, 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: 978-0-12-374591-0 ISSN: 0076-6879 Printed and bound in United States of America 09 10 11 12 10 9 8 7 6 5 4 3 2 1

CONTRIBUTORS

Brian Douglas Ames Department of Molecular Biology and Biochemistry, Department of Chemistry, University of California, Irvine, California, USA Jesu´s F. Aparicio Institute of Biotechnology INBIOTEC, and Microbiology Area, Biology Faculty, University of Leoń, Leoń, Spain Patrick Caffrey School of Biomolecular and Biomedical Science, University College Dublin, Dublin, Ireland David E. Cane Department of Chemistry, Brown University, Box H, Providence, Rhode Island, USA Yolande A. Chan Department of Bacteriology, University of Wisconsin-Madison, Madison, Wisconsin, USA Yi-Qiang Cheng Department of Chemistry and Biochemistry, and Department of Biological Sciences, University of Wisconsin-Milwaukee, Milwaukee, Wisconsin, USA Tarun Chopra Chemical Biology Laboratory, National Institute of Immunology, New Delhi, India Jane M. Coughlin Department of Chemistry, University of Wisconsin-Madison, Madison, Wisconsin, USA Russell J. Cox School of Chemistry, University of Bristol, Bristol, United Kingdom John E. Cronan Department of Biochemistry and Department of Microbiology, University of Illinois, Urbana, Illinois, USA Zixin Deng Laboratory of Microbial Metabolism and School of Life Sciences and Biotechnology, Shanghai Jiaotong University, Shanghai, China xiii

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Tadashi Eguchi Department of Chemistry and Materials Science, Tokyo Institute of Technology, Tokyo, Japan Rajesh S. Gokhale Chemical Biology Laboratory, National Institute of Immunology, New Delhi, India Hugo Gramajo Instituto de Biologıá Molecular y Celular de Rosario (CONICET), Facultad de Ciencias Bioquı´micas y Farmaceúticas, Universidad Nacional de Rosario, Rosario, Argentina Lutz Heide Pharmazeutische Biologie, Pharmazeutisches Institut, Universita¨t Tu¨bingen, Tu¨bingen, Germany Geoffrey P. Horsman Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Hui Jiang Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Leonard Katz Synthetic Biology Engineering Research Center, University of California, Berkeley, Emeryville, California, USA Fumitaka Kudo Department of Chemistry, Tokyo Institute of Technology, Tokyo, Japan Steven G. Van Lanen Division of Pharmaceutical Sciences, College of Pharmacy, University of Kentucky, Lexington, Kentucky, USA Si-Kyu Lim Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Hung-wen Liu College of Pharmacy, Department of Chemistry and Biochemistry, and Institute for Cellular and Molecular Biology, University of Texas-Austin, Austin, Texas, USA Tiangang Liu Laboratory of Microbial Metabolism and School of Life Sciences and Biotechnology, Shanghai Jiaotong University, Shanghai, China, and Department of Chemistry, Brown University, Box H, Providence, Rhode Island, USA

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Juan F. Martıń Universidad de Leoń, Dpto. Biologıá Molecular – A´rea de Microbiologıá, Fac. CC. Biolo´gicas y Ambientales and Institute of Biotechnology INBIOTEC, Leoń, Spain Hugo G. Menzella Instituto de Biologıá Molecular y Celular de Rosario (CONICET), Facultad de Ciencias Bioquı´micas y Farmaceúticas, Universidad Nacional de Rosario, Rosario, Argentina Salvador Peiru´ Instituto de Biologıá Molecular y Celular de Rosario (CONICET), Facultad de Ciencias Bioquı´micas y Farmaceúticas, Universidad Nacional de Rosario, Rosario, Argentina Wolfgang Piepersberg Department of Chemical Microbiology, Bergische University Wuppertal, Wuppertal, Germany Scott R. Rajski Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Christopher D. Reeves Amyris Biotechnologies, Inc., Emeryville, California, USA Eduardo Rodriguez Instituto de Biologıá Molecular y Celular de Rosario (CONICET), Facultad de Ciencias Bioquı´micas y Farmaceúticas, Universidad Nacional de Rosario, Rosario, Argentina Ben Shen Department of Chemistry and Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Thomas J. Simpson School of Chemistry, University of Bristol, Bristol, United Kingdom Yi Tang Department of Chemical and Biomolecular Engineering, University of CaliforniaLos Angeles, Los Angeles, California, USA Christopher J. Thibodeaux Institute for Cellular and Molecular Biology, University of Texas-Austin, Austin, Texas, USA Jacob Thomas Department of Microbiology, University of Illinois, Urbana, Illinois, USA

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Michael G. Thomas Department of Bacteriology, University of Wisconsin-Madison, Madison, Wisconsin, USA Shiou-Chuan (Sheryl) Tsai Department of Molecular Biology and Biochemistry, Department of Chemistry, Department of Pharmaceutical Sciences, University of California, Irvine, California, USA Udo F. Wehmeier Department of Sports Medicine, Bergische University Wuppertal, Wuppertal, Germany Kira J. Weissman Department of Pharmaceutical Biotechnology, Saarland University, Saarbru¨cken, Germany Jessica White-Phillip Institute for Cellular and Molecular Biology, University of Texas-Austin, Austin, Texas, USA Wenjun Zhang Department of Chemical and Biomolecular Engineering, University of CaliforniaLos Angeles, Los Angeles, California, USA Sergey Zotchev Department of Biotechnology, Norwegian University of Science and Technology, Trondheim, Norway

PREFACE

The complex structures of microbial natural products have fascinated chemists for decades. As the tools of chemistry and biochemistry were sharpened, huge advances in understanding natural product biosynthesis were made, but there were still barriers to a satisfactory understanding. Many such impediments were due to the instability of intermediates in the biosynthetic pathways, which hampered chemical analysis. At the same time, a frequent inability to obtain active cell–free preparations severely limited the success of biochemical approaches. A striking example of these limitations is provided by the polyketides, the largest and most important family of secondary metabolites. Chemistry and biochemistry had deduced the relationships between polyketide and fatty acid biosynthesis and had revealed the basic biochemical reactions involved, but there was little understanding of the ‘‘programming’’ of the enzymes, that is control of the variables that make the polyketides such a varied class of chemicals: choice of starter and extender units for carbon chain building, and control of chain length, degree of reduction of keto groups, and chirality of carbon and hydroxyl branches. Isolation of the actinomycete gene clusters that encode the polyketide synthases, their sequencing, and their manipulation into unnatural combinations in the early 1990s changed the landscape almost overnight. There followed a period in which genetics provided the primary stimulus to much of the research in natural product biosynthesis. Now, chemistry, genetics, enzymology, and structural studies are working synergistically to reveal the details of biochemical control. This two-volume set of Methods in Enzymology reflects these developments in the study of natural product biosynthesis. As expressed by Mel Simon in his invitation to edit the set, it is especially timely in view of the increasing need for novel bioactive natural products, especially antibiotics and anticancer drugs, and the new possibilities for addressing this need by carrying out ‘‘chemistry through genetics’’ and by studying the gamut of potential natural products revealed by the sequencing of microbial genomes. We begin Volume A with the isolation and screening of various kinds of microorganisms, to provide the raw material for subsequent fundamental studies or for the development of natural products as drugs. Then come three chapters dealing with the regulation of secondary metabolite production in actinomycetes—the group of filamentous soil bacteria that are preeminent secondary metabolite producers—and how an understanding of such regulation can furnish compounds that would otherwise be xvii

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hard to obtain. Next are chapters covering the cloning and analysis of biosynthetic pathway genes and computer-based methods for predicting the products encoded by gene sets for two key classes of secondary metabolites, the polyketides and nonribosomal peptides, from DNA sequence data, as well as articles describing innovative approaches to probing their biosynthesis. Two final chapters in the first section deal with the biosynthesis of sugars and their attachment to secondary metabolite aglycones, thereby conferring biological activity. The section on peptide natural products begins with an overview of nonribosomal peptide biosynthesis, followed by a detailed description of methods for studying the biosynthesis of the amino acids and other precursors that function as building blocks in their assembly, as well as a chapter on the heterologous expression of nonribosomal peptide synthetase genes. Next come chapters on a specific class of compounds in this superfamily, the cephem beta-lactams, on a special type of iron-chelating siderophore, and on the important glycopeptide and lipopeptide families of antibiotics. Moving to ribosomally synthesized peptide natural products, two chapters cover the lantibiotics, a topic of increasing current focus in the search for antibiotics effective against resistant pathogens. We end Volume B with another example of ribosomally synthesised peptides, this time coupled with techniques for metagenomics mining. Volume B is dominated by the polyketides, reflecting their preeminence as natural products. Kira Weissman introduces polyketide synthesis and the different types of polyketide synthases, and puts the 16 chapters in this section elegantly into context, making redundant any further remarks here, except to note the absence of a chapter on the type III polyketide synthases, an omission stemming from the last-minute withdrawal of the author chosen for this topic. The section on aminocoumarins contains a single chapter that provides a particularly fine example of the application of molecular genetics to another class of compounds, with considerable potential for the generation of ‘‘unnatural natural products’’ by genetic engineering techniques first developed for the polyketides. The volume ends with a section on carbohydrate-type natural products, with two chapters on aminoglycosides and one on the biosynthesis of the TDPdeoxysugars that play such a crucial role in conferring biological activity on a whole range of secondary metabolites, harking back to the chapters on sugar biosynthesis in Volume A. Inevitably, the choice of topics to include in these volumes is somewhat arbitrary. The peptides and polyketides chose themselves because of their importance among natural products, especially as antibiotics, and because of the huge amount of recent research devoted to them. Historically, the aminoglycosides were centre-stage in the early days of antibiotic discovery—streptomycin was the first important actinomycete antibiotic to be described, and only the second, after penicillin, from any source to be a

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medical marvel—and they probably still make up the third largest chemical family of antibiotics, earning them a place in Volume B. Several other classes of microbial natural products were contenders for inclusion: aminocoumarins, terpenoids, and tetrapyrroles, among others. However, space constraints precluded inclusion of all of them, and in the end only the aminocoumarins made the cut. Hopefully, other classes will take their place in a further volume in due course, along with fuller coverage of natural product production by a wider range of microorganisms outside of the actinomycetes. I am most grateful for the enthusiastic response that greeted my invitations to contribute to this project. Inevitably, leaders in the field have many calls on their time, but it was most gratifying that nearly all my invitees either accepted or offered suggestions for alternative authors. I am especially grateful to Greg Challis, Chaitan Khosla, Tom Simpson, and Chris Walsh for their insightful ideas. To those who accepted—as well as to the many co-authors who were recruited to the writing—thank you for the time and effort that went into the preparation of the chapters and to the friendly way in which you all responded to my—usually minor—editorial suggestions, making my task a very pleasant one. DAVID A. HOPWOOD

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 xxi

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


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

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


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

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


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

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


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

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


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

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


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

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


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VOLUME 239. Nuclear Magnetic Resonance (Part C) Edited by THOMAS L. JAMES AND NORMAN J. OPPENHEIMER VOLUME 240. Numerical Computer Methods (Part B) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 241. Retroviral Proteases Edited by LAWRENCE C. KUO AND JULES A. SHAFER VOLUME 242. Neoglycoconjugates (Part A) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 243. Inorganic Microbial Sulfur Metabolism Edited by HARRY D. PECK, JR., AND JEAN LEGALL VOLUME 244. Proteolytic Enzymes: Serine and Cysteine Peptidases Edited by ALAN J. BARRETT VOLUME 245. Extracellular Matrix Components Edited by E. RUOSLAHTI AND E. ENGVALL VOLUME 246. Biochemical Spectroscopy Edited by KENNETH SAUER VOLUME 247. Neoglycoconjugates (Part B: Biomedical Applications) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 248. Proteolytic Enzymes: Aspartic and Metallo Peptidases Edited by ALAN J. BARRETT VOLUME 249. Enzyme Kinetics and Mechanism (Part D: Developments in Enzyme Dynamics) Edited by DANIEL L. PURICH VOLUME 250. Lipid Modifications of Proteins Edited by PATRICK J. CASEY AND JANICE E. BUSS VOLUME 251. Biothiols (Part A: Monothiols and Dithiols, Protein Thiols, and Thiyl Radicals) Edited by LESTER PACKER VOLUME 252. Biothiols (Part B: Glutathione and Thioredoxin; Thiols in Signal Transduction and Gene Regulation) Edited by LESTER PACKER VOLUME 253. Adhesion of Microbial Pathogens Edited by RON J. DOYLE AND ITZHAK OFEK VOLUME 254. Oncogene Techniques Edited by PETER K. VOGT AND INDER M. VERMA VOLUME 255. Small GTPases and Their Regulators (Part A: Ras Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 256. Small GTPases and Their Regulators (Part B: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL

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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 VOLUME 274. RNA Polymerase and Associated Factors (Part B) Edited by SANKAR ADHYA


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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 VOLUME 293. Ion Channels (Part B) Edited by P. MICHAEL CONN

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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 VOLUME 311. Sphingolipid Metabolism and Cell Signaling (Part A) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN


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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 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

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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 VOLUME 345. G Protein Pathways (Part C: Effector Mechanisms) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT


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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 VOLUME 362. Recognition of Carbohydrates in Biological Systems (Part A) Edited by YUAN C. LEE AND REIKO T. LEE

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VOLUME 363. Recognition of Carbohydrates in Biological Systems (Part B) Edited by YUAN C. LEE AND REIKO T. LEE VOLUME 364. Nuclear Receptors Edited by DAVID W. RUSSELL AND DAVID J. MANGELSDORF VOLUME 365. Differentiation of Embryonic Stem Cells Edited by PAUL M. WASSAUMAN AND GORDON M. KELLER VOLUME 366. Protein Phosphatases Edited by SUSANNE KLUMPP AND JOSEF KRIEGLSTEIN VOLUME 367. Liposomes (Part A) Edited by NEJAT DU¨ZGU¨NES, VOLUME 368. Macromolecular Crystallography (Part C) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 369. Combinational Chemistry (Part B) Edited by GUILLERMO A. MORALES AND BARRY A. BUNIN VOLUME 370. RNA Polymerases and Associated Factors (Part C) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 371. RNA Polymerases and Associated Factors (Part D) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 372. Liposomes (Part B) Edited by NEJAT DU¨ZGU¨NES, VOLUME 373. Liposomes (Part C) Edited by NEJAT DU¨ZGU¨NES, VOLUME 374. Macromolecular Crystallography (Part D) Edited by CHARLES W. CARTER, JR., AND ROBERT W. SWEET VOLUME 375. Chromatin and Chromatin Remodeling Enzymes (Part A) Edited by C. DAVID ALLIS AND CARL WU VOLUME 376. Chromatin and Chromatin Remodeling Enzymes (Part B) Edited by C. DAVID ALLIS AND CARL WU VOLUME 377. Chromatin and Chromatin Remodeling Enzymes (Part C) Edited by C. DAVID ALLIS AND CARL WU VOLUME 378. Quinones and Quinone Enzymes (Part A) Edited by HELMUT SIES AND LESTER PACKER VOLUME 379. Energetics of Biological Macromolecules (Part D) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS VOLUME 380. Energetics of Biological Macromolecules (Part E) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS VOLUME 381. Oxygen Sensing Edited by CHANDAN K. SEN AND GREGG L. SEMENZA


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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 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

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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 VOLUME 417. Functional Glycomics Edited by MINORU FUKUDA VOLUME 418. Embryonic Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA


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VOLUME 419. Adult Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 420. Stem Cell Tools and Other Experimental Protocols Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 421. Advanced Bacterial Genetics: Use of Transposons and Phage for Genomic Engineering Edited by KELLY T. HUGHES VOLUME 422. Two-Component Signaling Systems, Part A Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 423. Two-Component Signaling Systems, Part B Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 424. RNA Editing Edited by JONATHA M. GOTT VOLUME 425. RNA Modification Edited by JONATHA M. GOTT VOLUME 426. Integrins Edited by DAVID CHERESH VOLUME 427. MicroRNA Methods Edited by JOHN J. ROSSI VOLUME 428. Osmosensing and Osmosignaling Edited by HELMUT SIES AND DIETER HAUSSINGER VOLUME 429. Translation Initiation: Extract Systems and Molecular Genetics Edited by JON LORSCH VOLUME 430. Translation Initiation: Reconstituted Systems and Biophysical Methods Edited by JON LORSCH VOLUME 431. Translation Initiation: Cell Biology, High-Throughput and Chemical-Based Approaches Edited by JON LORSCH VOLUME 432. Lipidomics and Bioactive Lipids: Mass-Spectrometry–Based Lipid Analysis Edited by H. ALEX BROWN VOLUME 433. Lipidomics and Bioactive Lipids: Specialized Analytical Methods and Lipids in Disease Edited by H. ALEX BROWN VOLUME 434. Lipidomics and Bioactive Lipids: Lipids and Cell Signaling Edited by H. ALEX BROWN VOLUME 435. Oxygen Biology and Hypoxia Edited by HELMUT SIES AND BERNHARD BRU¨NE

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VOLUME 436. Globins and Other Nitric Oxide-Reactive Protiens (Part A) Edited by ROBERT K. POOLE VOLUME 437. Globins and Other Nitric Oxide-Reactive Protiens (Part B) Edited by ROBERT K. POOLE VOLUME 438. Small GTPases in Disease (Part A) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 439. Small GTPases in Disease (Part B) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 440. Nitric Oxide, Part F Oxidative and Nitrosative Stress in Redox Regulation of Cell Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 441. Nitric Oxide, Part G Oxidative and Nitrosative Stress in Redox Regulation of Cell Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 442. Programmed Cell Death, General Principles for Studying Cell Death (Part A) Edited by ROYA KHOSRAVI-FAR, ZAHRA ZAKERI, RICHARD A. LOCKSHIN, AND MAURO PIACENTINI VOLUME 443. Angiogenesis: In Vitro Systems Edited by DAVID A. CHERESH VOLUME 444. Angiogenesis: In Vivo Systems (Part A) Edited by DAVID A. CHERESH VOLUME 445. Angiogenesis: In Vivo Systems (Part B) Edited by DAVID A. CHERESH VOLUME 446. Programmed Cell Death, The Biology and Therapeutic Implications of Cell Death (Part B) Edited by ROYA KHOSRAVI-FAR, ZAHRA ZAKERI, RICHARD A. LOCKSHIN, AND MAURO PIACENTINI VOLUME 447. RNA Turnover in Bacteria, Archaea and Organelles Edited by LYNNE E. MAQUAT AND CECILIA M. ARRAIANO VOLUME 448. RNA Turnover in Eukaryotes: Nucleases, Pathways and Analysis of mRNA Decay Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 449. RNA Turnover in Eukaryotes: Analysis of Specialized and Quality Control RNA Decay Pathways Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 450. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON


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VOLUME 451. Autophagy: Lower Eukaryotes and Non-Mammalian Systems (Part A) Edited by DANIEL J. KLIONSKY VOLUME 452. Autophagy in Mammalian Systems (Part B) Edited by DANIEL J. KLIONSKY VOLUME 453. Autophagy in Disease and Clinical Applications (Part C) Edited by DANIEL J. KLIONSKY VOLUME 454. Computer Methods (Part A) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 455. Biothermodynamics (Part A) Edited by MICHAEL L. JOHNSON, JO M. HOLT, AND GARY K. ACKERS (RETIRED) VOLUME 456. Mitochondrial Function, Part A: Mitochondrial Electron Transport Complexes and Reactive Oxygen Species Edited by WILLIAM S. ALLISON AND IMMO E. SCHEFFLER VOLUME 457. Mitochondrial Function, Part B: Mitochondrial Protein Kinases, Protein Phosphatases and Mitochondrial Diseases Edited by WILLIAM S. ALLISON AND ANNE N. MURPHY VOLUME 458. Complex Enzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles and Peptides Edited by DAVID A. HOPWOOD VOLUME 459. Complex Enzymes in Microbial Natural Product Biosynthesis, Part B: Polyketides, Aminocoumarins and Carbohydrates Edited by DAVID A. HOPWOOD

C H A P T E R

O N E

Introduction to Polyketide Biosynthesis Kira J. Weissman Contents 3 6 6 8 9 12 13 13

1. Introduction 1.1. Types of PKS 1.2. Type II PKS 1.3. Type III PKS 1.4. Type I PKS 1.5. Combinatorial biosynthesis: Prospects and progress Acknowledgment References

1. Introduction Even among natural product-derived medicines, the polyketides stand out. The fungal-derived statins (Fig. 1.1) are the most successful cardiovascular drugs of all time (Roberts, 1996; Shepherd, 2006). Erythromycin A, a macrolide antibacterial, has been in clinical usage for 53 years (Washington and Wilson, 1985), while its semisynthetic derivative azithromycin (Zithromax) is one of the best-selling antibiotics in the world. The enediynes, relative newcomers to the polyketide stage, are among the most powerful anticancer agents yet discovered (Galm et al., 2005). More than a third of natural product and natural product-derived compounds approved as drugs in the 2005–2007 period were polyketides, while a host of others are advancing through clinical trials (Butler, 2008). As testimony to their importance, annual sales of polyketide-derived medicines routinely top US$20 billion. The endless variety of polyketide shape and structure (Fig. 1.1) continues to fascinate synthetic and natural product chemists alike. While synthetic

Department of Pharmaceutical Biotechnology, Saarland University, Saarbru¨cken, Germany Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04601-1

#

2009 Elsevier Inc. All rights reserved.

3

HO

O

O

O

OH

HO

O O

Mycolic acid Cell membrane component

HO O H

O O

Lovastatin Cholesterol lowering agent

HO

OH OH

O

O

O

OMe OMe MeO OH H O N O

OMe

O

O

OH

OH O MeHN

Erythromycin A Antibiotic

O

HO

OH OMe

Neocarzinostatin Anti-cancer

OH

Pederin Anti-cancer

MeO OH

O

HO2C

HO

O

OH

O

O

O

O

O

OH

HO

OH

OH OH NH2

OH OH

O

H2N OH

O

OH

O

Orsellinic acid

O O

O

OAc Squalestatin S1 Cholesterol lowering agent

OH O O

COOH

O 6-Methyl salicylic acid Antibiotic

O

Doxorubicin Anti-cancer

OH

OH OH O

OH OH

Amphotericin B Anti-fungal

HO

OH

O OH

OH

O CO2H HO HO2C O

OH

O

O

OH

O

OH

Monensin A Antibiotic

O

O

H H O

HO2C

O

OMe

O

MeO O

O

NMe2

HO O

O HO

O

O

O O

O

O O

Maitotoxin Cytotoxin

O

O

OH

HO HOO COOH

HO OH

20:5n3 Eicosapentaenoic acid Membrane components

O O

OH OH

NaO SO 3 OH OH

O

O

OSO3Na HO HO

OH

O

O OH

OH

O OH

OH O

O O

O OH

O

OH O

OH

O

OH OH

HO

O OH

Figure 1.1 Structures of representative polyketides, including prominent bioactivities.

OH OH

Introduction to Polyketide Biosynthesis

5

chemists tackle the challenge of recreating the compounds in the laboratory, natural products researchers try to deduce how nature went about constructing the molecules. The smallest polyketides, such as triacetic acid lactone (TAL) and 6-methylsalicylic acid (6-MSA), incorporate a mere six to eight carbons, while the biggest, maitotoxin, weighs in at a staggering 3422 daltons, with 164 carbons in the chain; in fact, maitotoxin is the largest, and most toxic nonbiopolymer known (Murata et al., 1994). A considerable number of polyketides, including doxorubicin, are highly rigidified, multicyclic aromatic compounds, derived from simple polyketone chains by alternative modes of folding and ring formation. Others, like erythromycin A, are substantially reduced, and are instead constrained into an active conformation by intramolecular end-to-end cyclization. Some polyketides masquerade as simpler metabolites—for example, the straightchain polyunsaturated fatty acids (PUFAs) of deep-sea microbes. In contrast, the intricately folded polyethers like monensin are origami-like in their complexity. Most polyketides bristle with diverse functionalities, but a few, including the enediynes, are also notable for their high reactivity. The majority of polyketide skeletons are elaborated by post-assembly reactions, many of which are required for the biological activity of the products; oxygenative tailoring is most common, followed closely by group transfer reactions, such as glycosylation and acylation. By the late 1980s, it was evident that despite their vast structural heterogeneity, all polyketides are assembled by a process closely resembling that of fatty acid formation (covered in Chapter 17 of this volume). Simple carboxylic acid derivatives (e.g., acetyl-CoA, malonyl-CoA, and methylmalonyl-CoA) are concatenated into linear chains by iterative Claisen condensation, followed in some cases by reductive modification of the resulting b-ketone groups (Fig. 1.2). However, the biochemical basis for programming alternative modes of polyketide construction to yield polyketones, partially reduced polyketides or fully reduced fatty acids, remained obscure; no obvious mechanism was available to explain the choice or number of building blocks and the order in which they are incorporated, the extent of redox adjustment following each round of chain extension, and the pattern of cyclization or lactonization of the resulting chains. The answer to this conundrum has emerged over several decades through sequencing and biochemical studies of hundreds of genes encoding for polyketide synthases (PKSs) and associated decorating enzymes; conveniently, these biosynthetic genes are often clustered together within the genomes of the producing organisms. PKS machineries turn out to be as complicated and varied as the molecules they produce—indeed, 10 chapters in this volume alone are dedicated to describing different types of PKS. In general, as the biosynthetic apparatus increases in sophistication, so does the complexity of the product.

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Kira J. Weissman

O ACP/ CoA

O

O

S

R

OH

Condensation

O ACP/ CoA

KR

O R

O

KS

KS

S

O

S

Reduction

O ACP

S

O

HO

n Polyketone (type II, type III, fungal NR)

O

OH R

DH Dehydration

O

ER

O

O

ACP

S

R

Reduction

O ACP

S

OH O

HO

O R

Reduced polyketide (modular type I (cis and trans), fungal PR and HR, PKSE)

R

HO

n Fatty acid (PUFA synthases)

Figure 1.2 Generic reaction scheme for biosynthesis of both fatty acids and polyketides, illustrating how the basic reaction cycle of fatty acid biosynthesis can be shunted to generate diverse polyketide structures, incorporating different levels of reduction. PKSs that practice each type of chemistry are indicated.

1.1. Types of PKS PKSs have been classified into three broad categories: type I and type II— reflecting their relationship to previously characterized enzymes of fatty acid biosynthesis—and type III (an example of each type is given in Fig. 1.3, and the characteristic features are summarized in Table 1.1). However, these distinctions are neither hard nor fast, as many PKSs exhibit hybrid behavior (Moss et al., 2004; Shen, 2003; Wenzel and Mu¨ller, 2005a). Type II and III PKSs both generate aromatic molecules, but use a fundamentally different approach to building a reactive polycarbonyl chain, and channeling it into a particular cyclic fate.

1.2. Type II PKS Type II PKSs are iteratively acting complexes of discrete proteins, each of which has one or more specific functions in the pathway (Hertweck et al., 2007 and references therein). Chain initiation (most often with acetate) and elongation with malonate are accomplished by the ‘‘minimal PKS’’ which consists of two ketosynthase-like condensing enzymes (KSa and KSb, only the first of which contributes the active site for condensation), and an acyl carrier protein (ACP), to which the growing chain is covalently tethered as a thioester. A malonyl-CoA:ACP transferase (MCAT) activity likely helps recruit the extender units from primary metabolism. Additional enzymatic subunits (ketoreductases [KR], cyclases [CYC], and aromatases [ARO]) cooperate with the minimal PKS to direct the folding pattern of the nascent polyketide chain. The polyphenolic products of the PKS are often

7


A Type I PKS (noniterative) Load AT

Module 1

ACP

KS

KR

AT

S

ACP

KS

AT

ACP

KS

AT

DH

KR

ACP

S

S O

Offload

Module 3

Module 2

TE

S

O

O

OH

O

O

R OH

O

R OH R Reduced polyketide

B Type II PKS (iterative) n times KSa

KSb

KSa

ACP x S

S O

KSb

ACP x

y

y

SH

S

O

O

O

O

R O

n

R Polyketone

C Type III PKS (iterative and ACP-independent) CoAS KS

O O

S O R

O

n times

CoAS O

KS SH

O

n

R Polyketone

Figure 1.3 Schematic of the three types of PKS. (A) Type I PKSs consist of multifunctional polypeptides. Each subunit contains one or more modules that incorporate a set of individually folded catalytic domains. Each module typically acts once (noniteratively) to accomplish one round of chain extension and associated reductive processing reactions. For each module, both the building block added and the functionality established by the reductive activities are indicated (by color-coded bold bonds, and shading, respectively). (B) Type II PKSs comprise discrete catalytic functions that associate into a productive complex. The ‘‘minimal PKS’’ includes the KSa, KSb, and ACP domains, which iterate through a defined number of chain-extension cycles to construct a polyketone chain. The exact stoichiometry of the KSa/KSb/ACP complex remains to be determined. (C) Type III PKSs consist of a single multifunctional active site that, in cooperation with CoA-bound substrates, performs all the steps necessary to assemble a polyketone chain of defined length.

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Kira J. Weissman

Table 1.1 Characteristic features of type I, II, and III PKS Type of PKS

Mode operation

Mode of substrate activation

Typical product

I I II III

Modular Iterative Iterative Iterative

ACP ACP ACP CoA

Reduced Aromatic and reduced Aromatic Aromatic

extensively remodeled by oxygenases, and decorated by glycosyl- and N-, O-, and C-methyltransferases (Rix et al., 2002 and references therein). Despite these broad-brush functional assignments deduced from structural, genetic, and in vitro studies, the exact roles of many type II PKS components remain uncertain. In addition, there is likely to be a significant degree of functional interdependence, hindering efforts to analyze the enzymes in isolation. Another complicating factor is the high inherent reactivity of the intermediate poly-b-ketone chains, which often leads to spontaneous chemistry when a reconstituted PKS is incomplete. The latest test tubebased approaches to precisely define the function of type II PKS enzymes and modifying activities, and how these efforts are likely to inform strategies to reconfigure aromatic polyketide biosynthesis by genetic engineering are reviewed in Chapter 16 of this volume.

1.3. Type III PKS In contrast to the multienzyme organization of type II PKSs, enzymes from the type III PKS superfamily of plants, fungi, and bacteria (also known as chalcone and stilbene synthases) use a single KS-like active site to catalyze the repetitive condensation of acetate units to a CoA-derivatized starter molecule, typically yielding mono- and bi-cyclic aromatic products (Austin and Noel, 2003, and references therein). Chain extension is often followed by intramolecular condensation and aromatization of the linear intermediate, all within the same PKS active site cavity. This ‘‘u¨ber-adaptable’’ group of catalysts (Austin et al., 2008) generates high diversity by varying the choice of acyl-CoA starter unit, the number of elongation steps (1 to 8), and the mechanism of cyclization. Downstream enzymes cause additional pathway branching, variably transforming the initial scaffolds to produce a range of different compounds. The relative architectural simplicity of the type III systems has enabled the rapid development of a mechanistic framework for these enzymes, providing insight into the factors mediating starter unit selection, chain extension and the control of ring formation. Nonetheless, several fundamental questions remain, particularly concerning the fungal and bacterial variants, which should be the subject of future work in the field.


9

1.4. Type I PKS Type I PKSs have proven to be both the most complicated, and yet most versatile of the three classes of catalyst. The diagnostic feature of these systems is the presence of multiple active sites within each polypeptide. In the iterative subcategory of type I PKS, a single multienzyme is used repeatedly. Despite this mechanistic commonality, this machinery is employed to assemble very different types of metabolites in a variety of microorganisms. Fungi contain three categories of iterative type I PKSs, termed nonreducing (NR), partially reducing (PR) and highly reducing (HR), reflecting the increasing incorporation of ketoreductase (KR), dehydratase (DH), and methylase (MT) activities into the multienzymes (Cox, 2007, and references therein). Correspondingly, their products range from very simple (e.g., orsellinic acid) to the highly complex (e.g., squalestatin S1) (Fig. 1.1). The functions of all domains in the NR-type PKS have recently been determined (Crawford et al., 2008; Ma et al., 2006), but the mechanism of the PR and HR synthases remains poorly understood. Here, the central challenge is to elucidate the molecular basis for the sophisticated programming of the PKSs, which allows a different subset of the constituent active sites to operate in each round of chain extension. The latest methodologies for studying these complex enzyme systems, both in vitro and in vivo, and the new insights that are emerging are reported in Chapter 3 of this volume. Nature also exploits a type I iterative PKS to generate long-chain polyunsaturated fatty acids (PUFAs) in psychrophilic marine microbes. These C20þ metabolites are thought to adapt the biophysical properties of the microbial cell membrane to the frigidity of polar and deep-sea environments (Metz et al., 2001; Napier, 2002; Nichols et al., 1999; Wallis et al., 2002). PKS-based PUFA biosynthesis actually came as a surprise, because until 2001, it was believed to be the preserve of terrestrial eukaryotes. In this more conventional pathway, unsaturated fatty acids are derived by elongation and subsequent desaturation of a precursor saturated fatty acid (Kaulmann and Hertweck, 2002). The PUFA PKSs, of which there are several variants, are distinct in both organization and mechanism from the fungal iterative PKS enzymes, and notably contain as many as nine tandem ACP domains ( Jiang et al., 2008). The unexpected discovery of these catalysts, as well as ongoing efforts to unravel their unique mode of biosynthesis are discussed in Chapter 4 of this volume. Chapter 5 of this volume describes the identification and characterization of the third type of iterative type I PKS, the PKSE family (Van Lanen and Shen, 2008 and references therein). These PKSs participate in formation of the enediyne class of polyketides in bacteria, and show sequence homology to the PUFA synthases. The hallmark of the enediynes, such as neocarzinostatin (Fig. 1.1), is an unsaturated core structure (called the

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enediyne ‘‘warhead’’), comprising two acetylenic groups conjugated to a double bond, all of which are housed within a 9- or 10-membered ring. Maturation of the core typically involves addition of further building blocks, including deoxy- and thio-sugars, which together fine-tune the biological activity. The precursor to the core, a polyene, is assembled by an iterative type I PKS, which contains familiar catalytic activities (KS, AT, KR, DH), but additionally, an ACP, which does not resemble other known carrier proteins, and which occupies an atypical position within the linear domain sequence. As with the fungal systems, the most significant issue concerns PKS programming, as the DH operates in all chain extension cycles except the final one. Some clusters also contain a member of a separate family of iterative type I PKS, which show organizational similarity to the PR systems of fungi. In contrast to the single-module architecture of the iterative PKS, the modular type I PKSs (usually) incorporate a distinct set or module of enzymatic domains for every catalytic cycle (Fischbach and Walsh, 2006; Staunton and Weissman, 2001, and references therein). This design strategy has resulted in some of the largest proteins in nature even though, in the vast majority of cases, the multiple modules in each system are distributed over several protein subunits. For example, MLSA1 from the mycolactone PKS, which contains eight linked modules, has a molecular mass of 1.8 MDa (Stinear et al., 2004). The modular architecture is inherently flexible, a feature that presumably justifies its high metabolic expense. Each module can be designed to select a particular building block (the biogenesis of unusual extender units is addressed in Chapter 7 of this volume) and to perform a defined extent of reductive modification, often with important stereochemical consequences. The overall number of chain extension reactions in the pathway can also be varied, as can the portfolio of post-PKS tailoring reactions. The products of modular PKSs are correspondingly diverse, although large macrocyclic lactones predominate among known structures. The prototypical modular PKS (called ‘‘DEBS’’) is responsible for biosynthesis of the macrolide erythromycin A in the soil bacterium Saccharopolyspora erythraea (Fig. 1.1). It was the first to be discovered (Corte´s et al., 1990; Donadio et al., 1991), and remains the most intensively studied, serving as the lens through which all other modular PKS are viewed (Khosla et al., 2007). Notably, there is a strict colinearity between the complement of (active) domains in the PKS and the required sequence of biosynthetic transformations, which immediately revealed the biosynthetic logic of these systems. As this ‘‘one enzyme, one function’’ organization evokes the Ford method of car manufacture, modular PKSs have been dubbed molecular ‘‘assembly lines.’’ Chapter 6 of this volume discusses the DEBS story, from the initial discovery of the gene cluster in the early 1990s to the latest insights obtained from combined genetic and molecular biological approaches.


11

In parallel, Chapter 8 in volume 458 explains how the correlation between gene sequence and product architecture has enabled the development of bioinformatics approaches to predicting polyketide structural elements from available gene sequences. Biosynthesis of the polyene antifungal drugs such as candicidin and amphotericin B (Fig. 1.1) closely resembles that of erythromycin A, although post-assembly line decoration is comparatively minimal. The defining feature of these macrolides is a chromophore formed by three to seven conjugated double bonds. Among the polyene clusters is the largest modular PKS yet discovered, with 122 domains spread over nine proteins, for a total molecular mass of 4.7 MDa (McAlpine et al., 2005). Studies to elucidate the pathways to these molecules are reviewed in Chapters 10 and 11 of this volume. Linear polyene intermediates are also the precursors for a distinct structural class of polyketides produced by modular PKS systems, the polycyclic polyethers such as monensin A (Fig. 1.1). Late-stage cyclization to form the characteristic five- and six-membered ether rings is accomplished by epoxidation, followed by a cascade of epoxide openings and ring closures. Insights into the detailed mechanisms of polyether formation have recently emerged through the cloning and sequencing of several gene clusters, including those for monensin (Leadlay et al., 2001), nigericin (Harvey et al., 2007), nanchangmycin (Sun et al., 2003), tetronomycin (Demydchuk et al., 2008), and lasalocid (Smith et al., 2008). Chapter 9 of this volume covers the latest advances in this rapidly moving area. As discussed in Chapter 12 of this volume, modular PKS machinery is also used to biosynthesize an array of exotic lipids in mycobacteria, including the dimycocerosate esters, mycolic acids (Fig. 1.1), sulfolipids and mannosyl-b-1-phosphomycoketides (Gokhale et al., 2007, and references therein). In each case, the PKS is charged with a long-chain fatty acid recruited from primary metabolism. One particularly notable system is PKS12, a polypeptide that contains two fused modules. This multienzyme self-associates five times through terminal docking domains (Broadhurst et al., 2003) to form a supramolecular complex, which then catalyzes a total of five rounds of chain extension (Chopra et al., 2008). Although monomodular PKS subunits from other systems have been shown to act repeatedly (Olano et al., 2003; He and Hertweck, 2003), this is the first example in which iterative action in a pathway is achieved through the presence of multiple copies of the same polypeptide. Remarkably, nature has fashioned a second class of modular PKS (the socalled ‘‘trans-AT’’ or ‘‘AT-less’’ PKS), an apparent example of convergent evolution (Nguyen et al., 2008). The hallmark of these systems is that one or more AT domains is supplied as a free-standing protein, instead of being integrated into each module (Piel, 2002; Cheng et al., 2003). In fact, AT-less PKS systems are formally hybrids of type I and type II architecture, as the trans-AT domains perform chain loading in iterative fashion. The resulting

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structures are also often chimeric, as many synthases incorporate machinery for chain extension with amino acids (so-called ‘‘non-ribosomal peptide synthetases [NRPSs]’’). Although the PKS domains sometimes occur in the same order as their ‘‘cis’’ counterparts, a large number of trans-AT modules exhibit conspicuous variations in domain composition, stoichiometry, and arrangement. Nonetheless, the polyketide portions of the metabolites exhibit remarkable overall structure similarity to those arising from classical cis systems. One distinctive feature of many trans-AT-derived polyketides, however, is the introduction of methyl-, or alternatively, ethyl-branching functionality at initially formed b-centers, as in the symbiont metabolite pederin (Fig. 1.1). Chapter 8 of this volume describes some representative AT-less PKSs, and details biochemical protocols to characterize essential components of these systems.

1.5. Combinatorial biosynthesis: Prospects and progress Even before the advent of the genetics era in polyketide biosynthesis, a central goal of research was to generate large collections of structural variants for evaluation as lead compounds. The discovery of the modular architecture of type I PKS immediately suggested a new strategy for polyketide drug discovery, as an adjunct to total and semisynthesis: the ‘‘mixing and matching’’ of different PKS components both within and between different synthases by genetic engineering (Weissman and Leadlay, 2005). Over the last several decades, this so-called ‘‘combinatorial biosynthesis’’ has resulted in over 200 new polyketides, but key challenges remain before the technology can be made routine. Chapter 13 of this volume presents a ‘‘how-to’’ guide for PKS engineering, including the latest tips for carrying out domain swaps and gene inactivations. The authors also provide pointers on the complementary mutasynthetic approach, in which chemically synthesized precursors are introduced into a pathway by targeted inactivation of an essential, early-stage enzyme (Kennedy, 2008; Weissman, 2007; Weist and Su¨ssmuth, 2005). One very significant impediment to genetic engineering is the enormous size of the genes encoding for modular PKS enzymes. This problem is further compounded by the lack of tools for genetic manipulation in a large number of polyketide-producing strains. A proven solution is to reconstitute biosynthesis in genetically friendlier hosts by wholesale transfer of gene sets (Wenzel and Mu¨ller, 2005b; Zhang et al., 2008), an approach that is not limited to type I PKS gene clusters. This heterologous expression technology, with an emphasis on host selection, is discussed in detail in Chapter 15 of this volume. E. coli is among the favored alternative hosts, as expression in this strain opens up the possibility of using new recombinogenic methods to manipulate gene clusters (Zhang et al., 1998, and see Chapter 7 of volume 458). Alternatively, PKS genes can be synthesized


13

from scratch, and then introduced into a strain of choice. This strategy simultaneously enables optimization of codon usage for efficient protein expression, and the introduction of specific restriction sites for convenient domain or module exchange experiments. Such de novo synthesis of polyketide pathways, and its application to the erythromycin system, is covered in detail in Chapter 14 of this volume. It is also clear that our ability to productively reconfigure PKS enzymes is intimately tied to our understanding of their function, and therefore the structural biology of PKS systems has been a central focus of research. Chapter 2 reviews methodologies and progress in the structure elucidation by NMR and X-ray crystallography of components from both type I and type II PKS, and how these efforts have informed our models for interdomain interactions in these systems. The powerful, complementary approach of using synthetic small molecules as structural and mechanistic probes is covered in Chapter 9 of volume 458. Taken together, the chapters in this volume reveal how nature exploits a common biosynthetic concept and a limited pool of simple building blocks to generate a remarkable assortment of polyketide metabolites, from linear, fatty-acid like structures, to tightly coiled aromatic molecules. However, as the recent discovery of the PUFA synthases demonstrates, our knowledge of PKS-catalyzed chemistry and architecture is likely to be incomplete, at best. Undoubtedly, the explosive growth in microbial genome data, encompassing sequences from soil, marine and symbiont metagenomes (Daniel, 2004; Taylor et al., 2007), will continue to reveal unexpected applications of PKS machineries. The real challenge will be to integrate the emerging information into coherent models for PKS operation, and to use the new insights to more efficiently engineer PKS enzymes to create novel metabolites of clinical value.

ACKNOWLEDGMENT Jim Staunton is thanked for critical reading of this manuscript.

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from sequencing of the monensin biosynthetic gene cluster. J. Ind. Microbiol. Biotechnol. 27, 360–367. Ma, Y., Smith, L. H., Cox, R. J., Beltran-Alvarez, P., Arthur, C. J., and Simpson, T. J. (2006). Catalytic relationships between type I and type II iterative polyketide synthases: The Aspergillus parasiticus norsolorinic acid synthase. ChemBioChem 7, 1951–1958. McAlpine, J. B., Bachmann, B. O., Piraee, M., Tremblay, S., Alarco, A. M., Zazopoulos, E., and Farnet, C. M. (2005). Microbial genomics as a guide to drug discovery and structural elucidation: ECO-02301, a novel antifungal agent, as an example. J. Nat. Prod. 68, 493–496. Metz, J. G., Roessler, P., Facciotti, D., Levering, C., Dittrich, F., Lassner, M., Valentine, R., Lardizabal, K., Domergue, F., Yamada, A., Yazawa, K., Knauf, V., et al. (2001). Production of polyunsaturated fatty acids by polyketide synthases in both prokaryotes and eukaryotes. Science 293, 290–293. Moss, S. J., Martin, C. J., and Wilkinson, B. (2004). Loss of co-linearity by modular polyketide synthases: A mechanism for the evolution of chemical diversity. Nat. Prod. Rep. 21, 575–593. Murata, M., Naoki, H., Matsunaga, S., Satake, M., and Yasumoto, T. (1994). Structure and partial stereochemical assignments for maitotoxin, the most toxic and largest natural non-biopolymer. J. Am. Chem. Soc. 116, 7098–7107. Napier, J. A. (2002). Plumbing the depths of PUFA biosynthesis: A novel polyketide synthase-like pathway from marine organisms. Trends Plant Sci. 7, 51–54. Nguyen, T., Ishida, K., Jenke-Kodama, H., Dittmann, E., Gurgui, C., Hochmuth, T., Taudien, S., Platzer, M., Hertweck, C., and Piel, J. (2008). Exploiting the mosaic structure of trans-acyltransferase polyketide synthases for natural product discovery and pathway dissection. Nat. Biotechnol. 26, 225–233. Nichols, D., Bowman, J., Sanderson, K., Nichols, C. M., Lewis, T., McMeekin, T., and Nichols, P. D. (1999). Developments with antarctic microorganisms: Culture collections, bioactivity screening, taxonomy, PUFA production and cold-adapted enzymes. Curr. Opin. Biotechnol. 10, 240–246. Olano, C., Wilkinson, B., Moss, S. J., Branã, A. F., Meńdez, C., Leadlay, P. F., and Salas, J. A. (2003). Evidence from engineered gene fusions for the repeated use of a module in a modular polyketide synthase. Chem. Commun. 2780–2782. Piel, J. (2002). A polyketide synthase-peptide synthetase gene cluster from an uncultured bacterial symbiont of Paederus beetles. Proc. Natl. Acad. Sci. USA 99, 14002–14007. Rix, U., Fischer, C., Remsing, L. L., and Rohr, J. (2002). Modification of post-PKS tailoring steps through combinatorial biosynthesis. Nat. Prod. Rep. 19, 542–580. Roberts, W. C. (1996). The underused miracle drugs: The statin drugs are to atherosclerosis what penicillin was to infectious disease. Am. J. Cardiol. 78, 377–378. Shen, B. (2003). Polyketide biosynthesis beyond the type I, II and III polyketide synthase paradigms. Curr. Opin. Chem. Biol. 7, 285–295. Shepherd, J. (2006). Who should receive a statin these days? Lessons from recent clinical trials. J. Intern. Med. 260, 305–319. Smith, L., Hong, H., Spencer, J. B., and Leadlay, P. F. (2008). Analysis of specific mutants in the lasalocid gene cluster: Evidence for enzymatic catalysis of a disfavoured polyether ring closure. ChemBioChem 9, 2967–2975. Staunton, J., and Weissman, K. J. (2001). Polyketide biosynthesis: A millennium review. Nat. Prod. Rep. 18, 380–416. Stinear, T. P., Mve-Obiang, A., Small, P. L., Frigui, W., Pryor, M. J., Brosch, R., Jenkin, G. A., Johnson, P. D., Davies, J. K., Lee, R. E., Adusumilli, S., Garnier, T., et al. (2004). Giant plasmid-encoded polyketide synthases produce the macrolide toxin of Mycobacterium ulcerans. Proc. Natl. Acad. Sci. USA 101, 1345–1349.

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Sun, Y., Zhou, X., Dong, H., Tu, G., Wang, M., Wang, B., and Deng, Z. (2003). A complete gene cluster from Streptomyces nanchangensis NS3226 encoding biosynthesis of the polyether ionophore nanchangmycin. Chem. Biol. 10, 431–441. Taylor, M. W., Radax, R., Steger, D., and Wagner, M. (2007). Sponge-associated microorganisms: Evolution, ecology, and biotechnological potential. Microbiol. Mol. Biol. Rev. 71, 295–347. Van Lanen, S. G., and Shen, B. (2008). Biosynthesis of enediyne antitumor antibiotics. Curr. Top. Med. Chem. 8, 448–459. Wallis, J. G., Watts, J. L., and Browse, J. (2002). Polyunsaturated fatty acid synthesis: What will they think of next? Trends Biochem. Sci. 27, 467–473. Washington, J. A., and Wilson, W. R. (1985). Erythromycin: A microbial and clinical perspective after 30 years of clinical use (1). Mayo Clin. Proc. 60, 189–203. (http://www. emedexpert.com/facts/azithromycin-facts.shtml; http://www.wipo.int/sme/en/case_ studies/pliva.htm). Weissman, K. J. (2007). Mutasynthesis—uniting chemistry and genetics for drug discovery. Trends Biotechnol. 25, 139–142. Weissman, K. J., and Leadlay, P. F. (2005). Combinatorial biosynthesis of reduced polyketides. Nat. Rev. Microbiol. 3, 925–936. Weist, S., and Su¨ssmuth, R. D. (2005). Mutational biosynthesis—a tool for the generation of structural diversity in the biosynthesis of antibiotics. Appl. Microbiol. Biotechnol. 68, 141–150. Wenzel, S. C., and Mu¨ller, R. (2005a). Formation of novel secondary metabolites by bacterial multimodular assembly lines: Deviations from text book biosynthetic logic. Curr. Opin. Chem. Biol. 9, 447–458. Wenzel, S. C., and Mu¨ller, R. (2005b). Recent developments towards the heterologous expression of complex bacterial natural product biosynthetic pathways. Curr. Opin. Biotechnol. 16, 594–606. Zhang, H. R., Wang, Y., and Pfeifer, B. A. (2008). Bacterial hosts for natural product production. Mol. Pharm. 5, 212–225. Zhang, Y., Buchholz, F., Muyrers, J. P., and Stewart, F. A. (1998). A new logic for DNA engineering using recombination in Escherichia coli. Nat. Genet. 20, 123–128.

C H A P T E R

T W O

Structural Enzymology of Polyketide Synthases Shiou-Chuan (Sheryl) Tsai* and Brian Douglas Ames† Contents 18 19 21 22 22 24 27 30 32 36 37 39 40 40

1. 2. 3. 4.

Introduction Fatty Acid Synthase Different Types of PKS and a Summary of Structural Work Structural Enzymology of Individual Domains 4.1. The acyltransferase 4.2. The ketosynthase 4.3. The ketoreductase 4.4. The dehydratase of modular type I PKS 4.5. The aromatase/cyclase (ARO/CYC) of type II PKS 4.6. Acyl carrier protein 4.7. Thioesterase 5. Summary and Future Prospects Acknowledgments References

Abstract This chapter describes structural and associated enzymological studies of polyketide synthases, including isolated single domains and multidomain fragments. The sequence–structure–function relationship of polyketide biosynthesis, compared with homologous fatty acid synthesis, is discussed in detail. Structural enzymology sheds light on sequence and structural motifs that are important for the precise timing, substrate recognition, enzyme catalysis, and protein–protein interactions leading to the extraordinary structural diversity of naturally occurring polyketides.

* {

Department of Molecular Biology and Biochemistry, Department of Chemistry, Department of Pharmaceutical Sciences, University of California, Irvine, California, USA Department of Molecular Biology and Biochemistry, Department of Chemistry, University of California, Irvine, California, USA

Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04602-3

#


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Shiou-Chuan (Sheryl) Tsai and Brian Douglas Ames

1. Introduction The past decade has witnessed significant advances in PKS structural biology for different types of PKSs that help visualize polyketide biosynthesis at all stages, including chain initiation, elongation, reduction, cyclization, and chain termination. Using x-ray crystallography or NMR, these studies help correlate PKS three-dimensional structures with substrate specificity and also help in elucidating the sequence–function–structure relationships that predict product outcome. With the recent publication of the striking porcine FAS crystal structure, we summarize advances in PKS structural biology during the past decade, and compare different types of FASs and PKSs, as well as their individual enzyme domains. The major methods used to determine PKS protein structures are NMR and x-ray crystallography. With the current advances in NMR instrumentation, software, and pulse sequence development, the upper limit of protein molecular weight for a given target can be as high as 80 kDa (Redfield, 2004; Zeeb and Balbach, 2004). Various pulse sequences offer a further powerful probe for protein conformational change, as well as residues that are important for protein–protein interaction and protein-ligand binding (Redfield, 2004; Zeeb and Balbach, 2004). The application of these NMR methodologies to PKS enzymology is presented in section 4 on the structure of the acyl-carrier protein (ACP). The general methodology for x-ray crystallography includes protein expression/purification, crystallization, crystal harvesting, x-ray diffraction, data processing, structure solution, model refinement and model validation. While mammalian fatty acid synthase (FAS) architecture can be visualized by electron microscopy (EM) (Asturias et al., 2005), PKS flexibility has resulted in highly variable conformations as detected by electron microscopy (Grant Jensen, personal communication), and such flexibility may also contribute to the relative difficulty in obtaining PKS megasynthase crystals (as compared to crystallizing the mammalian FASs). Nevertheless, the majority of single and didomain PKS fragments can be cloned into standard expression vectors (such as pET vectors), expressed in large quantity (up to 200 mg/l culture), and purified to more than 95% purity using Ni-affinity chromatography. The pure single and didomain PKS fragments can then be crystallized using standard vapor diffusion methods (Derewenda, 2004; McPherson, 2004; Rupp and Wang, 2004), followed by crystal harvest in cryoprotectant and cooling in liquid nitrogen (Pflugrath, 2004). Robotic automation of crystallization has greatly accelerated the discovery of initial crystal leads (Bard et al., 2004). Due to the high structural homology to FAS domains, many PKS domains have been solved by molecular replacement, using programs such as CNS (Brunger et al., 1998) or the CCP4 suite (Xx, 1994), although heavyatom methods are also applied using programs such as SOLVE (Terwilliger, 2004). Protein models are then built and refined by programs such as Coot

Structural Enzymology of Polyketide Synthases

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(Emsley and Cowtan, 2004), further refined by programs such as CNS (Brunger et al., 1998) or the CCP4 suite (Xx, 1994), and verified by programs such as PROCHECK (Laskowski et al., 1993). As discussed in detail in Section 4, the conformational flexibility of many PKS domains renders the binding of enzyme inhibitors or substrates very useful to stabilize protein conformation for crystallization and improve diffraction data quality. However, the reactivity of these PKS enzyme domains sometimes also results in difficulty in the detection of substrate electron density maps. The above issues in the methodology of PKS structural enzymology, as well as the application and subsequent outcome of these methodologies, are discussed in details in Sections 3 and 4.

2. Fatty Acid Synthase The fatty acid synthase (FAS) is a multidomain protein complex (Maier et al., 2006) consisting of seven conserved protein domains (MAT, KS, KR, DH, ER, ACP and TE) that catalyze more than 50 reactions en route to the final fatty acid product. FASs can be classified as type I or type II (Fig. 2.1A). The crystal structures of all type II FAS domains have been solved, including the type II KS, MAT, KR, ER, DH, ACP, and TE domains (Leesong et al., 1996; Olsen et al., 1999; Price et al., 2001; Serre et al., 1995), and White et al. have provided an excellent review of type II FAS structural biology (White et al., 2005). The crystal structure of a type I FAS TE domain, as well as the NMR structure of a type I FAS ACP domain, have also been reported (Chakravarty et al., 2004) and reviewed (Smith and Tsai, 2007). The recently ˚ , has greatly published full-length mammalian type I FAS, solved to 3.2 A expanded our knowledge about the complicated domain–domain interactions in the megasynthase (Maier et al., 2008) (Fig. 2.1B, C), showing a homodimer that confirms the head-to-head model based on electron microscopy (Asturias et al., 2005) and biochemical results ( Joshi et al., 1997, 1998). The porcine FAS is separated into two portions: the lower condensing portion which contains the KS and MAT domains, and the upper chain-modifying portion which contains the DH, ER, and KR domains (Fig. 2.1B, C). Two additional nonenzymatic domains, termed cMe (an inactive methyltransferase) and cKR (a truncated KR), lie at the periphery of the upper portion (Fig. 2.1B, C).The central core of the X-shape architecture of type I FAS consists of KS, DH, and ER domains from both monomers, with an extensive dimer interface between KS, DH, and ER domains. The dimer is held together by KS–KS, DH–DH and KS–DH interactions. The head-to-head arrangement also implies that the ACP in either monomer (A or B) can interact with active sites of both monomers, consistent with previous biochemical studies ( Joshi et al., 1997; Witkowski et al., 2004). The DH–KS interaction connects the top and bottom of the

A Type I FAS

NH+3

Type II FAS

AT

KS

DH

AT

KS

ER

DH

ER

KR

KR

TE

ACP

ACP

COO−

TE

ER

B ER⬘

KR

KR⬘

ΨKR

ΨKR⬘

C ER KR

DH1

DH⬘ ΨME⬘

ACP anchor

Central connection

ΨKR

DH2

DH Mo

dif

Con

yin

g

ΨME

ΨME ACP

den

sing

TE C

KS LD

LD⬘ KS⬘ KS MAT⬘

LD MAT

N

MAT

˚ Figure 2.1 (A) Domain organization of type I and type II FAS. (B) Cartoon ribbon representation and (C) domain illustration of the 3.2-A porcine type I FAS structure.


21

X-shaped architecture, and strongly suggests that the DH fold is important in maintaining FAS architecture. The AT and KR domains extend from the bottom and top of the protein, respectively, to form two asymmetric reaction chambers, one on either side of the central KS–DH–ER core. The flexible linker regions between KS–AT, AT–DH, DH–ER and ER–KR (Fig. 2.1C) facilitate the opening and closing of each reaction chamber, thus allowing the ACP-bound fatty acyl intermediate access to the active-site of each domain. Although there is no solid proof, the type II FAS is proposed to adopt a similar X-shaped architecture (Smith and Tsai, 2007) due to the high degree of conservation between type I and II FAS domains.

3. Different Types of PKS and a Summary of Structural Work Based on domain architecture, there are at least four distinct PKS types: modular type I, iterative type I, type II, and type III PKSs (Shen, 2003). Extensive progress has been made on the structural biology of DEBS (the erythromycin PKS) (Khosla et al., 2007), including crystal structures of KS3–AT3 (Tang et al., 2007), KS5–AT5 (Tang et al., 2006), KR1 (Keatinge-Clay and Stroud, 2006), DH4 (Keatinge-Clay, 2008), and TE (Tsai et al., 2001), and an NMR structure of ACP2 (Alekseyev et al., 2007) (the number following the domain name abbreviation indicates the module number), as well as KR1 of the tylosin PKS (Keatinge-Clay, 2007) and the TE domain from the pikromycin PKS (PIKS) (Akey et al., 2006; Giraldes et al., 2006; Pan et al., 2002; Tsai et al., 2002). An NMR structure of the intermodular linker region in DEBS has also been reported (Weissman, 2006). For the type II PKS, extensive progress has been made on the structural biology of Streptomyces coelicolor MAT (Keatinge-Clay et al., 2003), actinorhodin (act) KS/CLF (Keatinge-Clay et al., 2004), act KR (Hadfield et al., 2004; Korman et al., 2004, 2008), the R1128 priming ketosynthase ZhuH (Pan, et al., 2002; Witkowski et al., 2004), and the NMR structure of holo and apo act ACP (Crump et al., 1997; Evans et al., 2008), frenolicin ( fren) ACP (Li et al., 2003), and oxytetracycline (otc) ACP (Findlow et al., 2003). The crystal structures of the tetracenomycin (tcm) ARO/CYC, the whiE ARO/CYC, and the ZhuI ARO/CYC have been solved (Ames et al., 2008). Three fourth-ring cyclase structures have also been reported that are not included in this chapter (Kallio et al., 2006; Sultana et al., 2004; Thompson et al., 2004). For iterative type I PKSs, the unpublished crystal structures of the PksA PT and TE domains are discussed briefly. The linker regions in type I PKSs have been reviewed elsewhere (Smith and Tsai, 2007; Weissman, 2006).

22


4. Structural Enzymology of Individual Domains 4.1. The acyltransferase The crystal structures of DEBS AT3 and AT5, S. coelicolor MAT, E. coli MAT (FabD) and the porcine FAS AT domain have been reported (Keatinge-Clay et al., 2003; Serre et al., 1995; Tang et al., 2006, 2007). All ˚ . The AT five structures are highly similar, with an RMSD of 1.3 to 1.9 A structure has two domains: the larger core subdomain is similar to an a/bhydrolase fold, with a parallel b-sheet flanked on two sides by a-helices (Fig. 2.2A in grey); the smaller subdomain, an insertion typically between residues 130 to 200, has a ferredoxin fold that consists of a four-stranded antiparallel b-sheet capped by two helices (Fig. 2.2A in black). The active site lies in a cleft formed between the two subdomains. Protein–protein docking simulations were conducted using the DEBS AT3 and ACP3 homology models (Tang et al., 2007), although no apparent solution was obtained. The most important structural difference between the type I modular PKS AT domains and other MAT domains is that DEBS AT3 and AT5 have a much longer C-terminal helix (residues 857 to 867), presumably important for protein–protein interactions between AT and the KS-AT linker (Tang et al., 2006, 2007). Based on enzymological studies of the type II FAS MAT domain, a ping-pong bi bi mechanism is proposed for both type I and type II ATs of FAS and PKS that involves the active site Ser and His (Fig. 2.2B) (Ruch and Vagelos, 1973), while the 3-carboxylate of malonyl- or methylmalonyl-CoA can form charge–charge interactions with the side-chain of a highly conserved Arg (Keatinge-Clay et al., 2003; Rangan and Smith, 1997; Serre et al., 1995). In the S. coelicolor MAT structure, the backbone amides of Gln9 and Val98 were identified as the oxyanion hole (Keatinge-Clay et al., 2003). Remarkably, for both type I and type II AT domains, the acyl-enzyme complex is stable enough for detection (Dreier et al., 2001; Lau et al., 2000; Liou et al., 2003; Szafranska et al., 2002). The deacylation of acyl-enzyme occurs only in the presence of specific thiol nucleophiles such as phosphopantetheinylated ACP. This is a unique behavior among structurally related enzymes with an a/b hydrolase fold (Serre et al., 1995). In addition, based on the crystal structures of all five AT domains, the hydrophobic nature of the substrate-binding pocket should presumably discourage water binding in the cleft. For type I modular PKSs, the loading AT domain is promiscuous and can accept acetyl-, propionyl-, isopropionyl-, isobutyryl-, crotonyl-, phenylacetyl-, hydroxybutyryl-, and isopentyl-CoA both in vivo and in vitro (Del Vecchio et al., 2003; Hong et al., 2005; Lau et al., 1999, 2000; Liou and Khosla, 2003; Liou et al., 2003). On the other hand, the AT domains in the extender

23


A 1 A 4

3

2

B B

His

H N N HO

His R O S

His

His

O Ser

H N

His O

Ser

O

NH R

O− NH Oxyanion S CoA

NH R ACP-SH

H N

NH R NH Oxyanion

CoA

Ser

H N

NH Oxyanion R = rest of acyl chain

O

CoA

O Ser

H N NH O Ser

R R

O− NH Oxyanion S ACP

O S ACP

Figure 2.2 (A) Overall fold of the AT domain (large domain in grey, small domain in black), and the four substrate motifs (see text for details) in red, green, purple, and blue, respectively. (B) Proposed mechanism for the AT domain.

modules (such as DEBS modules 1 to 6) are highly specific toward (2S)-methylmalonyl-CoA and the (2S)-methylmalonyl thioester of N-acetylcysteamine (NAC) (Lau et al., 1999; Liou and Khosla, 2003; Marsden et al., 1994). Therefore, the extending AT in DEBS serves as an important gatekeeper in macrolide biosynthesis (Khosla et al., 1999). Past biochemical and structural work identified four motifs to explain the observed AT specificity (Fig. 2.2A): (1) the ‘‘RVDVVQ’’ motif lies 30 residues upstream of the active-site Ser (Haydock et al., 1995; Yadav et al., 2003); (2) the GHSXG motif around the catalytic Ser (Haydock et al., 1995; Yadav et al., 2003); (3) the YASH motif 100 residues downstream of the active-site Ser (Haydock et al., 1995; Reeves et al., 2001), which based on a

24


systematic mutational analysis of DEBS AT domains is the dominant substrate specificity motif in type I modular PKSs among motifs 1, 2, and 3 (Reeves et al., 2001); and (4) the C-terminal region shown to be important for substrate specificity from domain swapping experiments (Lau et al., 1999). A detailed review of the above four motifs can be found in reference (Smith and Tsai, 2007). The recent 3.2-A˚ porcine FAS structure further demonstrates that F682 (part of Motif 1) and F553 (part of Motif 3) form a hydrophobic cavity, which may allow M499 to flip in and out to accommodate both methylmalonyl and malonyl moieties. In conclusion, the specificity between malonyl- versus methylmalonyl-CoA (or methylmalonyl- vs. propionyl-CoA) is likely to be a combinatorial result of different structural elements that interact throughout the entire protein fold, rather than an influence of a limited number of residues.

4.2. The ketosynthase The crystal structures of DEBS ketosynthase (KS) 3 (Tang et al., 2007) and KS5 (Tang et al., 2006), the actinorhodin KS/CLF (act KS/CLF) (Keatinge-Clay et al., 2004) and the R1128 priming KS (ZhuH) (Pan et al., 2002) will be compared with those of the type III PKS (Austin and Noel, 2003), type I (the porcine FAS) (Maier et al., 2008), and type II (the E. coli FabH, FabB, and FabF) KS domains (Olsen et al., 1999; Price et al., 2003; Qiu et al., 2005). All KS crystal structures from FASs and PKSs reveal a highly similar thiolase fold (Austin and Noel, 2003), consisting of two copies of a-b-a-b-a folds that form a five-layered core (2a-5b-2a-5b-2a): three layers of a-helices interspersed by two layers of b-sheet, with extensive connecting loops. A pseudotwofold axis lies between Na3 and Ca3 parallel to the dimer axis. The KSs can be divided into three subfamilies: (1) KAS I and II, including FabB, FabF, DEBS KS3 and KS5, the porcine FAS KS domain, and the act KS/CLF (Keatinge-Clay et al., 2004; Maier et al., 2008; Olsen et al., 1999; Price et al., 2003; Tang et al., 2006, 2007); (2) KAS III and the CHS-like type III PKS enzymes, including the priming KS ZhuH (Pan et al., 2002) from the R1128 PKS; (3) the biosynthetic and degradative thiolases. All three subfamilies conserve the core structural features, the extensive dimer interface, and the location of the active-site residues. There is also absolute conservation of the active-site Cys for covalent attachment of substrates and intermediates. However, there are major structural differences among the three subfamilies, mainly concerning the extent and structure of the loops on the opposite side of the substrate-binding pocket. These loops affect the position and identity of key catalytic residues (except the universally conserved Cys) as well as the different substrate chain-length specificities for CoA-linked or ACP-linked thioesters (Austin and Noel, 2003). The KSs are dimers with an extensive dimer interface stabilized by a pair of hydrogen-bonded, antiparallel b-strands, which creates a 14-stranded b-sheet spanning both monomers. In the DEBS KS5 structure, there is a long helix at the N-terminal that may facilitate KS


25

dimerization and serve as a docking point for the upstream C-terminal docking domain of the previous module 4 (Tang et al., 2006). In the type I FAS and PKS, KS deletion experiments show that the dimeric nature of the KS domains is key in facilitating dimerization of the entire megasynthase (Witkowski et al., 2004). Similarly to the well-studied FabB and FabF KSs of E. coli, the extending KSs, including DEBS KS3 and KS5 and act KS/CLF, employ a Cys-His-His triad at the active center. The active-site triad and oxyanion hole of DEBS KS3 and KS5, act KS/CLF and porcine FAS KS domains can be overlapped perfectly, suggesting a similar catalytic mechanism (Witkowski et al., 2002; Zhang et al., 2006), initiated by the docking of acyl-ACP to an electropositive patch of KS. The proposed mechanism is similar between the priming (ZhuH) and extending (DEBS KS3 and KS5 and act KS/CLF) KSs, except that the catalytic triad of a priming KS consists of His-Asn-Cys. Furthermore, acyl-CoA, rather than acyl-ACP, first binds to ZhuH. The Asn versus His difference between the priming and elongation KSs helps explain why inhibitors such as cerulenin and thiolactomycin preferentially bind the elongation KSs (White et al., 2005). Despite a similar thiolase fold and enzyme mechanism, the specificity of type I and II FAS and PKS KSs varies significantly. The type I FAS KS domains are highly specific toward saturated acyl chains (Witkowski et al., 2002). In contrast, type I modular KS domains, such as the six KS domains in DEBS, have a wide range of substrate specificities that vary in length from diketide to decaketide, although some PKS KS domains appear to possess some specificity with regard to different b-carbon status (Khosla et al., 1999). The type II systems possess highly specific chain-length control that is pathway-specific for the priming and extending KSs. Examples include the C2–C4 priming preference of ZhuH (Pan et al., 2002; Qiu et al., 2005), or the act, tcm, and whiE KS/CLF which extend the polyketide chain to 16, 20, and 24 carbons, respectively (Tang et al., 2003). The observed substrate specificity of each KS domain can be explained by the size and shape of the KS substrate-binding channel, which can be divided into two halves, corresponding to the substrate-binding and PPT-binding regions (Fig. 2.3). The PPT-binding region stretches from the enzyme surface to the active-site Cys and this region is relatively well conserved, reflecting its universal role in binding the PPT moiety. In contrast, the acylbinding region varies significantly. While the binding pockets of FAS KS domains are hydrophobic and promote fatty acyl binding (Maier et al., 2008; Olsen et al., 1999; Price et al., 2003; Qiu et al., 2005), the acyl-binding pockets of PKS KS domains (such as DEBS KS3 and KS5, act KS/CLF and ZhuH) are amphipathic and allow hydrogen-bonding interactions with the carbonyl groups of the growing polyketide chain (Keatinge-Clay et al., 2004; Pan et al., 2002; Tang et al., 2006, 2007). The FAS KS domains have a hydrophobic, narrow pocket of a suitable size to specifically

A

B

D

E

C

F

G

Figure 2.3 The substrate binding channel of (A) the priming KS ZhuH, (B) the E. coli KAS I, (C) actinorhodin KS/CLF, (D) chalcone synthase, (E) porcine FAS KS domain, (F) DEBS KS5. The active site Cys is shown in spheres, and (G) The cyclization chamber of chalcone synthase (in green) compared to the linear polyketide-extending chamber in the act KS/CLF (in purple).

27


accommodate their corresponding fatty acyl substrates (Fig. 2.3B) (Maier et al., 2008; Olsen et al., 1999; Price et al., 2003; Qiu et al., 2005), whereas the substrate pockets in DEBS KS3 and KS5 are much wider (Fig. 2.3F) (Tang et al., 2006, 2007), consistent with the substrate tolerance reported for type I modular PKS KS domains. In contrast, both priming and extending KSs of the type II PKS are more substrate-specific, reflected by the narrower acyl-binding pocket, similar to those in FAS KS domains. Significantly, mutations of four residues that define the bottom of the acyl pocket in act and tcm KS/CLF confirmed that the pocket size and shape indeed control polyketide-product chain length (Tang et al., 2003), with mutations in CLF being sufficient to alter chain length.

4.3. The ketoreductase Three ketoreductase (KR) crystal structures have been reported: for type I modular PKSs, crystal structures of the DEBS KR1 (EryKR1) (Keatinge-Clay and Stroud, 2006) and KR1 of tylosin PKS (TylKR1) (Keatinge-Clay, 2007) have been solved (Fig. 2.4A-B) and for type II PKS, the actinorhodin KR (act KR) crystal structure has been reported (Hadfield et al., 2004; Korman et al., 2004, 2008) (Fig. 2.4D). EryKR1 and TylKR1 reduce a diketide substrate C¼O to C-OH with ‘‘2R, 3R’’ and ‘‘2R, 3S’’ stereochemistry, respectively, so these two KRs choose opposite diketide epimers (at the 2 position) for the A

D

LID

B

LID

C

E

Figure 2.4 Crystal structures of (A) EryKR1, (B) TylKR1, (C) porcine FAS KR, (D) act KR dimer, (E) the act KR tetramer, colored as in (C), NADP in spheres. A to D are in the same orientation.

28


reduction reaction. In contrast, the actinorhodin KR (act KR) specifically reduces the C9 carbonyl group of a 16-carbon (octaketide) preassembled polyketide chain, which folds into the C7–C12 first-ring cyclized shuntproduct mutactin when expressed without downstream enzymes. The ketoreduction catalyzed by act KR, as well as by other type II PKS KR domains, is chemically identical to the corresponding fatty acid ketoreduction, yet with very different regio-specificities (O’Hagan, 1993). In both FASs and PKSs, the type I KR has two domains with the same protein fold: the catalytic subdomain and a truncated, noncatalytic structural subdomain. Both EryKR1 and TylKR1 are monomeric in solution and in crystal structure (Keatinge-Clay, 2007; Keatinge-Clay and Stroud, 2006). In contrast, the type II KR exists as a tetramer (Fig. 2.4) (Hadfield et al., 2004; Korman et al., 2004), and each monomer contains a single domain. Each domain (or each subdomain in type I KR) contains a short-chain dehydrogenase/reductase (SDR) fold consisting of a highly conserved Rossmann fold with two right-handed a-b-a-b-a motifs connected by a3, and the core region consists of a seven-stranded b-sheet flanked by a-helices (Fig. 2.4). The cofactor NADPH is bound at the junction of two a-b-a-b-a motifs in a highly conserved groove characteristic of the Rossmann fold (Persson et al., 2003). The polyketide substrate-binding pocket consists of a large cleft formed by helices a6-a7 and the loops between a4 and a5 (Fig. 2.4). The catalytic subdomains of EryKR1 and TylKR1, as well as act KR, have the typical SDR motifs, such as the TGxxxGxG motif (residues 2 to 19), the D63 and NNAG motifs (residues 89 to 92), the active-site tetrad Asn–Ser–Lys–Tyr, and the PG motif (residues187 to 188) (Persson et al., 2003). The biggest difference between the type I and II PKS KRs is a long insertion between helices 6 and 7 for act KR, and this may account for the different substrate specificities of type I and II PKS KRs. The monomeric type I KR orients its two subdomains (Fig. 2.4A-B) very similarly to a type II KR dimer (Fig. 2.4D), with extensive, mainly hydrophilic, interactions. The structural subdomains in EryKR1 and TylKR1 lack a cofactor-binding motif and the substrate-binding portion, thus rendering the structural subdomain inactive (Keatinge-Clay, 2007; KeatingeClay and Stroud, 2006). The type I KRs have additional b1-b8 and a4-aF interactions bridging the structural and catalytic subdomains and stabilizing the pseudodimeric KR protein fold. The act KR active-site tetrad consists of N114–S144–Y157–K161 (42 to 44). In contrast, the Asn and Lys positions are switched in EryKR1 and TylKR1 (K1776–S1800–Y1813–N1817) (Keatinge-Clay, 2007; KeatingeClay and Stroud, 2006). The KR tetrad lies near the nicotinamide ring of NADPH, where Tyr and Lys form hydrogen bonds with the NADPH ribose and nicotinamide ring. In act KR, four crystalline water molecules form extensive hydrogen bonds with N114 and K161 (Fig. 2.5A). These waters form a proton-relay network that is very similar to the one observed

29


A

K161

Y157

N114

S144

O

O

H

H O

H

H O

H

⊕ H N H

H

H O

H

HO

H2

O3 O

H

H

O

O H

1 NADPH

N

H

O B

NADPH

Monomer B closed form

NADPH

Monomer A open form

NADPH

7.9 Å

Figure 2.5 (A) Proposed proton-relay mechanism for act KR. (B) The asymmetric unit contains monomers A (middle panel) and B (left panel) in open and closed conformations, respectively. Surface potentials are colored from negative (red) to positive (blue). When the open and closed conformations are overlapped (right panel, open in yellow, closed in purple), the major conformational change is in the flexible a6–a7 region.

in E. coli FabG-NADPþ (Price et al., 2004), leading to the hypothesis that the water-relay mechanism for FabG may also be applicable to act KR (Fig. 2.5A). In vitro assays indicate that type II PKS KRs have a different substrate specificity from that of FAS (Dutler et al., 1971; Joshi and Smith, 1993) and modular type I PKS KRs (Ostergaard et al., 2002), both of which can reduce linear and monocyclic ketones. The strong preference of act KR toward bicyclic polyketides supports the conclusion that its natural substrate is a cyclic polyketide, and detailed kinetic analysis showed that act KR proceeds through an ordered bi bi mechanism (Korman et al., 2008), in which the cofactor NADPH binds KR prior to the substrate

30


trans-1-decalone. The above results imply that the first ring is cyclized prior to ketoreduction, thus sterically constraining the ketoreduction and leading to a highly specific C9-reduction. The inhibitor emodin-bound act KR structure also revealed that act KR can exist with at least two different conformations (Korman et al., 2008): open and closed forms that differ in the 10-residue loop region (residues 199 to 209) between helices 6 and 7 (Fig. 2.5B). EryKR1 and TylKR1 also show similar conformational changes and loop movement in this region (Keatinge-Clay, 2007; Keatinge-Clay and Stroud, 2006), which may reflect different binding motifs during ketoreduction (Keatinge-Clay, 2007; Korman et al., 2008). The stereoselective signature motifs for the modular PKS KRs were previously proposed to be ‘‘LDD’’ and PxxxN (Caffrey, 2003; Reid et al., 2003), and the presence or absence of these motifs produce the 3R or 3S stereomer, respectively. The EryKR1 crystal structure shows that the 93–95 ‘‘LDD’’ motif lies in a loop adjacent to the active site (Keatinge-Clay and Stroud, 2006), and KR1 mutation indeed resulted in a switch of alcohol stereochemistry (Baerga-Ortiz et al., 2006; O’Hare et al., 2006). The 2-position stereochemistry is also affected by KR and its upstream KS domain. Based on extensive bioinformatic analysis guided by the crystal structures of EryKR1 and TylKR1, Keatinge-Clay categorized the type I KRs into six types to explain their observed stereochemistry at the 2- and 3-positions and developed a protocol to assign substituent stereochemistry accordingly (Keatinge-Clay, 2007). Similar studies on type II KRs (Korman et al., 2008) support the hypothesis that type II KR substrate specificity is defined by a combination of enzyme conformation, specific molecular interactions between the substrate and active-site residues, as well as substrate and protein flexibility due to the dynamic nature of the binding cleft.

4.4. The dehydratase of modular type I PKS DEBS DH4 catalyzes dehydration of a 2R-methyl-3R-OH pentaketide to afford a trans double bond (Keatinge-Clay, 2008). The recent report of the 1.8-A˚ DEBS DH4 crystal structure, combined with the porcine FAS structure, reveals that the DH domain in type I PKSs and FASs consists of two subdomains with limited sequence homology, yet each subdomain consists of the ‘‘hot-dog-in-a-bun’’ fold (Dillon and Bateman, 2004). The double hot-dog (DHD) fold exhibited by type I FAS and PKS DHs is similar to that of the dimeric type II bacterial DHs (such as E. coli FabA and FabZ ( Kimber et al., 2004; Leesong et al., 1996)), consisting of one hot-dog fold per monomer (Fig. 2.6A-B). The E. coli and human TE II are also reported to contain the DHD fold (Li et al., 2000). In each hot-dog subdomain, the long central helix—the hot-dog—is packed against a seven-stranded antiparallel b-sheet that forms the bun. In both the porcine FAS and DEBS DH4 structures, the two hot-dog subdomains of DH interact extensively to

A

B

C

H44

H44

D

N H

O

H N

H

O

D206 Q210

O

R1 H

H R2

O H O H H N

S55

+

N H

N

O

L51

L51

S

ACP

O

“Trans-” dehydration

H N

H

O

O− H

D206 Q210

R1

N H

O N H

N H R 2 S H

H O Wat1 H N

ACP

O

S55

Figure 2.6 Comparison of the double hot dog–fold and interior pocket of (A) DEBS DH4, (B) porcine FAS DH domain, and (C) PKSA PT domain. The active-site His and Asp are shown in sphere and sticks in upper and lower panels, respectively. The N- and C-terminal subdomains in each model are shown in darker and lighter shades, respectively. (D) Proposed mechanism for DH4.

32


form a 14-strand b-sheet with additional interactions between the hot-dog helices from each subdomain. In DH4, the catalytic H44 lies in the first subdomain as part of the HXXXGXXXXP motif (Joshi and Smith, 1993), where the conserved Gly is necessary to make a turn that enables van der Waals interactions between H44 and P53. The catalytic D206 within the DXXX(Q/H) motif lies in the second subdomain and hydrogen-bonds with the side chain of Q210 (itself anchored to Y158). The organization and interaction of catalytic residues described for DH4 are conserved in the porcine FAS DH. Additionally, both the ‘‘GYXYGPXF’’ and ‘‘LPFXW’’ motifs are highly conserved and help define the substrate pocket. The reaction catalyzed by type I and II FAS DHs is freely reversible with equilibrium favoring hydration (Heath and Rock, 1995; Witkowski et al., 2004). In type I PKS DHs, the equilibrium may also lean toward the hydrated polyketide, and a downstream KS or TE may pull the reaction forward toward the dehydrated polyketide (Tang et al., 1998; Wu et al., 2005). However, there is no obvious sequence motif associated with DHs that dehydrate substrates with or without a-substituents, indicating that the a-substituents may not be recognition factors for the PKS DHs. A catalytic mechanism was proposed for DH4 (Fig. 2.6D) in which H44 serves as the active-site base to deprotonate the a-carbon, while the b-hydroxyl group may be polarized by the helix-1 dipole, facilitating water elimination. The DH4 structure shows that the stereochemistry of the b-hydroxyl group in an incoming polyketide substrate may be the primary factor that determines if a cis or trans double bond is produced by the DH domain. For example, when an A-type KR provides DH with the 3S-OH substrate, the 3S-OH is hydrogen-bonded to D206 and the polyketide chain must rotate 120 degrees about the Ca–Cb bond. Thus, an elimination results in the formation of a cis double bond, as in the phoslactomycin DH1 and DH2 (Palaniappan et al., 2008) or rifamycin DH10 (Tang et al., 1998). Sequence comparison indicates that if DH does catalyze epimerization, the Leu and Pro in the ‘‘LPFXW’’ motif may be important (Keatinge-Clay, 2008). Further work is necessary to distinguish the above hypotheses.

4.5. The aromatase/cyclase (ARO/CYC) of type II PKS Based on previous studies of first-ring cyclization, there are at least three different classes of ARO/CYCs (Fig. 2.7A): (1) C9–C14 cyclization associated with monodomain ARO/CYCs, such as tcm ARO/CYC, WhiEORFVI (whiE ARO/CYC), and RemI (Alvarez et al., 1996; Fritzsche et al., 2008; Moore and Piel, 2000; Motamedi and Hutchinson, 1987; Zawada and Khosla, 1997); (2) C7–C12 cyclization in the absence of KR by monodomain ARO/CYCs such as ZhuI from the R1128 biosynthetic pathway (Tang et al., 2004), or didomain ARO/CYCs such as MtmQ from the mithramycin pathway (Lombo´ et al., 1996; Zhang et al., 2008) (3) the

O

O

O

O

O

O

Class 1: C9-C14 C7-C16

O 1

S-ACP

TcmN WhiE-ORFVI Reml

OH

OH

14

O

HO

H3CO

16

OH CO2CH3

O

7

whiE-encoded spore pigment (C24, structure unknown)

O 1

OCH3

OH OH O

O

OH

9

O

Tetracenomycin C (antibiotic/anticancer)

O O

OH

OH

S-ACP Resistomycin (antibiotic/antimor)

HO O

OH

Class 2: C7-C12 Zhul MtmQ

O

OH

O

OH

OH

R

R1128 A-D (anticancer) A: R = (CH2)2CH3 B: R = (CH2)3CH3 OH C: R = (CH ) CH(CH ) 2 2 3 2 D: R = (CH2)4CH3

O

12

HO

O HO

OH

O

O

O

S-ACP

7

O OSug H O

KR

OH

Mithramycin (antibiotic/antimor)

SugO OCH3 OH 2

OH

O O

O

O

OH

O

Class 3: C7-C12

12 9

7

HO

O

O S-ACP

ActVII Gris-ORF4

O

OH

O 7

Actinorhodin (antibiotic)

O

12 O

O

CO2H

O

OAC

S-ACP

OH

O

OH

O O

O

OH

Griseusin (antibiotic)

CO2H

Figure 2.7 The three types of ARO/CYCs. Class 1 proteins possess monodomain architecture and promote C9-C14 first-ring and C7-C16 second-ring cyclization of an unreduced polyketide intermediate. Class 2 ARO/CYCs are monodomain or didomain proteins that promote C7-C12 first-ring cyclization of an unreduced intermediate. Class 3 enzymes are didomain proteins that aromatize the first ring of an intermediate that has undergone C9-specific ketoreduction and C7-C12 first-ring cyclization.

34


didomain ARO/CYCs associated with KR-containing type II PKSs that aromatize C7–C12 first-ring cyclized polyketides, such as the actinorhodin (McDaniel et al., 1995) and griseusin (Zawada and Khosla, 1997) ARO/ CYCs. The best-studied monodomain ARO/CYC is tcm ARO/CYC, which consists of the N-terminal 160 residues of the bifunctional protein, TcmN (McDaniel et al., 1995; Zawada and Khosla, 1999). Following the production of the linear decaketide intermediate, tcm ARO/CYC is proposed to fold, cyclize and aromatize the ACP-tethered polyketide via aldol condensation and dehydration reactions (Shen and Hutchinson, 1996). Based on genetic analyses, it had been argued that the first cyclization of a linear polyketide chain may occur either in the active site of KS–CLF (KeatingeClay et al., 2004), in solution (without enzyme catalysis) (Hertweck et al., 2004), or in the binding pocket of KR or ARO/CYC (Zawada and Khosla, 1999). Similar observations on more than ten aromatic PKSs (except in enterocin, discussed below (Hertweck et al., 2004)) have led to the general conclusion that KR promotes C7–C12 cyclization (Crump et al., 1997; McDaniel et al., 1994), whereas the monodomain ARO/CYCs in nonreducing type II systems promote C9–C14 cyclization. The crystal structure and mutagenesis of act KR and tcm ARO/CYC support the hypothesis that cyclization in the KR or ARO/CYC substrate pocket may be a likely event. However, further validation is necessary. Three ARO/CYC structures have been solved: the tcm, whiE, and ZhuI ARO/CYCs, all three of which have a helix-grip fold (Gajhede et al., 1996; Iyer et al., 2001) consisting of a seven-stranded antiparallel b-sheet that partially surrounds (‘‘grips’’) a long C-terminal a-helix (Fig. 2.8). Two small helices between b1 and b2 form a helix-loop-helix motif that acts to seal one end of the b-sandwich. The topology of ARO/CYC is highly similar to that of members of the Bet v1-like superfamily (Iyer et al., 2001; Radauer et al., 2008), which commonly includes a large solvent-accessible pocket that binds small molecules such as phytosteroid hormones, lipids, enediyne, and cholesterol (Markovic-Housley et al., 2003; Mogensen et al., 2002; Pasternak et al., 2005; Radauer et al., 2008; Tsujishita and Hurley, 2000). However, unlike many Bet v1-like proteins with hydrophobic pockets, the ARO/CYC pocket is amphipathic with an approximately equal distribution of hydrophobic and hydrophilic residues. The ARO/CYC pocket dimensions and residue composition are appropriate for binding cyclized polyketide intermediates (Fig. 2.8E). Whereas the type II PKS ARO/CYC contains a helix-grip fold, all FAS DH (such as E. coli FabA) contain a hotdog fold (Dillon and Bateman, 2004; Leesong et al., 1996) (Fig. 2.8A-B) that is topologically different but similar in shape. In the hot-dog fold, the bsheets have strand-order 1–2–3–5–6–7–4, whereas the strand-order is 1–7– 6–5–4–3–2 for the helix-grip fold. Also, the central helix is tucked against the b-sheets in FabA, thus precluding formation of an interior pocket between the b-sheet and central helix aC. As multiple dehydration events

35


C

TcmN ARO/CYC

A

A

N 1

C

B

6 C

7

90⬚

5

3

4

2

N

FabA

B

B

A

N

1

2

3 C

90⬚

5

4

C

D

N

N

C

C

WhiE ORFVI

Zhul

E F88

R82

20 Å 12 Å

7

N

C

C

6

A80 M91 L93 R69 S67 F50

W95 W108

W28

W65

L52 Q110 L129 Y35

T54

W65

N136

A80 M91 L93 S67 R69 W95 F50

T54

W108

W28 L52 Q110 L129

T133 F32

H128 E34 T132 D57

F88

R82

Y35

T133 F32

H128 E34 T132 N136 D57

Figure 2.8 Crystal structures of (A) tcm ARO/CYC, (B) FabA, (C) whiE ARO/CYC, (D) ZhuI ARO/CYC, and (E) The tcm ARO/CYC interior pocket.

are necessary in order to aromatize the rings formed during polyketide biosynthesis it has been suggested that PKS ARO/CYCs may act as DHs to catalyze aromatic ring formation (Hopwood, 1997). That both ARO/ CYC and DH catalyze dehydration reactions in evolutionarily related complexes suggests that they may play similar biological roles implied by their similar topologies. Based on site-directed mutagenesis, the tcm ARO/CYC crystal structure, and computer-simulated docking, a catalytic mechanism was proposed for tcm ARO/CYC (Ames et al., 2008) in which the polyketide carbonyl

36


oxygens are anchored in close proximity to S67, R69, Y35, and R82, and subsequent aldocyclization is promoted by electrostatic stabilization of pocket residues (with special attention to the possible involvement of the strictly conserved pocket residue, R69). The above proposal is further supported by the crystal structures of whiE and ZhuI ARO/CYC (Fig. 2.8C, D). The whiE ARO/CYC promotes C9-C14 first-ring and C7-C16 second-ring cyclizations of a 24-carbon polyketide, and ZhuI is the only reported monodomain C7–C12 first-ring-only cyclase (Marti et al., 2000; Yu et al., 1998). The overall structure and residue composition are similar between whiE and tcm ARO/CYCs, but subtle conformational and residue changes in the whiE ARO/CYC pocket increases the pocket space (compared to tcm ARO/CYC) to accommodate the binding of a C24 polyketide. In contrast, ZhuI has a much smaller pocket that accommodates a monocyclic C7–C12 cyclized intermediate (Fig. 2.8C, D). There are two observed folding patterns of aromatic polyketides which generally lead to unique cyclization regio-specificities (Thomas, 2001). S-type folding, promoted by type II PKSs, leads to C7–12 or C9–C14 first-ring cyclization depending on whether KR is present. In contrast, F-type folding patterns, promoted by the produce template (PT) domain of fungal nonreducing type I PKSs (Crawford et al., 2006, 2008; Udwary et al., 2002), include C2–C7, C4–C9, and C6–C11 first-ring cyclization. ˚ crystal structure and mutational analyses of the PksA PT domain The 1.8-A (manuscript in preparation) revealed that PT also has a DHD fold (Fig. 2.6C) and nearly all secondary structure elements are aligned with both DEBS DH4 ˚ of RMSD. Significantly, and porcine FAS DH domain with only 3 A similarly to ARO/CYC, PT has an interior pocket (Fig. 2.6C), and the reported F-fold patterns may be directly related to their corresponding PKS PT domain, in which the cyclization pattern is determined by pocket shape, while chain length is correlated with pocket size. In conclusion, PT may bind a fully extended linear polyketide that is ‘‘kinked’’ in the cyclization chamber to promote the F-folded cyclization pattern, while the ARO/CYC likely bends the ACP-bound polyketide into a hairpin, thus promoting the S-folded cyclization pattern.

4.6. Acyl carrier protein Recently, the first solved structure of an acyl carrier protein (ACP) from a type I modular PKS was reported for DEBS ACP2 (Alekseyev et al., 2007). Similar to the type I FAS ACP structure, the 10-kD DEBS ACP2 contains a three-helical bundle, and an additional short helix in the second loop also contributes to core helical packing. The conserved Ser in the universal ‘‘DSL’’ motif (where PPT is covalently attached) lies at the N-terminal end of helix-2, which is regarded as a universal ‘‘recognition helix’’ involved in interactions with other proteins (Crump et al., 1997; Findlow


37

et al., 2003; Li et al., 2003; Reed et al., 2003; Zhang et al., 2003). Homology models of DEBS ACP domains (ACP1–6) (Alekseyev et al., 2007) suggest that protein–protein recognition of ACP domains is highly specific for their corresponding KS domains (Chen et al., 2006). Similar results were reported for type II PKS ACP domains, such as the act apo-ACP (Crump et al., 1997), the fren holo-ACP and otc ACP (Findlow et al., 2003; Li et al., 2003). These results are consistent with the ‘‘switch blade’’ theory based on the yeast FAS crystal structure (Leibundgut et al., 2008), in which the growing acyl chain (the blade) switches its nestling cavity from ACP to the KS (or other PKS enzymes) binding pocket, and the timing of blade switching is closely related to the degree of exposure between the polyketide intermediate and the solvent, which depends not only on the ACP ‘‘recognition helix’’ property, but also on the chemical structure of a given polyketide intermediate.

4.7. Thioesterase Three PKS thioesterase (TE) structures have been reported: the DEBS TE (Tsai et al., 2001), and the homologous pikromycin (PIKS) TE (Akey et al., 2006; Giraldes et al., 2006; Tsai et al., 2002), and the PksA TE in a nonreducing iterative type I PKS from aflatoxin biosynthesis (manuscript in preparation). All three structures exhibit the classic features of the a/b hydrolase fold (Fig. 2.9A-B), which consists of a central seven-stranded b-sheet with the second strand (b2) antiparallel to the remaining strands (Chakravarty et al., 2004; Giraldes et al., 2006; Tsai et al., 2001, 2002). While the active-site triad (Ser-His-Asp) and nearly all important secondary structure components are highly conserved, substrate specificity and regiospecificity vary significantly among different TEs. The PKS TE domains lack the characteristic aD helix of the a/b-hydrolase family, instead having an inserted ‘‘lid’’ region that is consistently observed in the DEBS (Tsai et al., 2001), PIKS (Akey et al., 2006; Giraldes et al., 2006; Tsai et al., 2002), surfactin synthetase (SrfA-C) (Bruner et al., 2002), fengycin synthetase (FenB) (Samel et al., 2006), and enterobactin (EntF ) (Frueh et al., 2008) TE structures. The lid region is also the most variable region among the TE domains (Fig. 2.9C). Because the substrate-binding region of the megasynthase TE is formed between the a/b-hydrolase core and the lid region inserted between b6 and b7, variability of the lid region is reflected in the highly variable substrate channel shape among different TEs. In the modular DEBS and PIKS TEs (Akey et al., 2006; Giraldes et al., 2006; Tsai et al., 2001, 2002), an unusual, 20-A˚ long amphipathic substrate channel passes through the entire protein, implying passage of the substrate through the protein (Fig. 2.9C) with the catalytic triad that carries out macrolactonization of a hydrophilic polyketide substrate located at the center. In contrast, the PksA TE substrate pocket adopts a closed conformation,

A

B 2005

Lid loop

1976

L1

1986

1853 1897

2018

L2

G1874 1870 1910 1936

S1937

2045 D

2048

2038

2083

D1964 1938

H2088 2097

N-term 2069 1878 2

4

A

3

B

5

C

6

7

N 1853 1891

E

8

F

2073

1882 1866

1925

1931 1953 1959 2032

2078

2108 C

C

PksA TE (closed)

SrfTE (closed)

FenTE (open)

Human TE (open)

Figure 2.9 (A, B) A typical TE structure with the PksA TE as an example. (C) Comparison of the lid region (yellow) of different megasynthase TE domains. The active site is red.


39

sealed from both ends of the channel, to turn the pocket into a sealed hydrophobic ‘‘cyclization chamber’’ and protect the polyketide substrate from hydrolysis. The flexibility of the lid region is evident when a series of PIKS TE structures solved from crystals at different pH values were compared (Tsai et al., 2002), showing that the size of the substrate channel increases with increasing pH. The variable channel geometry is proposed to influence the regio-selectivity between the hydrolysis and macrocyclization/Claisen cyclization activities of type I PKS TEs. The crystal structures of DEBS and PIKS TE help rationalize the observed substrate specificity of modular PKS TEs (Giraldes et al., 2006; Tsai et al., 2001, 2002), and the chemical structures of the polyketide substrates are as important as the TE substrate residues in determining cyclization versus hydrolysis activity (Gokhale et al., 1999; Lu et al., 2002). Further work is necessary to fully determine residues important for PKS TE substrate specificity.

5. Summary and Future Prospects Recent revelations from crystallographic analysis of type I and type II PKSs have raised awareness of the extraordinary architecture of these megasynthases and offered a new perspective in visualizing some of the unsolved questions concerning polyketide biosynthesis. The 3.2-A˚ porcine FAS crystal structure has provided a framework for megasynthase architecture that may also apply to type I PKSs (Maier et al., 2008). Further, the DEBS KS-AT structures clearly show that the KS-AT and post-AT linkers are highly structured and closely interacting with both KS and AT domains, so that the linker regions contribute extensively to stabilization of the overall KS-AT structure (Tang et al., 2006, 2007). Clearly, we need to reconsider the original notion that the linkers merely serve as semi-flexible tethers that hold adjacent domains in proximity (Gokhale et al., 1999). In the future, studies of interdomain, intermodule, and interpolypeptide linker regions in type I PKSs should further determine their importance to dimer formation, polyketide chain transfer, and reaction timing. In the arena of type II PKSs, the detection of multienzyme complex formation should help shed light on how the ACPs gain access to each of the PKS enzyme domains. Because of the dynamic nature of this complex, x-ray crystallography, if successful at all, may only trap one snapshot of such a transient complex, and other techniques such as electron microscopy or NMR may be necessary to capture a series of protein motions during different stages of polyketide biosynthesis. The early successful application of electron microscopy to capture the FAS dynamic structures (Asturias et al., 2005) can serve as an excellent example for a similar study with type I and type II PKSs.

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ACKNOWLEDGMENTS Our sincere thanks to David Hopwood, Chaitan Khosla, Joel Bruegger, Pouya Javidpour, and Ming Lee for their helpful suggestions and critical reading of the manuscript. Sheryl Tsai is supported by the Pew Foundation, the American Heart Association (0665164Y), and National Institutes of Health R01GM076330 and R21GM077264.

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C H A P T E R

T H R E E

Fungal Type I Polyketide Synthases Russell J. Cox and Thomas J. Simpson Contents 50 54 57 57 61 64 66 66 69 72 74 74

1. Introduction 2. Partially Reducing PKSs: 6-Methylsalicylate Synthase 3. Nonreducing PKSs 3.1. Norsolorinic acid synthase 3.2. Tetrahydroxynaphthalene synthase 3.3. Bikaverin nonaketide synthase 4. Highly Reducing PKSs 4.1. Lovastatin (LNKS and LDKS) 4.2. HR PKS-NRPS: Fusarin and tenellin synthetases 5. NR/HR PKS Hybrid Systems: Zearalenone (ZAE1 and ZAE2) 6. Conclusions References

Abstract Fungi produce a wide variety of biologically active compounds, a large proportion of which are produced by the polyketide biosynthetic pathway. Fungal polyketides comprise a very large and structurally very diverse group, and many display important biological activities, including lovastatin, aflatoxins, and strobilurins. These are produced by very large multifunctional iterative enzymes, the iterative type I polyketide synthases (PKSs) whose closest structural and functional analogues are the mammalian fatty acid synthases. Although fungal polyketides were one of the first classes of secondary metabolites to be subject to extensive biosynthetic studies, they remain the least studied and understood at the enzyme level. This chapter presents an overview of methodologies that have been applied to in vivo and in vitro genetic and biochemical studies on the PKSs responsible for both aromatic and highly reduced polyketide metabolites, and which are providing an improved insight into how these highly complex enzymes function.

School of Chemistry, University of Bristol, Bristol, United Kingdom Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04603-5

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Russell J. Cox and Thomas J. Simpson

1. Introduction Polyketides have long been recognized as one of the most important classes of fungal metabolites (Cox, 2007; O’Hagan, 1991). Fungal polyketides (Fig. 3.1) vary from the simplest monocyclic aromatic compounds, such as orsellinic acid and 6-methylsalicylic acid (6-MSA), to polyclic aromatics such as citrinin, deoxyherqueinone, and norsolorinic acid (NSA). Although initially associated with the formation of aromatic OMe R

Me

Me

Me

CO2H

O

Me

Me OH O

Norsolorinic acid

Deoxyherqueinone Me O

Me

OH

O

O HO

OH

O

O

OH

O

O

OH O

O

OH

O

O O

Me

O

O H

8’

7’

Me H

Me Zearalenone

Me

Me

Cl

OMe

MeO2C Strobilurin B

O Me

Me Me Xenovulence A

Lovastatin

O

OH

O

CO2H

Ph N H Ochratoxin A

O Me

CO2H O HO2C

O

OH

O

OH OH

O OH

Penicillic acid

MeO HO

H

O

H OH

O

Me

O OMe Aflatoxin B1

T-toxin

O O

HO Me

H Me

HO

O

O

O

O

OH Decarestrictine D

Me

OH

H Me

O

HO

Me Me

Citrinin

Orsellinic acid (R=OH) 6-methylsalicylicacid (R=H)

OH

OH

HO

O

HO2C

OH

OH O

O

HO

Me

O

Cl OH 8

O

Me

OH 1

Me

O

OH

O

R

Me

6 3 OH HO O MeO OMe THN (R=H) Bikaverin OH O Fumonisin B1 CO2H O ATHN (R=acetyl, MeCO-) AATHN (R=acetoacetyl, MeCOCH2CO-) OH OH Me O HO O Me OH N O O OH Me OH O HO O OH H Me O Me O SMA76a H Me Me Me HO LNKS hexaketide OH O Me 12 H Me 11 Monacolin J Equisetin 15 Me Me Me O N Me O Tenellin OH H O Me OH O HO CO2Me Me HO Me O Me H N Me HO Fusarin C H Dihydromonacolin L Me Me O N H R NH2

HO2C

OH Me

O

Me

pretenellin (R=H) protenellin D (R=OH)

Figure 3.1 Examples of polyketides synthesized by fungal type I iterative PKSs.

Fungal Type I Polyketide Synthases

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compounds, many fungal polyketides are nonaromatic, such as the macrolides decarestrictine D, long-chain polyfunctional molecules exemplified by T-toxin, and the decalin, lovastatin. Many other metabolites consist of an aromatic ring attached to a more highly reduced moiety, such as zearalenone. Further diversity arises from extensive oxidative metabolism of preformed aromatic polyketides, such as penicillic acid from orsellinic acid and aflatoxin B1 from NSA. Yet other molecules contain a polyketide-derived moiety as part of a larger molecule, the remainder of which comes from other biosynthetic routes, such as the terpenoid humulene in the case of xenovulene A. Fungal polyketides have found wide use in pharmaceutical (Butler, 2005) and agricultural applications. Lovastatin, from Aspergillus terreus, was the progenitor of the now widely used statin group of cholesterol-lowering agents, and strobilurin, from Bolinea lutea, was the key lead for the development of the extensively used methoxyacrylate group of antifungal agents (Clough, 2000). In addition, mycotoxins such as the aflatoxins, ochratoxins, fuminosins, and zearalenone are of major economic importance due to the widespread problems in both human and animal health that can result from fungal contamination of growing and stored crops and foodstuffs. Although diverse in structure, fungal polyketides are defined by their common biosynthetic origin from condensation of simple acyl CoA thiolesters. As long ago as 1953, Birch realized that polyketide biosynthesis is related to fatty acid biosynthesis (Birch and Donovan, 1953), and some of the earliest applications of radioisotopes to natural product biosynthesis were to fungal polyketide metabolites, where the ease of fermentation and isolation of metabolites in pure form, and relatively efficient uptake of simple labeled precursors, facilitates the work (Hanson, 2008). In more recent years, fungal metabolites in general, and polyketides in particular, were the focus of the rapidly expanding applications of stable isotope–labeling methods in the 1970s and 1980s (Simpson, 1987). Fungal PKSs comprise large multifunctional multidomain proteins that assemble these simple acyl units in a highly programmed and iterative fashion to make a huge range of complex natural products, using the simple reactions of Claisen C–C bond formation, followed by varying degrees of reduction and C-methylation via S-adenosylmethionine, each of which is controlled by specific catalytic domains on the PKS (Simpson, 1995). The understanding of the relationship between fatty acid synthase (FAS) and PKS proteins, the application of molecular genetics, and more recently genomics, has greatly facilitated the discovery and understanding of polyketide synthases from diverse sources. The homology in catalytic function between FAS and PKS enzymes is preserved in their respective gene sequences. It is now clear (Cox, 2007) that fungal PKSs belong to the class of type I iterative synthases represented by mammalian FAS. Type I FAS proteins are large multifunctional proteins in which single (or occasionally two)

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peptides contain the sequences for ketosynthase (KS), acyl carrier protein (ACP), malonyl/acyl transferase (MAT), ketoreductase (KR), dehydratase (DH), enoyl reductase (ER) and thiolesterase (TE) activities; these catalytic functions are carried out by particular functional domains. Similarly, the genes for type I PKS proteins are correspondingly large single open reading frames (ORFs) and type I PKSs consist of very large multifunctional proteins, typically 180 to 250 kDa, with individual functional domains. Fungal PKS genes often contain multiple introns, which must be removed prior to heterologous expression in bacteria or yeast. Thus, PKSs use much the same array of chemical reactions as FASs—but the key difference is that of programming: FASs have to control only chain length (i.e. the number of extensions), but PKSs are able to additionally control starter unit selection and the extent of reduction and dehydration during each condensation cycle. Fungal PKSs are also able to program the extent of chain methylation and the off-loading mechanism. The issue of programming is key to understanding and exploiting PKSs. In the case of the bacterial modular polyketide synthases, each condensation cycle is catalyzed by a discreet module containing all the catalytic domains required (Staunton and Weissman, 2001, and other chapters in this volume). In this case the program is explicit in the order and composition of the modules. However, for the iterative type I fungal PKSs, the program is cryptic—encoded in the PKS itself. Understanding the factors that control this programming is arguably the greatest remaining problem in natural product biosynthesis and also represents a great fundamental problem in enzyme catalysis. Thus, the huge structural variety of fungal polyketides is due to differences in programming of their PKS proteins—apparent increases in structural complexity are due to increasing use and control of reductive, dehydrative and methylating steps by the PKS. This must be due to differences in PKS protein sequence and structure. This fact has been exploited in the development of rapid methods for the cloning of fungal PKS genes associated with the biosynthesis of particular fungal polyketide types. Lazarus and Simpson realized that these subtle protein sequence differences should be reflected in DNA sequence and that polymerase chain reaction (PCR) primers could be designed to selectively amplify fragments of fungal PKS genes from fungal genomic DNA (or cDNA). In early work (Bingle et al., 1999), they hypothesized that fungal polyketides could be grouped into two classes: nonreduced (NR) compounds, such as orsellinic acid, NSA and 1,3,6,8-tetrahydroxynaphthalene (THN), and partially reduced (PR) compounds, such as 6-MSA. At the time, very few fungal PKS genes were known, and based on very limited sets of sequences they designed degenerate PCR primers that were complementary to conserved DNA sequences in the KS domains in fungal PKSs responsible for the biosynthesis of NR and PR compounds. Thus, sequences for the available PKS protein sequence KS domains were aligned and inspected

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for regions of high homology, and in particular, sequences that were conserved between PKS sequences for structurally similar compounds but different from conserved sequences for FAS proteins or PKS proteins for structurally different compounds. Additionally, the primers needed to be located away from regions known to harbor introns and be far enough apart to yield PCR products which would be easily analyzed by gel electrophoresis (about 700 bp in this case). Thus, one primer pair (designated LC1/LC2c) was designed to be selective for unreduced PKSs and another set (LC3/LC5c) (Fig. 3.2) for partially reduced PKSs. These primers incorporated degeneracy—that is, included either mixed bases or inosines at defined positions—in order to allow for codon variation in different template organisms. In initial experiments, genomic DNA was prepared from a number of fungi known to produce polyketides. PCR reactions were then performed using the PCR primer pairs, and the products separated by gel electrophoresis. In these reactions conditions were optimized for efficient product formation by varying the ratio of primer concentrations and by including varied amounts of Mg2þ ions and/or varying the PCR annealing temperature. In all cases it was possible to amplify sequences of the expected length. The PCR products were then cloned and sequenced. This showed that the cloned products were always the expected 700-bp fragments of KS domains of PKS genes. Tree-plot analysis, using both the known PKS sequences and

2270

0 1000

KS

KR

LC3 LC1 Cys (1600)

LC2c LC5c

Figure 3.2 PKS primers designed to selectively amplify regions of unreduced and partially reduced fungal PKS genes.

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the PCR products, showed a clear relationship between the clades and the chemical structures of the PKS products (where known). Thus, the primers designed to amplify fragments of unreduced PKSs (LC1/LC2c) gave PCR products which closely grouped with the unreduced clade of known sequences. Likewise the LC3/LC5c primer-amplified products most closely grouped with the partially reduced PKS clade. This was an important result as it showed a clear link between sequence and chemical structure. Later (Nicholson et al., 2001), the same analysis was extended to the KS domains of highly reduced (HR) compounds, such as lovastatin, when DNA sequence data became available for the lovastatin nonaketide and diketide synthases (LNKS and LDKS, respectively) (Hendrickson et al., 1999). The availability of these sequences also allowed the development of selective PCR primers for C-methyltransferase (C-MeT) domains. This sequence analysis has been significantly extended as genomic approaches have been applied to fungi (Kroken et al., 2003). Full genome sequences have now been obtained for more than a dozen fungi. In each organism, many PKS genes have been discovered. For example, Aspergillus niger contains 34 PKS genes (Pel et al., 2007), so there are now several hundred fungal PKS genes known. Sequence comparison of all these new PKS genes, however, shows that the three classes of fungal PKS genes predicted by Simpson and Lazarus are the same three classes observed in the most recent sequence comparisons. Despite the fact that so many fungal PKS genes have been discovered, relatively few genes have been definitively linked to the biosynthesis of specific metabolites. In this chapter, examples of in vivo enzymological studies and associated in vitro genetic manipulations will be presented in outline, and where appropriate in more detail, by discussing progress with a selection of molecules chosen to represent the range of fungal polyketide structures. Further discussion of genetics (Hoffmeister and Keller, 2007), gene cloning (Schu¨mann and Hertweck, 2006), and other biosynthetic aspects (Simpson and Cox, 2009) can be found in a number of recent reviews.

2. Partially Reducing PKSs: 6-Methylsalicylate Synthase 6-Methylsalicylate synthase (MSAS) from Penicillium patulum, at 191 KDa, is one of the smallest type I PKS. It was the first fungal PKS to be purified, in 1970 (Dimroth et al., 1970), and purifications were subsequently described by a number of other workers (Beck et al., 1990; Scott et al., 1974). In 1992, a greatly improved method was described by ShoolinginJordan (Spencer and Jordan, 1992). The synthase was isolated in homogeneous form from P. patulum grown in liquid culture to ensure that the cells


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were all of approximately the same age and morphology. The mycelium was harvested after about 26 h when the enzyme level assayed by fluorimetric determination of 6-MSA (using an incubation mixture (in 2 ml) of Tris/ sulfate buffer, pH 7.6; 160 mM acetyl CoA, 0.4 mole; NADPH, 0.4 mM; BSA 2.5 mg; 0.2 to 1.0 m-unit of MSAS in a total volume of 2 ml at 25 for 10 min, initiated by addition of 0.4 mM of malonyl CoA) was at a maximum. Longer fermentation times lead to decreased yields of purified synthase due to increasing proteolysis in the older cultures. MSAS copurifies initially with P. patulum FAS, but the latter is removed during the purification steps of poly (ethyleneglycol) 6000 precipitation, DAE-sepharose ion-exchange, followed by hydroxyapatite chromatography, and finally FPLC Mono Q purification. The enzyme is highly susceptible to proteolytic degradation, so the purification is carried out in the presence of protease inhibitors phenylmethylsulfonyl fluoride (PMSF), benzamidine and 15% glycerol. It exists as a homotetramer, as determined by gel filtration using a Sephacryl S400 column. It is inactivated by 1,3-dibromopropane, leading to formation of cross-linked dimers, as evidenced by SDS gel electrophoresis. Acetyl CoA and malonyl CoA protect the enzyme against inhibition and dimerization. It will accept acetoacetate as an alternative starter to acetyl CoA. The enzyme also catalyses the formation of small amounts of triacetic acid lactone (TAL) and, in the absence of NADPH, this becomes the exclusive product. This truncation of polyketide chain elongation in the absence of a reductive capability is reminiscent of the lack of fidelity of in vivo chain elongation when the lovastatin and fusarin PKSs, LNKS and FUSS, respectively, are expressed in the absence of the accompanying trans-acting ER domains (see Section 4.1). When the incubation is carried out in the absence of both NADPH and acetyl CoA, other acyl CoA starter units can be utilized, but again only two chain-extending condensations take place, to give a series of alkyl-TAL derivatives, indicative of a ‘‘counting’’ mechanism for chain length control rather than the ‘‘measuring’’ mechanism established for the bacterial type II actinorhodin minimal PKS (Nicholson et al., 2003). Schweizer and coworkers used the purified MSAS to raise antibodies which were used for immunological screening of an Escherichia coli expression library of P. patulum genomic DNA (Beck et al., 1990). This led to isolation of the MSAS gene along with flanking sequences: 7.1 Kb of the cloned genomic DNA was sequenced, the MSAS gene being identified as a 5322-bp ORF coding for a protein of 1774 amino acids. The obtained cDNA revealed a 69-bp intron at the N-terminal part of the MSAS gene. It has a relatively low degree of similarity to the yeast and Penicillium FAS, and a significantly higher sequence similarity was found to the mammalian (rat) FAS. The domain structure is similar to the FAS, with an N-terminal KS followed by AT, DH, and KR domains, terminating with an ACP and no TE or similar release-mechanism domain (Fig. 3.3A).

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A


AT

KS

Loading B

SAT

Core

DH

Processing

Extension KS

AT

ACP

KR

ACP TE/CLC

PT

lovC

ER

C

KS KS

D

KS

AT AT

AT

DH DH

C–MeT

(ER)

C–MeT

DH C–MeT (ER)

KR

ER

KR ACP

Other

ACP ACP

KR

C

A

lovB

C lovF

T R/DCK

Figure 3.3 Representative domain architectures of iterative PKS genes in fungi. (A) MSAS, a partially reducing (PR) PKS from Penicillium patulum; (B) general structure of a nonreducing (NR) PKS; (C) highly reducing (HR) PKSs, LNKS (lovb), and LDKS (lovF) from Aspergillus terreus, and the accessory reductase gene lovC; and (D) general structure of a HR PKS-NRPS. KS, b-ketoacylsynthase; AT, acyl (acetyl and/or malonyl) transferase; C-MeT, C-methyltransferase; DH, dehydratase; KR, ketoreductase; ACP, acyl carrier protein; SAT, starter unit, ACP transcylase; PT, product template; TE, thiolesterase; CLC, Claisen cyclase; ER, enoyl reductase; C, condensation; A, adenylation; T, thiolation/peptidyl carrier protein; R, reductase; DCK, Dieckmann cyclase.

Expression of P. patulum MSAS in heterologous hosts has allowed functional experiments to be performed. It was initially expressed in Streptomyces coelicolor CH999 (Bedford et al., 1995) and subsequently in E. coli and Saccharomyces cerevisiae (Kealey et al., 1998). The latter bacterial and yeast systems required coexpression with a phosphopantetheinyl transferase (PPTase), Sfp from Bacillus subtilis (Lambalot et al., 1996), to ensure that the encoded apo-ACP was converted to its active holo-form. The yields of 6-MSA in the bacterial systems were lower than in P. patulum, but the yeast expression resulted in yields which were twofold higher (1.7 g/l) than in the native host. A 6-MSA encoding gene, atX, was also identified in the genome of A. terreus via Southern blot analysis with the P. patulum MSAS gene (Fujii et al., 1996). Sequencing revealed a 5.5-kb ORF with a 70-bp intron,


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again near the N-terminus. The gene was cloned using the fungal expression vector PTAex3 into Aspergillus nidulans—the consequent production of 6-MSA establishing the function of ATX as a MSAS. The same group went on to develop an S. cerevisiae expression system in which two independent copies of ATX can be produced simultaneously (Moriguchi et al., 2006). This allowed deletion experiments showing that at the N-terminal, 44 amino acids could be removed without loss of activity but that a deletion of 88 amino acids resulted in removal of key KS residues and loss of 6-MSA production. At the C-terminus, removal of as few as nine amino acids resulted in loss of activity. In vivo recombinations of separate deletion mutants, each inactive in isolation, led to restoration of 6-MSAS production, showing that, for example the KS of one peptide chain must interact with the ACP of another. Further deletion analysis allowed the identification of a core interdomain (ID) region, between the DH and KR domains, which is essential for subunit–subunit interactions in the tetrameric structure. These experiments have recently been extended to introduce point mutations in all of the individual domains (Moriguchi et al., 2008). Coexpression of the same single domain mutants gave no 6-MSA production, but coexpression of mutants with different point mutations could all complement one another to restore 6-MSA production. Interestingly, coexpression of two KR mutants gave production of TAL.

3. Nonreducing PKSs 3.1. Norsolorinic acid synthase Norsolorinic acid (NSA) is the first isolable intermediate in the biosynthesis of the potent hepatocarcinogen aflatoxin B1 (Brown et al., 1999). Although initially believed to be a decaketide, formed from an acetate starter unit and nine malonate extender units, isotope labeling experiments suggested that it was likely to be an octaketide primed by a hexanoate starter (McKeown et al., 1996). This was given credence by, inter alia, genetic analysis which showed that transformation of Aspergillus parasiticus with a disruption construct, PXX, blocked aflatoxin biosynthesis (Chang et al., 1995). The disruption was attributed to a single crossover homologous integration event at a specific locus in the A. parasiticus genome, designated pksA. Sequence analysis suggested that pksA is a homologue of the A. nidulans wA gene, a PKS gene involved in conidial pigment biosynthesis (Fig. 3.3B). The sequence contained KS, AT and ACP domains, but no KR or ER were found. The pksA gene lies in the aflatoxin pathway gene cluster adjacent to nor-1, a gene required for conversion of NSA to the next proposed intermediate, averantin, suggesting that PksA was the PKS responsible for the biosynthesis of NSA. Subsequently, Townsend realized that hexA and

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hexB, two genes encoding the a and b subunits of a typical yeast FAS, which were clustered with the NSA PKS, probably produced hexanoate for use as the NSA starter unit (Hitchman et al., 2001). He was able to partially purify (Watanabe and Townsend, 2002) a 1.4-MDa complex, NorS, consisting of these three proteins, from A. parasiticus by diafiltration of a cell-free extract (CFE) prepared from flash-frozen mycelium. The powdered mycelium was suspended in buffer comprising 100 mM potassium phosphate, pH 7.5, 50% glycerol, 2 mM DTT and 1 mM EDTA. The resulting suspension was stirred at 4 for 2 h and centrifuged at 20,000g to give the crude CFE. This was filtered through four layers of cheese cloth, followed by dialysis against buffer supplemented with 30% glycerol, and the final diafiltered CFE was prepared using an Amicon RA 2000 apparatus equipped with a 100,000molecular weight cut-off membrane. This CFE (10 ml) was assayed by incubation with 10 ml [2-14C]-malonyl CoA (25 mCi/ml) and 1 ml hexanoyl CoA (1 mg/ml) and 80 ml of cofactor solution containing NADPH, SAM and FAD (each 1 mg/ml) in distilled water. NSA was detected by TLC autoradiography or HPLC and scintillation counting of collected fractions. The problems associated with working with such a large, labile complex at wild type concentrations precluded further detailed studies. This led to two groups adopting a ‘‘deconstruction’’ approach involving cloning and heterologous expression of individual domains and subsets of the NSAS catalytic domains. A problem associated with this approach is the need to identify the domain boundaries and so the problem of cloning and expressing the part of the PKS gene which will produce soluble, stable, and active protein. The Townsend group has developed a bioinformatics method, the UMA algorithm, which predicts the inter-domain linker regions within related multidomain systems by combining protein sequence similarity, predicted secondary structure and local hydrophobicity (Udwary et al., 2002). The weightings ascribed to the different factors can be varied to give the best indication (low UMA score) of domain boundaries and hence potential cutting and cloning sites. This analysis has revealed (Fig. 3.3B) two hitherto unsuspected domains in NR PKSs (Crawford et al., 2006). The first is a central product template (PT) domain that may be involved in the control of polyketide intermediate chain-length, folding, and stabilization. The PT domain appears to be unrelated to the core ‘‘docking’’ domain of NR PKSs. Cox has carried out a phylogenetic analysis of PT domains from NR PKSs whose products are known and this does reveal a distinct relationship between chain length and clade structure of the resulting treeplot (Bailey et al., 2007). The second is an extended N-terminal domain that can now be seen to function as a starter unit-selecting domain: starter unit-ACP transacylase (SAT). Detailed in silico analysis of SAT domains revealed an unexpected similarity to known malonyl CoA: acyl carrier protein transacylases (malonyl transferases, MAT). The active sites of MAT enzymes contain a


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conserved GHSXG motif in which the conserved serine is known to form an O-acyl intermediate, with a downstream conserved histidine, to give the catalytic dyad (Serre et al., 1995). In the NSAS SAT domain, sequence alignment reveals a similar conserved GXCXG motif and conserved histidine, and it is proposed that the SAT domain substitutes the more reactive thiolester chemistry for the oxy-esters used by MATs in transferring the starter unit to the PKS ACP for subsequent chain elongation. The ACP and SAT domains were cloned, separately expressed, and purified. UMA was used to predict the linker region between the SAT and KS domains to select appropriate oligonucleotides for domain amplification. Three sites were chosen within the linker region for attachment of a C-terminal hexa-histidine (His6) tag in pET28a(þ) to generate pENTC1, pENTC2, and pENTC3. The SAT domain was expressed at 20 in E. coli Rosetta2(DE3), which codes for extra tRNAs to assist expression of eukaryotic genes. Of the three active enzymes, SAT3, arising from pENTC3, demonstrated the greatest stability, and so was used for all further experiments. To demonstrate Cys-117 involvement in the acyl transfer reaction, this proposed catalytic residue was mutated to yield C117A by overlap extension PCR (Ho et al., 1989). Priming sites within putative linker regions surrounding the ACP domain were predicted as above. Because this domain is internal, oligonucleotides were designed with multiple restriction endonuclease sequence tags to facilitate cloning of both the N- and C-terminal His6 tag fusions. The N-terminal His6 tag-fusion expression construct in pET28a(þ), pEACP, gave the soluble ACP domain in E. coli BL21(DE3). MALDI-TOF mass spectrometry indicated a mixture of apo- and holo-ACP, so the purified mixture was fully converted to the holo-ACP by reacting with CoA in the presence of Svp, a PPTase from Streptomyces verticillus (Sanchez et al., 2001). As a control, the invariant phosphopantetheine attachment site was mutated to yield S1746A. The ability of the isolated SAT domain to transfer hexanoate to the ACP domain was demonstrated by incubation of combinations of SAT, SATC117A, holo-ACP and ACP-S1764A, after dialysis against 100 mM potassium phosphate buffer (ph 7.0) in 10 mM final concentrations of each in the presence of 200 mM [1-14C]-hexanoyl CoA at 25 for 2 min. Reactions were quenched with SDS/loading buffer and separated on SDS/12% polyacrylamide gel which, after drying, was exposed to BioMax XAR film. This showed the covalent attachment of hexanoate to SAT and transfer to the holo-ACP. The ability of SAT to transfer different acyl groups was tested by using pantetheine as an acceptor in a modification of a procedure developed by Smith to monitor the ability of KS in mammalian FAS to accept and transfer fatty acyl groups to the ACP (Witkowski et al., 1997). To determine the relative rates, pantetheine (2.5 mM final) and acyl-CoAs (1 mM ) were mixed on ice in 100 mM potassium phosphate (pH 7.0). The mixture was preincubated at 28 for 1 min, and the reaction was started by the addition

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of enzyme preincubated to 28 (0.5 mM final in 100 mM potassium phosphate, pH 7.0). The reaction mixture was incubated at 28 , and 100 ml aliquots were quenched in 100 ml of 8 M urea at room temperature for 2, 4, 6, and 8 min. The quenched reaction samples were frozen in liquid nitrogen and stored at -80 for HPLC analysis. For substrates ranging from acetyl to palmityl, SAT exhibits marked chain-length selectivity, with a strong preference for hexanoyl and octanoyl, with hexanoyl having the faster (about twofold) rate. In an extension of this work, Townsend has used protein database analysis to identify similar FAS-PKS hybrid systems in the pathogenic fungi Coccidioides immitis and Coccidioides posadasii (Crawford et al., 2008a). The NR PKS contains an SAT domain, and similar expression and substrate specificity studies indicate in this case a marked preference for octanoate. This may guide isolation and structure elucidation studies of the, as yet unidentified, metabolites. Further related work on THN synthase will be discussed in the following section. Complementary experiments on NSAS deconstruction have been reported by Cox and coworkers (Ma et al., 2006). They successfully expressed the NSAS AT and ACP as single biochemically active domains. The approach to identifying domain boundaries was simpler and involved performing multiple alignments using a commercial alignment package (AlignX, Invitrogen) and comparisons with type II PKS and FAS components. In contrast to the UMA approach, this allowed the isolation of an active AT (MAT) domain from A. parasiticus. The AT was initially expressed in insoluble form. A refolding strategy involving initial solubilization in CAPS buffer (N-cyclohexyl-3-aminopropanesulfonic acid) and N-lauryl sarcosine resulted in good yields of soluble AT protein, but this protein preparation proved unstable toward precipitation and was inactive in enzyme assays. Similarly, insoluble recombinant AT protein was obtained from the type I mammalian FAS when expressed in E. coli (Rangan and Smith, 1996). They, however, were able to successfully refold the FAS AT in vitro to give soluble active protein. They also showed that a range of AT clones of differing lengths behaved similarly (Rangan et al., 1997). Thus, two additional NSAS AT clones were constructed based on homology to expressed AT domains of the mammalian FAS. The longer of the two AT clones (ATlong) extends by an additional 26 aa at the C-terminus compared with the first AT clone, while the shorter of the two (ATshort) is six amino acids shorter than the first AT clone at the C-terminus. Both new constructs avoid the proline- and lysine-rich sequence (PKSKPK) at the N-terminus of the previous clone. These two new clones were subjected to expression trials, with similar results—very limited soluble protein was observed, with almost all protein produced as inclusion bodies. Good yields of soluble and active NSAS AT were finally achieved using commercially available TB medium (Overnight Express Instant TB Medium, Novagen), and slow overnight expression at 18 led to the


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production of significant amounts of soluble protein for all three AT clones. A single colony was picked and used to inoculate LB medium plus kanamycin (100 ml, 100 mg/ml), which was incubated until the OD600 reached 0.8-1.0 (about 5 h). The mixture was then stored overnight at 4 . The cells were pelleted by centrifugation (SLA 3000, 11,000 rpm, 15 min), then resuspended in fresh LB medium (50 ml) plus kanamycin (100 mg/ml). The resuspended pellet was used to inoculate instant TB medium (Novagen) plus kanamycin (1 l, 100 mg/ml), which was incubated at 37 and 250 rpm for 2 to 3 h, then at 18 and 250 rpm overnight. The cells were harvested by centrifugation (SLA 3000, 11,000 rpm, 15 min), resuspended in an appropriate buffer and stored at –20 until use. In particular, the ATlong clone gave exceptional amounts of soluble protein that could be purified by using Ni2þ affinity chromatography on His-Bind resin (Novagen) and gel filtration (Hiload 26/60 Superdex 75 column). The AT catalyzed the transfer of malonyl groups from CoA onto ACP, but failed to transfer potential starter groups, such as hexanoate, from CoA or FAS-ACP, confirming that it is not involved in PK chain-priming. A mass spectrometric acyl transfer assay was used in which NSAS holo-ACP (50 mM ) was incubated with malonyl CoA or hexanoyl CoA (50 mM—1 mM ) and NSAS ATlong (10 mM ) in phosphate buffer (20 mM, pH 7.5) at 30 . The total volume was 100 ml. After ˚ , Phenomenex) purification method 30 min, the C4 ( Jupiter 15, 300 A (Winston and Fitzgerald, 1998) was used to purify and desalt the reaction mixture prior to analysis by ESMS (QStar). Recently Townsend has reported that NSAS activity can be reconstructed in vitro from partially deconstructed systems (Crawford et al., 2008b). NSA synthetic activity can be observed in a series of systems in which a SAT-KS-MAT tridomain is incubated with varying combinations of the PT, ACP, and TE domains as single-, di- or tri-domain constructs. The importance of the PT domain is shown by the lack of product when it is omitted from the incubations. In the absence of the TE/CLC domain, efficient formation of the final carbocyclic ring is impaired, with a naphthopyrone product predominating. The necessity for the intact SAT-KS-MAT tridomain and its ability to then interact productively with single domains may have important structural implications and again finds parallels in the results from minimal type II PKS systems.

3.2. Tetrahydroxynaphthalene synthase The formally pentaketide-derived 1,3,6,8-terahydroxynaphthalene (THN) is produced in a number of plant pathogenic fungi where it is the key intermediate in the formation of melanin, which is essential for appressorial penetration of plants and thus for the mechanism of pathogenicity towards many commercially important crops (Bell and Wheeler, 1986). As such, the pathway is of great economic importance and a number of effective

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agrochemicals work by inhibiting melanin formation. The conversion of THN to melanin is well understood and, in addition to the PKS responsible for its formation, there are clustered ‘‘ketoreductase’’ and dehydratase genes that act to convert THN to 1,3,8-tri- and 1,8-dihydroxynaphthalenes by successive reduction and dehydration sequences. These are interesting examples of postaromatization deoxygenations of polyketides that contrast with the normal processive mode of carbonyl modification. Interestingly, the fungal melanin ‘‘KRs’’ group with other fungal genes encoding postaromatization reductases, such as versicolorin reductase, and the S. coelicolor type II PKS actIII KR, rather than with the processive KRs of modular PKS and FAS systems (Nicholson, 2000). The dihydroxynaphthalene then undergoes an oxygenase (laccase)-mediated polymerization to produce the melanin pigment. THN is a symmetrical compound with no apparent starter unit. It was presumed to be a pentaketide formed by condensation of acetate with four malonates. A number of possibilities then exist for the folding and condensation of the presumed polyketide intermediate. The THNS-encoding gene has been cloned and sequenced from several fungi, including, as pks1, from Colletotrichum lagenarium (Fujii et al., 1999). Pks1 has been heterologously expressed in Aspergillus oryzae and partially purified (Fujii et al., 2000). The fungal expression plasmid pTAex3, containing the a-amylase (amyB) promoter of A. oryzae and auxotrophic marker argB of A. nidulans, was used. Pks1 gene expression plasmid pTAPSG was constructed based on the PKS1 genomic DNA plasmid pBSPKS. Thus, the N-terminal part of the PKS1 gene amplified by PCR, using its cDNA as a template, was ligated into Xba1/EcoR1-digested pBSPKS to give pBSPSG. The reconstructed pks1 ORF was then inserted into the EcoR1 cloning site just downstream of the amyB promoter of pTAex3 to construct expression plasmid pTAPSG. The argB host fungus A. oryzae M-2–3 was transformed by the protoplast-ethylene glycol method (Gomi et al., 1987). The transformants appearing on minimal plates after about 1 week were precultured in Czapek-Dox medium containing glucose and then transferred into an induction medium of Czapek-Dox containing starch. After 3 days of incubation, the culture medium was acidified with 1 M HCl and extracted with ethyl acetate. This led to the identification of a number of penta- and tetra-ketide metabolites, with THN as the major component. A cell-free extract was then prepared from mycelium from a 3-day-old culture grown at 30 on a rotary shaker at 200 rpm. The mycelium was harvested on a Buchner funnel, washed with distilled water, and flash frozen before pulverization. The resulting powder was suspended in 50 mM potassium phosphate buffer, pH 7.5, containing 30% glycerol, 2 mM b-mercaptoethanol, 1 mM EDTA, and 0.1 mM benzamidine. The mixture was stirred on ice for 20 min and then centrifuged at 10,000g for 20 min. The supernatant was filtered through four layers of gauze and the resulting


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CFE was subjected to ultracentrifugation at 50,000g for 1 h. The supernatant was further fractionated by ultracentrifugation at 21,000g for 2 h. SDS-Page analysis indicated an intense band at 230,000 kDa, corresponding to the molecular weight of the deduced PKS1 protein. Enzymatic synthesis of THN was confirmed by incubation of 150 ml of the CFE in 50 ml of 50 mM potassium phosphate buffer, pH 7.5, 25 ml of 100 mM acetyl CoA, 25 ml of 180 mM [2-14C]-malonyl CoA (5.55 105 dpm) for 30 min at 30 . After acidification with 50 ml of 6 M HCl, the reaction mixture was extracted with ethyl acetate. THN was detected as its tetra-acetate by TLC autoradiography after acetylation with acetic anhydride and pyridine, to establish in vitro synthesis of a multiaromatic ring fungal metabolite for the first time. The incubation and analysis conditions were then slightly modified to allow direct detection by HPLC. The incubations were repeated in the presence of [1-14C]-acetyl CoA and unlabeled malonyl CoA, and in the presence and absence of acetyl CoA with 14C-labeled malonate. In the absence of labeled malonate, THN production was detected by HPLC but it contained no radiolabel. In the other experiments, labeled THN was detected at the same level as before, regardless of the presence or absence of acetate. These experiments clearly demonstrated that no acetate primer was necessary, and this was rationalized by proposing that THNS uses malonate for both chain priming and elongation. While there is no precedent for this in fungal polyketide biosynthesis, the bacterial type III PKSs, such as RPPA, a THNS from S. coelicolor, has been shown to use only malonyl CoA (Funa et al., 1999). In common with several other fungal NR PKSs, PKS1/THNS has a terminal TE/CLC domain (Fig. 3.3B). Claisen cyclase (CLC) domains have been demonstrated to effect release of polyketide product from the PKS by a resorcinol ring-forming Claisen condensation reaction (Fujii et al., 2001). Subsequent work (Watanabe and Ebizuka, 2004) reexamined the products resulting from heterologous expression of PKS1 in A. oryzae. In addition to the major pentaketide product, THN (50%), they isolated the monocyclic pentaketide, a-acetylorsellinic acid (25%), the tetraketide orsellinic acid (10%) and, surprisingly, 15% of a hexaketide, 2-acetyl-1,3,6,8-tetrahydroxynaphthalene (ATHN). They then studied two PKS1 mutants in which the CLC domain had been excised or inactivated by point mutation of the presumed active-site serine (S2009A). Both of these mutants now produced predominantly (95%) the hexaketide product. They explained these observations by proposing a new role for the CLC in controlling chain length, specifically that it intercepted the polyketide intermediate from the ACP domain during chain elongation to produce the shorter chain-lengths. An alternative scenario which obviates the unusual need for malonate as a chain-starter in fungal polyketide biosynthesis, and that is consistent with the formation of THN as the major product of WT PKS1, is that the hexaketide precursor to ATHN is produced by PKS1, and that chainshortening occurs by loss of acetyl, to produce THN. This was indeed

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suggested as long ago as 1983 (Bardshiri and Simpson, 1983), and has recently received experimental verification from Townsend who showed that the SAT domain from PKS1 and a number of other NR PKSs show a marked preference for acetyl CoA over longer-chain acyl CoAs and is essentially inactive towards malonyl CoA (Crawford et al., 2008c). It is possible that C. lagenarium harbors a homologue of AygP, which is known to effect loss of acetoacetate to form THN from the acetoacetyl analogue AATHN in Aspergillus fumigatus (Fujii et al., 2004). However, it is equally likely that THNS/PKS1 catalyses loss of acetyl via a retro-Claisen reaction as an essential part of the mechanism of cyclization and formation of the second aromatic ring.

3.3. Bikaverin nonaketide synthase Bikaverin is a tetracyclic nonaketide-derived metabolite of Gibberella fujikuroi. Its biosynthesis is mediated by the NR PKS gene, pks4. It provides the first example of an active PKS to be expressed and purified from E. coli. Tang and coworkers (Ma et al., 2007) used splice overlap extension PCR to amplify the uninterrupted gene from geonomic DNA from G. fujikuroi. The 6.1-kb gene was inserted into the expression vector pSMa76 which was introduced by transformation into E. coli BL21(DE3) for protein expression. For 1 l of liquid culture, the cells were grown at 37 in LB medium with 35 mg/ml kanamycin to an OD600 of 0.4, at which time the cells were incubated on ice for 10 min, and then induced with 0.1 mM IPTG for 16 h at 16 . The cells were harvested by centrifugation (3500 rpm, 10 min, 4 ), resuspended in 30 ml lysis buffer (20 mM Tris-HCl, pH 7.9, 0.5 M NaCl, 10 mM imidazole), and lysed using sonication on ice. Cellular debris was removed by centrifugation (15,000g, 1 h, 4 ). Ni-NTA agarose resin was added to the supernatant (1 ml/l of culture) and the solution was stirred at 4 for at least 2 h. The protein resin mixture was loaded into a gravityflow column and proteins were purified with increasing concentrations of imidazole in Buffer A (50 mM Tris-HCl, pH 7.9, 2 mM EDTA, 2 mM DTT). Purified proteins were concentrated and buffer exchanged into Buffer A þ 10% glycerol with Centriprep filter devices (Amicon Inc.). The final PKS4 enzyme was concentrated to 8 mg/ml, aliquoted, and flash frozen. SDS gel analysis showed a single band at 220 kDa. PKS4 was produced in an excellent yield of about 8 mg/l. The purified PKS4 (10 mM ) was assayed by incubation with 2 mM [2-14C]-malonyl CoA (4.5 mCi/mmol) in Buffer R (100 mM NaH2PO4, pH 7.4, 10% glycerol, 2 mM DTT) at room temperature. Individual aliquots (20 ml) were removed at different time points and were quenched with 250 ml of 99% ethyl acetate/1% acetic acid. The organic phase was separated, evaporated to dryness, re-dissolved in 20 ml of ethyl acetate, and the reaction mixture was separated by TLC with 99% ethyl acetate/1% acetic acid as the mobile phase. The resultant


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TLC plate, which was imaged and quantified by phosphoimager (InstantImager, Packard). This showed a new compound, which LC-MS analysis confirmed has a molecular weight of 322, in agreement with an unmethylated and unoxidized precursor of bikaverin. The identity of the new compound, SMA76a, was confirmed by scale-up of the in vitro reaction using the malonyl CoA synthase MatB from Rhizobium trifolii to generate malonyl CoA in situ from sodium malonate. The new compound was produced in a remarkable yield of about 100 mg/l. PKS4 appears to be efficiently phosphopantetheinylated by the endogenous E. coli holo-ACP synthase (ACPS), as assay of PKS4 expressed in E. coli BAP1, which contains a chromosomal copy of the broad-specificity PPTase Sfp, produced similar results to BL21(DE3) expression. This is in marked contrast to all previous studies on expression of PKS proteins or domains, such as expression of type II PKS ACPs, where efficient post-translational modification required coexpression of the ACP genes with a copy of the ACPS gene (Cox et al., 1997). Another feature is the apparent lack of requirement for acetyl CoA. This seemed to be consistent with the PKS4 lacking the coding sequence for the required cysteine in the CXCXG motif normally observed in SAT domains. However, Townsend has reported a frame-shift mutation in the published sequence on resequencing exons 1 and 2, which results in an alternative splicing pattern that does indeed contain the required motif (Crawford et al., 2008c). Using his SAT phosphopantetheine assay, he showed that the PKS4 SAT domain shows a preference for acetyl CoA and negligible activity with malonyl CoA. In light of these results and the THNS results discussed above, the suggested use of malonyl CoA as a starter appears unlikely and an alternative explanation for the apparent lack of requirement for acetyl CoA may be required. When various alkylacyl CoAs are added to the PKS4 incubation, PKS4 did not utilize any of the short-chain (C2 to C6) acyl CoAs but did produce two new benzopyrone metabolites when incubated with octanoyl CoA. This decreased the yield of SMA76a 10-fold. Significantly, the overall size of the new products is 18 carbons, the same as bikaverin and SMA76a. This is reminiscent of the bacterial type II minimal PKS where increasing the length of the starter unit decreases the number of condensations so that overall chain length remains constant (Nicholson et al., 2003). For the starter unit assays, alkylacyl-CoAs were each added to a final concentration of 2 mM. For LC-MS analysis, the same reaction mixture (100 ml) was prepared with unlabeled malonyl CoA (2 mM ). The organic residue was redissolved in methanol and analyzed by LC-MS with a Finnigan LCQ Deca XP quadrupole ion trap mass spectrometer using negative electrospray ionization and a Waters 2.1 100 mm C18 reverse-phase column. Samples were separated on a linear gradient of 5 to 95% CH3CN (v/v) over 30 min and 95% CH3CN (v/v) for a further 30 min in H2O supplemented with 0.05% (v/v) formic acid at a flow rate of 0.125 ml/min at room temperature.

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4. Highly Reducing PKSs 4.1. Lovastatin (LNKS and LDKS) Lovastatin, a metabolite of A. terreus, is an inhibitor of the enzyme 3-hydroxy-3-methlyglutaryl (HMG) CoA reductase and thus is a potent inhibitor of cholesterol biosynthesis. Lovastatin and other natural and synthetic derivatives constitute the statin group of pharmaceutical agents which are amongst the major selling pharmaceutical agents of all time (Tobert, 2003). It is also the archetypal example of a metabolite produced via a highly reducing (HR) PKS and its biosynthesis has been the subject of extensive isotopic labeling, molecular genetic and biochemical studies (Sutherland et al., 2001). It comprises a decalin, formed by a presumed Diels-Alder cyclization of a highly reduced and monomethylated nonaketide precursor, to which is attached, via an ester linkage, a diketide-derived 3-methylbutyrate moiety. The biosynthetic genes were isolated by screening mutants of lovastatin-producing strains (Hendrickson et al., 1999). Several mutants were found which were deficient in production of either the nonaketide- or diketide-derived moieties, indicating that two separate PKSs were likely involved. A genomic cosmid library was constructed in an A. terreus/E. coli shuttle cosmid (pLO9) and this was used to complement the BX102 mutant, which was deficient in lovastatin production but which, when fed the nonaketide-derived monacolin J, efficiently converted it to lovastatin, suggesting that it was deficient in the gene encoding the lovastatin nonaketide synthase (LNKS). Comparison of crude protein extracts of the wild type and BX102 strains indicated that BX102 was completely deficient in a about 250-kDa protein. Conversely, a different polypeptide of about 220 kDa was missing from monacolin J–producing mutants. On transformation, one lovastatin-producing transformant was identified and the cosmid was recovered by in vitro packaging of its genomic DNA. In addition, cDNA libraries were prepared in the E. coli vector lgt11 using mRNA from a lovastatin-producing culture and were screened with antiserum raised against the 250-kDa protein. This led to the isolation of 18 clones. Two of these were found to hybridize to the BX102-complementing cosmid clone. One of the two isolated cDNA clones was then used to isolate overlapping clones from the genomic library to give an 11.6-kb region coding for the LNKS (Fig. 3.3C). Specific primers were then used to obtain the cDNA sequence by PCR. Using the LNKS gene (lovB) as a probe, Hutchinson and coworkers (Kennedy et al., 1999) isolated cosmids containing lovB and the surrounding genomic DNA from an A. terreus genomic library. DNA sequencing and analysis of cosmids pWHM1263 and pWHM1265 revealed 10 potential genes over 64 kb, whose functions could be predicted by sequence comparisons. Of these, two—lovB and lovF—encode PKSs, the latter the lovastatin diketide synthase (LDKS).


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To further investigate the role of lovB, it was heterologously expressed in A. nidulans under the control of the alcA promoter. This showed a major high molecular weight protein on SDS-polyacrylamide gel electrophoresis. The protein cross-reacted with antibodies to LNKS, confirming its overexpression. Further growth and induction by the alcA promoter resulted in the isolation of two compounds which were not immediately related to monacolin J. These were two monomethylated fully unsaturated hexaketide- and heptaketide-derived pyrones. An A. terreus mutant disrupted in the lovC gene, encoding a putative enoyl reductase (ER), was blocked in lovastatin production, but also produced the hexaketide pyrone metabolite. The formation of this truncated and incompletely processed polyketide suggested that LNKS and LovC must interact to produce the polyketide of the correct length and correct reduction and cyclization pattern. Thus, lovC was placed under the control of the alcA promoter and was used to transform strain WMH1738 containing LNKS. Transformant colonies were grown under inducing conditions for alcA and the extracts were analyzed by TLC and HPLC. Several transformants, including WHM1750, were found to produce dihydromonacolin L, which therefore must be the immediate product of LNKS (plus LovC), and greatly reduced amounts of the aberrant pyrones. The fact that LNKS and LovC together make the functional PKS for monacolin production was unexpected at the time and had few precedents, such as the formation of TAL from MSAS alluded to in Section 2 (Spencer and Jordan, 1992). While LNKS does possess a putative ER domain, this suggests that it is inactive, or at least inactive on its own, and that the PKS requires the trans association with the ER encoded by lovC to produce a fulllength, correctly reduced nonaketide. This has subsequently been observed with a number of other, hybrid, PKS-NRPS systems (Halo et al., 2008). An important step in lovastatin biosynthesis is the intramolecular DielsAlder cyclization of the polyunsaturated nonaketide or an earlier precursor. Using a combination of synthesis of putative substrates and purified LNKS, along with a careful analysis of presumed spontaneous cyclization and enzyme-mediated products, Vederas has provided evidence that LNKS does harbor such a Diels-Alderase activity (Auclair et al., 2000), but the evidence suggests that cyclization occurs on an enzyme bound hexaketide intermediate, suitably activated for the Diels-Alder reaction, which is subsequently extended to the nonaketide level. Further biochemical characterization of LovB has been obtained by expressing some of its constituent domains as stand-alone mono- and di-domains (Ma and Tang, 2007). Domain boundaries were identified via sequence alignment with bacterial type I PKSs and mammalian and fungal FASs. PCR primers were designed to amplify the designated target domains using the uninterrupted lovB sequence obtained by removal of the seven introns, using splice by overlap PCR. PET28a(þ) expression vectors were constructed for the KS-MAT didomain, the ACP monodomain and the

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ACP-CON didomain. The KS-MAT (pSMA30) and CON (pSMA61) constructs were introduced by transformation into BL21(DE3) for protein expression and the ACP-containing constructs were introduced by transformation into BAP1 containing the broad spectrum PPTase gene sfp. Good yields of soluble protein were generally obtained. Addition of the CON domain (see Section 4.2 for further discussion) increased yields from 1 mg/l for the ACP alone to 15 mg/l for the didomain. These results contrast with the work of Townsend and Cox who both found E. coli expression of fungal NSAS domains to be very difficult (Section 3.1). The ability of the domains to accept acetate and malonate was assayed by incubation of the proteins (1—10 mM ) in buffer l (100 mM NaH2PO4, pH 7.4, 2 mM DTT, 10% glycerol) along with added acyl CoA (180 mM, 55 mCi/mmole) in a final volume of 10 ml. After 10 min at room temperature or on ice, reactions were quenched with one volume of SDSPAGE loading buffer lacking any reducing agents such as DTT or mercaptoethanol. The proteins were separated on 6% or 12% SDS-PAGE gels and analyzed using a phosphoimager. During purification of the KAS-MAT didomain, a truncated MAT domain was obtained as a result of proteolytic cleavage during cell lysis. Interestingly, the cleavage site, EY448/M449EPEQ, corresponds to a region that is highly conserved among fungal and bacterial type I PKSs (Rangan et al., 1997). Comparison with the crystal structure of the DEBS module-5 KS-AT from Saccharopolyspora erythrea (Tang et al., 2006) suggests that the cleavage site is at the start of a structurally highly ordered linker region. No similar proteolytic susceptibility was observed, however, during purification of the DEBS KS-AT or mammalian FAS-MAT didomains (Witkowski et al., 2004). Using this sequence information, the MAT domain was cloned and expressed in E. coli. An exceptionally high yield (50 mg/l) was obtained. The lovB MAT active site region contains two consecutive serines (GHS656S657G) and this diad appears to occur frequently in fungal PKSs. This contrasts with FAS MAT, where a single active-site serine is normal (GHSXG). Both sites in lovB MAT were separately lovB mutated to alanine, and labeling assays indicated that S656, which corresponds to the normal serine position, was the only one to be labeled from malonyl CoA. The amount of soluble protein obtained from the S656A mutant was similar to the wild type domain (50 mg/l), but the S657A mutant gave only poor yield (2 mg/l). It is suggested, therefore, that the second serine may contribute to the overall folding and structural integrity of the domain. Attempts to express the KS as a standalone protein failed to give soluble protein, similarly to efforts to express the NSAS KS. Proteolytic cleavage of the KS domain is also observed for the LovB protein itself. It can be expressed as soluble protein in reasonably high yields (5 mg/l) with a C-terminal His-tag in BAP1. During Ni affinity chromatography, however, a slightly smaller protein coeluted. Both bands were strongly labeled from [14C]-malonyl CoA, indicating that the smaller band must contain an intact MAT domain and has likely lost the KS.


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4.2. HR PKS-NRPS: Fusarin and tenellin synthetases LovB appears to have part of a non–ribosomal peptide synthase (NRPS) condensation (C or CON) domain downstream of the ACP. While this was surprising at the time, it was proposed (Cox, 2007) that this may be involved in release of the product from the synthase, either by activating water or the C-5 hydroxyl to form the d-lactone moiety of lovastatin directly. However, the presence of NRPS domains downstream of a fungal HR PKS is now known to be rather common. Cox and coworkers were investigating the biosynthesis of fusarin C, a toxic metabolite of inter alia Fusarium monilforme and Fusarium venenatum. Fusarin C is representative of a large group of fungal metabolites in which a highly reduced and often highly methylated polyketide moiety is fused to a pyrrolidone, tetramic acid, or pyridone moiety, of presumed Krebs cycle or amino acid origin. Using a cloning strategy previously applied to squalestatin (Cox et al., 2004), based on the use of degenerate C-MeT PCR primers, they were able to rapidly clone about 26 kb of genomic DNA from F. monilorme containing the fusarin synthetase and several other ORFs (Song et al., 2004). FUSS was shown to be the fusarin synthetase by inactivation of the gene and concomitant loss of metabolite production. They found that the fusarin synthetase (FUSS) combines an HR PKS with a full NRPS module (Fig. 3.3D), presumably in this case selective for the activation and incorporation of homoserine. This was confirmed by the synthesis and intact incorporation of [2-13C,15N] homoserine into fusarin C (Rees et al., 2007). Subsequent to this initial observation, a number of other tetramic acid/pyridone-containing metabolites, including equisetin (Sims et al., 2005) and tenellin (Eley et al., 2007), have been shown to be produced via similar PKS-NRPS systems. All of the PKS-NRPS genes so far reported terminate with an apparent reductase (R) domain (Fig. 3.3D) and, in the case of fusarin, this would be consistent with the required reductive cleavage of the homoserine thiolester linkage to the NRPS thiolation (peptidyl carrier protein/T) domain to give the corresponding aldehyde, followed by an aldol cyclization to give the final pyrrolidone ring. However, this is not the case with equisetin and tenellin, which both require an unreduced carboxylate/thiolester for tetramic acid formation. The reductase domain has been shown through heterologous expression (Halo et al., 2008) to act alternatively as a condensing (Dieckmann cyclase [DCK]) domain. This proposal has since been supported by overexpression of the equisetin R-domain and studies of its reactivity with substrate analogues (Sims and Schmidt, 2008). The C-terminal R-domain was cloned as a free-standing domain and also as a didomain with the adjacent T-domain and both were expressed in E. coli in His-tagged form and purified to homogeneity using Ni-NTA resin, followed by FPLC with a Superdex SD200 size-exclusion column. Synthetic substrates, N-acetyl- and N-acetoacetyl-alanine, were synthesized as their

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N-acetylcysteamine (SNAC) and CoA thiolester derivatives. The CoA analogues were covalently linked to the T-domain using the usual sfp PPTase. Incubation with excess NADH or NADPH failed to give any evidence for reduction of the substrates when assayed by ESI-MS. The purified R-domain was incubated with the acetoacetyl-alanine thiolester substrates, both with and without NADPH, and the reactions monitored by TLC and GC-MS. At pH 8, nonenzymatic background formation of tetramic acid was rapid, while at pH 6, no tetramic acid formation was observed in the presence or absence of enzyme. It was found that pH 7.0 gives negligible nonenzymatic reaction but rapid enzyme-catalyzed production. The acetyl analogues were not substrates and no aldehyde or alcohol products were observed in the presence of NADPH, which was not required for tetramic acid formation. Two mechanisms are possible: either the enzyme catalyses a Dieckmann cyclization, or it recycles bound NADPH to both reduce and oxidize the substrate. Although this appears complicated, this mechanism has been demonstrated for sugar epimerase enzymes that are homologous with the R-domains (Mayer and Tanner, 2007). Interestingly, 3-methylorcinaldehyde synthase (MOS), which is responsible for the tetraketide-derived moiety in xenovulene A, is an NR PKS, which terminates with a C-terminal R-domain very similar in sequence to the fusarin and equisetin R-domains. In this case it is active as a reductase, removing the polyketide product from the PKS by reduction of the ACP-thiolester linkage to give the aldehyde product (Bailey et al., 2007). The fusarin PKS contains an ER domain which appears to be inactive, as no ER-mediated reduction is actually required for fusarin C production. The tenellin PKS (TENS) also contains an ER domain, which also appears to be inactive, cf. LNKS. However, the TENS gene cluster also contains an encoded LovC homologue, ORF3. Like LovC, this is essential for correct programming and fidelity in assembly of the polyketide precursor. When TENS was heterologously expressed in A. oryzae on its own, polyketidederived products were obtained. These were all tetramic acids, not pyridones, and contained polyketide moieties that not only lacked the essential ER-mediated reduction at carbons 11 and 12, but also differed in chain length, and methylation (Halo et al., 2008). In order to establish the essential role of the putative trans-acting ER encoded by orf3, a pTAex3 derivative was constructed in which selection was provided by a Basta-resistance (bar) cassette. B. bassiana is sensitive to the herbicide ammonium glufosinate (Basta) and can be used as a selection marker for the construction of both knockout (KO) and antisense RNA (see below) vectors. The argB gene was inactivated by truncation. This allowed cointroduction by transformation of the tenS/argB vector into A. oryzae. Dual selection was achieved by plating the transformants on a minimal medium lacking arginine and overlaying after 24 h with Basta, followed by subculturing single colonies into Bastacontaining liquid medium lacking arginine. Eleven clones were isolated


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which showed both Basta resistance and arginine prototrophy. PCR analysis of their genomic DNA confirmed the presence of both tensS and orf3. Five clones were grown in production medium and the extracts were examined by LC-MS. Four of the clones produced a single new compound, pretenellin A, which contained the correctly assembled polyketide portion fused to a phenylalanine-derived tetramic acid. To establish that pretenellin A was indeed an intermediate in tenellin biosynthesis, it was incubated in a CFE. A 10-day-old culture (500 ml) of a B. bassiana blocked mutant (tenS KO strain, Eley et al., 2007) grown in tenellin production medium was harvested by vacuum filtration. The extract was prepared using the method of Watanabe and Townsend (1998). The mycelium was thoroughly washed with distilled water, 35g wet-weight of the mycelium was flash frozen in liquid nitrogen, pulverized using a pestle and mortar, and re-suspended in 70 ml buffer solution (50 mM potassium phosphate, pH 7.5; 30% glycerol; 2 mM DTT; 100 mM PMSF, 100 mM benzamidine hydrochloride and 1 mM EDTA). The mixture was stirred for 2 h at 4 , then centrifuged at 20,000g for 20 min. The supernatant was decanted and used as CFE. 20 ml of 1 mg/ml solution of pretenellin A in acetone was added to 5 ml of CFE in a precooled 10-ml tube. The reaction was incubated at 25 for 30 min, then quenched by shaking with ethyl acetate (5 ml). The organic layer was carefully decanted after the mixture was allowed to settle and concentrated in vacuo. The sample was dissolved in 100 ml of HPLC methanol and subjected to LCMS analysis. Control experiments were conducted following the same protocols, using a boiled CFE solution. Two pyridone-containing metabolites, 15-hydroxytenellin and pyridovericin (also 15-hydroxylated but lacking the N-hydroxyl), were detected. These have also been detected as minor metabolites of the wild type strain of B. bassiana. Thus, it appears that the CFE has been elevated in the oxidase responsible for this extra side-chain hydroxylation. The gene responsible for the ring expansion has been demonstrated to be encoded by orf1 by gene KO, selective gene silencing and coexpression approaches. Gene KO strategies were previously used in B. bassiana through the insertion of an antibiotic-resistance cassette into the target gene via homologous recombination. However, the tenS KO required examination of nearly 100 individual clones to find a bone fide KO as the rate of ectopic integration of the selection marker in B. bassiana was high (>90%). In order to improve the chance of success, an RNA silencing approach was also used—specifically antisense RNA—as this avoids the requirement for homologous recombination (Heneghan et al., 2007). In order to construct KO vectors for orf1 and orf2, PCR was performed on B. bassiana genomic DNA to amplify 50 - and 30 -targeting fragments (about 700 to 1200 bp) for each ORF. All the PCR products were initially cloned into pENTR/ D-TOPO (Invitrogen), and the KO plasmids were constructed by inserting

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a bar cassette (A. nidulans trpC promoter driving a Basta resistance gene) between the targeting fragments. Creation of an antisense RNA vector required choice of a suitable promoter. The A. nidulans gpdA (glyceraldehyde-3-phosphate dehydrogenase) promoter has previously been demonstrated to drive strong constitutive expression in B. bassiana (dos Reisa et al., 2004), so it was selected to ensure high levels of antisense RNA production. The promoter was amplified by PCR and inserted into vector pCB1530, which contains a selectable bar cassette. To enable rapid cloning of diverse gene fragments a Gateway cloning cassette (Shafran et al., 2008) was inserted ‘‘in reverse orientation’’ downstream of the gpdA promoter (i.e., with attR2 adjacent to the promoter), creating fungal expression vector pCBgpdA-GA. Orf1 and orf2 were amplified by PCR and cloned directionally into pENTR/D-TOPO (Invitrogen). Gateway recombination was then used to insert the fragments in antisense orientation with respect to the gpdA promoter into pCBgpdA-GA. The two linearized KO and two circular antisense RNA plasmids were then introduced into B. bassiana by PEGmediated protoplast transformation and selection for Basta resistance. A total of 18 potential Dorf1 KO and 17 Dorf2 KO transformants were obtained, in which the intended double crossovers would have resulted in gene KO. A total of 29 antisense RNA transformants were obtained for orf1 and 39 antisense RNA transformants for orf2. The KO and antisense transformants were grown in production medium, extracted and analyzed. Of the KO transformants, all but one showed the wild type phenotype, with only one orf1 transformant producing pretenellin A and its 16-hydroxy derivative, protenellin D. Several of the orf1 antisense RNA strains, however, produced pretenellin A and no detectable pyridones. Overall, analysis of these and related results confirm the role of TENS and ORF3 in production of pretenellin A, with ORF1 being responsible for ring-expansion of tetramic acid to pyridone and ORF2 for the final N-hydroxylation to produce tenellin.

5. NR/HR PKS Hybrid Systems: Zearalenone (ZAE1 and ZAE2) The studies described above on NSA biosynthesis (Section 3.1) attest to the ability of NR PKSs to accept chain primers from other synthases, in this case a dedicated FAS. It is now apparent that this is a more widespread phenomenon and that HR PKSs can also provide more complex polyketide-derived primers to be chain extended by an NR PKS. The best-studied example to date is the fungal toxin zearalenone, formally a nonaketide, whose structure indicates a high level of reductive modification early in the biosynthesis and no reduction during the latter (three) chain


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elongations. It has now been shown that the zearlaneone gene cluster from Gibberella zeae (Gaffoor and Trail, 2006) contains two iterative PKSs, PKS4 (ZEA2) and PKS13 (ZEA1), both of which are essential for zearalenone biosynthesis. The former contains the expected domains for a HR PKS, whereas the latter has the expected SAT, KS, MAT, and ACP, terminating with a thioesterase (TE). This has been given biochemical verification by Reeves and coworkers, who sequenced the gene clusters for the zearalenone analogues, hypothemycin, and radicicol biosynthesis from Hypomyces subiculosis and Pochonia chlamydosporia, respectively (Reeves et al., 2008), cloned and expressed a number of the genes from the hypothemycin cluster in both E. coli and yeast and showed that both the HR-PKS and NR-PKS must be simultaneously expressed in yeast for production of the zearalenone analogue 70 ,80 -dehydrozearalenone (DHZ). Hypothemycin cDNAs were expressed from the adh2 promoter in 2m-plasmid–based yeast-E. coli shuttle vectors (Mutka et al., 2006) in an S. cerevisiae strain developed for fungal PKS expression (Kealey et al., 1998). The HR-PKS cDNA was cloned into pKOS187-98A (LEU2 marker) as an NdeI-SbfI fragment to obtain pKOS518-118A. The NR-PKS was cloned into pKOS247-14-3 (TRP1 marker) cut with NdeI and EcoRI in a threepiece ligation with the 0.75-kb NdeI-AlwNI and 5.4-kb AlwNI-EcoRI fragments from pKOS518-117A to obtain pKOS518-120A. The P450 and FMO cDNAs were cloned into pKOS187-98A as NdeI-AvrII fragments to obtain pKOS518-116 and pKOS518-137A, respectively. The OMT cDNA was cloned as a 1.6-kb NdeI-EcoRI fragment from pKOS51892C into pKOS247-14-3 to obtain pKOS518-136A. Each cDNA expression cassette flanked by the adh2 promoter and terminator was moved into pKOS187-4e (URA3 marker) (Mutka et al., 2006), using the BssHII sites, to obtain pKOS518-137B (P450), pKOS518-138A (FMO), and pKOS518138D (OMT). Plasmids were introduced into yeast by the LiCl-PEG procedure, with plating on the appropriate complete minimal dropout agar plates (Teknova). Colonies were transferred to the appropriate liquid complete minimal dropout medium and grown at 30 to generate inoculum for YPD medium containing 2% glucose and 20 g/l of Amberlite XAD1180 resin (Alfa Aesar). After 24 h of incubation at 30 , the resin was collected and washed with water, and the products were eluted with methanol. The methanol was evaporated, and the residue was dissolved in 80% acetonitrile for HPLC analysis, which confirmed that expression of both PKS genes, but neither alone, gave DHZ. Adding expression of the O-methyltransferase (OMT), flavin mono-oxygenase (FMO) or cytochrome P450 to the strain brought about methylation, epoxidation, or hydroxylation of DHZ, respectively. In a related study, Tang and coworkers cloned, overexpressed, and purified the G. fujikuroi PKS13 and, using a small molecule mimic (the SNAC thiol ester of 10-hydroxydecanoic acid) of the natural hexaketide

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precursor, showed that it is efficiently utilized to chain-prime PKS13, which then carries out the full set of chain extension, cyclization, aromatization, and macrolactone formation reactions (Zhou et al., 2008). Other fatty acyl CoAs from C-6 to C-16 were also utilized, with the highest activity with decanoyl CoA. In E. coli, PKS13 synthesized new compounds using endogenous acyl starter units or when the cultures were supplemented with a variety of synthetic precursors, demonstrating its potential for precursor-directed biosynthesis in vivo as well as in vitro.

6. Conclusions A combination of molecular genetic and enzymological studies have begun to shed light on how these complex iterative multifunctional enzyme systems actually work, but we still have little understanding of the fundamental programming mechanisms that govern starter unit selection and control of chain length, degree of methylation, and reduction. However, the studies described above do confirm that it will be possible to dissect these complex systems to study the functions of individual domains and subsets of domains at a biochemical level. This must be the key to achieving a full understanding of how programming works, and thus how it might be exploited for synthetic biology.

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Tandem Acyl Carrier Protein Domains in Polyunsaturated Fatty Acid Synthases Hui Jiang,* Scott R. Rajski,* and Ben Shen*,† Contents 1. Introduction 2. Methods 2.1. Production of PUFAs in E. coli by expressing the PUFAS genes 2.2. Mapping the active sites of PfaA-ACPs by site-directed mutagenesis 2.3. Overproduction of each of the PfaA-ACPs 2.4. Overproduction of PfaE and Svp PPTases 2.5. In vivo and in vitro preparation of the holo-form of PfaA-ACPs 2.6. Elucidation of the relationship between PUFA production and the number of active ACPs 3. Conclusion Acknowledgement References

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Abstract Polyunsaturated fatty acids (PUFAs) can be biosynthesized via aerobic pathways that rely on combinations of desaturases and elongases to convert saturated fatty acids to PUFAs or anaerobic pathways that exploit polyketide synthase (PKS)-like enzymes known as PUFA synthases for de novo synthesis from acyl CoA precursors. In contrast to most fatty acid synthases (FASs) and PKSs that contain a single acyl carrier protein (ACP) domain for each cycle of fatty acid or polyketide chain elongation, all PUFA synthases known to date contain tandem ACPs (ranging from five to nine). The roles and engineering potential of such tandem ACPs in PUFA synthases remain largely unknown, although the growing demand for PUFAs and decline of current sources dictate that a greater understanding of these PUFA synthases is not only warranted, but urgently needed. This chapter describes methods and protocols developed to

* {

Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Department of Chemistry, University of Wisconsin-Madison, Madison, Wisconsin, USA


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dissect the role and underlying biochemistry of each of the PfaA-ACPs in the Shewanella japonica PUFA synthase for eicosapentaenoic acid (EPA) and docosapentaenoic acid (DPA) biosynthesis. These studies have set the stage to interrogate the roles of the other domains and subunits of the Pfa PUFA synthase in EPA and DPA biosynthesis. Applications of the methods and protocols described here to other PUFA synthases are therefore envisioned to help close the knowledge gap currently limiting microbial production of PUFAs via PUFA synthase engineering and heterologous expression.

1. Introduction Polyunsaturated fatty acids (PUFAs), such as arachidonic acid (AA), eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA), are essential to human health and nutrition (Berge´ and Barnathan, 2005; Graham et al., 2007; Muskiet and Kemperman, 2006; Wallis et al., 2002) (Fig. 4.1). Oceanic fish and fish oil products are the typical sources of PUFAs (Graham et al., 2007). However, these sources possess a number of problems including poor taste, instability, and variability. Moreover, PUFAs from these sources are produced as complex mixtures requiring extensive purification (Berge´ and Barnathan, 2005). Global warming, pollution, and overfishing represent further challenges to the present and future availability of these vital nutrients (Graham et al., 2007). These considerations, compounded by the increasing knowledge of PUFAs’ beneficial roles in human health and a rapidly expanding global population, necessitate the development of new and sustainable PUFA sources to meet growing demand (Berge´ and Barnathan, 2005). Although industrial-scale production 5

1 CO2H CH3 w

Arachidonic acid (AA) 20:4Δ5,8,11,14 20:4n6 CO2H CH3 w Eicosapentaenoic acid (EPA) 20:5Δ5,8,11,14,17 20:5n3

10

7 CO2H CH3

13 16 19 Docosapentaenoic acid (DPA) 22:5Δ7,10,13,16,19 22:5n3 4 CO2H CH3 Docosahexaenoic acid (DHA) 22:6Δ4,7,10,13,16,19 22:6n3

Figure 4.1 Structures of selected polyunsaturated fatty acids, arachidonic acid, eicosapentaenoic acid, docosapentaenoic acid, and docosahexaenoic acid.

ACP Domains of PUFA Synthases

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of PUFAs by total chemical synthesis is not yet a viable option, microbial fermentation offers an attractive alternative. The recently discovered genes and gene clusters encoding PUFA biosynthesis are sure to hasten the development of microbial sources of PUFAs, although much remains unknown about PUFA biosynthesis in microorganisms (Metz et al., 2001; Okuyama et al., 2007; Warude et al., 2006). PUFAs differ from other fatty acids in that they contain more than one double bond. The PUFA olefins, most commonly in the cis configuration, are at three-carbon intervals, with each double bond separated by a methylene group (Huang et al., 2004). Olefin positions are indicated by D and the number of the first carbon atom, bearing in mind that the carboxylic acid moiety contains C1; the most commonly used nomenclature starts with a number indicating the PUFA carbon chain length (Wallis et al., 2002). Thus, using this naming convention, AA is termed 20:4D5,8,11,14. More commonly, PUFAs are named in a fashion whereby the carbon count is followed by the number of double bonds and the position of the double bond closest to the terminal methyl group (o carbon) (Wallis et al., 2002). Using this nomenclature, AA is called 20:4n6. Similarly, EPA is 20:5D5,8,11,14,17 or 20:5n3, docosapentaenoic acid (DPA) is 22:5D7,10,13,16,19 or 22:5n3, and DHA is 22:6D4, 7,10,13,16,19 or 22:6n3 (Fig. 4.1). Both aerobic and anaerobic pathways for PUFA biosynthesis are known (Fig. 4.2). Aerobic synthesis of PUFAs utilizes saturated fatty acids as precursors, and desaturases introduce double bonds at specific positions, while elongases extend the resulting precursors in two-carbon increments (Berge´ and Barnathan, 2005; Uttaro, 2006; Warude et al., 2006). The specific set of desaturases and elongases present within any given cell dictates the biosynthetic route exploited and, accordingly, which PUFAs are produced (Uttaro, 2006). As depicted in Fig. 4.2, route A is believed to operate in marine primary producers of PUFAs, initiating the food chain of ‘‘oceanic’’ PUFAs that end in large carnivorous fish, currently the principal source of PUFAs, while route B is typical of mammalian cells (Berge´ and Barnathan, 2005). Mammals lack D12 and n3 desaturase activities and must therefore obtain linoleic acid (18:2n6) and a-linolenic acid (18:3n3) from their diets. Both 18:2n6 and 18:3n3 are produced in plants; conversion of 18:0 to 18:1n9 by D9 desaturase and subsequent processing by D12 desaturase affords 18:2n6 that either can be acted upon by D6 desaturase to afford 18:3n6 or by n3 desaturase to afford a-linolenic acid 18:3n3 (Berge´ and Barnathan, 2005). Aerobic biosynthesis of PUFAs has been rigorously evaluated, and interested readers are referred to recent reviews (Berge´ and Barnathan, 2005; Warude et al., 2006). Numerous bacterial strains are now also known to produce PUFAs under strictly anaerobic conditions, thus precluding the participation of oxygen-dependent enzymes such as the desaturases noted above

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Acetyl-CoA Malonyl-CoA Aerobic pathway

A

Anaerobic pathway

B

C

Linolenic acid 18:3Δ9,12,15 18:3n3

Linolenic acid 18:3Δ9,12,15 18:3n3

PUFA synthase

Δ6-desaturase Δ6-elongase 20:3Δ8,11,14 20:3n6

20:3Δ8,11,14 20:3n6 Δ5-desaturase

Δ5-elongase Δ4-desaturase DHA 22:6Δ4,7,10,13,16,19 22:6n3

DHA 22:6Δ4,7,10,13,16,19 22:6n3

EPA 20:5Δ5,8,11,14,17 20:5n3

EPA 20:5Δ5,8,11,14,17 20:5n3

EPA 20:5Δ5,8,11,14,17 20:5n3

Δ5-elongase Δ7-elongase Δ6-desaturase

DPA 22:5Δ7,10,13,16,19 22:5n3

Tetracosahexaenoic acid (THA) 24:6Δ6,9,12,15,18,21 24:6n3 b-oxidation DHA 22:6Δ4,7,10,13,16,19 22:6n3

Figure 4.2 Pathways for PUFA biosynthesis in different organisms. Routes A and B represent the aerobic PUFA biosynthetic pathways relying on the action of desaturases (O2-dependent) and elongases; transformations shared by the two paths are shaded. Route A is a direct pathway proposed to operate in many marine primary producers and to initiate the food chain of ‘‘oceanic’’ PUFAs. Route B is typical for mammalian cells and embedded in this route, DHA production from EPA results from two successive elongations, a D6 desaturation and a b-oxidation chain–shortening event. The anaerobic route C exploits a PKS-like PUFA synthase.

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(Pereira et al., 2003). Instead, the anaerobic pathways exploit polyketide synthase (PKS)-like enzymes known as PUFA synthases for de novo synthesis of PUFAs from acyl CoA precursors (Hauvermale et al., 2006; Kaulmann and Hertweck, 2002; Metz et al., 2001; Okuyama et al., 2007). Gene clusters encoding PUFA synthases and associated enzymes have been found in both prokaryotic and eukaryotic marine organisms (Okuyama et al., 2007). Successful expression of the bacterial genes encoding EPA and DHA biosynthesis in heterologous hosts, leading to production of EPA and DHA, represented a landmark finding securing the importance and ubiquity of PUFA synthases in anaerobic PUFA biosynthesis (Okuyama et al., 2007). Thus, the pfa genes responsible for EPA and DHA production were first cloned as a cosmid carrying approximately 38 kb of DNA from Shewanella sp. SCRC2738 (Metz et al., 2001). Although this fragment contained 18 open reading frames, only 5 of them, named pfaA, pfaB, pfaC, pfaD, and pfaE, were subsequently found to be needed for EPA and DHA production in E. coli, thereby establishing the minimal gene set for PUFA biosynthesis. Homologues of the pfa genes have since been found in a myriad of PUFA-producing microorganisms, principally from marine sources (Kaulmann and Hertweck, 2002; Okuyama et al., 2007). While the pfaABCDE genes are both necessary and sufficient for EPA and DHA biosynthesis, subtle differences are apparent across the various PUFA biosynthetic gene clusters, as exemplified by those from Shewanella japonica (an EPA and DPA producer), Moritella marina (a DHA producer) (Orikasa et al., 2006), Photobacterium profundum (an EPA producer) (Sugihara et al., 2008), and the marine protist Schizochytrium (a DHA and DPA producer) (Hauvermale et al., 2006) (Fig. 4.3). Most notable among these differences are (1) the genetic organization of pfaE and homologues within PPTase

KS MAT 6

X

ACP

KR DH

AT

KS CLF DH

DH

ER

Shewanella (EPA/DPA) A PfaB

PfaA

PfaE PPTase

KS MAT 5

X

KS MAT 5

X

ACP

KR DH

PfaC

KS AT

KS CLF

DH

PfaD DH

ER

Moritella (DHA) B PfaE

Orf8

Orf9

ACP

KR DH

AT

Orf10 KS CLF DH

Orf11 DH

ER

Photobacter (EPA) C PfaA KS MAT

9

X

PfaB ACP

KR DH

KS CLF

PfaC AT

ER

PfaD DH

DH ER

Schizochytrium (DHA/DPA) D OrfA

OrfB

OrfC

Figure 4.3 Organization of selected PUFA biosynthetic gene clusters: (A) S. japonica (an EPA and DPA producer); (B) M. marina (a DHA producer); (C) P. profundum (an EPA producer); and (D) the marine protist Schizochytrium (a DHA and DPA producer).

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the clusters and (2) the tandem acyl carrier protein (ACP) domains (ranging from five to nine) encoded by pfaA and homologues. Thus, pfaE or homologues have been localized either within or remote from the PUFA biosynthetic gene clusters, encoding either discrete proteins or a domain fused to other PUFA synthase subunits. That PfaA or homologues known to date all contain tandem ACPs stands in striking contrast to the current paradigm for most fatty acid synthases (FASs) and PKSs that are characterized by a single ACP for each cycle of chain elongation. It is these unprecedented architectural and mechanistic features that have motivated careful dissection of PUFA gene clusters as well as efforts to correlate PUFA synthase functions implied by bioinformatics to bona fide biosynthetic transformations in vitro and in vivo. In spite of the significant progress made recently in our understanding of PUFA synthases in PUFA biosynthesis, the majority of studies focused on cloning and sequencing of relevant PUFA biosynthetic gene clusters. Accordingly, there remains a large gap in our ability to correlate sequence information to experimentally verifiable function. Crucial to the full realization of the potential of microbial production methods for PUFAs is the development of new methods specially designed to understand the intricate workings of PUFA synthases. Toward this end, we have recently expressed the S. japonica pfaABCDE genes in E. coli to produce EPA and DPA ( Jiang et al., 2008). Using the S. japonica genes as a model for PUFA synthases, we conducted in vitro and in vivo characterization of each ACP domain of the PfaA subunit. Specifically, we demonstrated that (1) each of the bioinformatically predicted ACPs can be phosphopantetheinylated by the promiscuous PPTase Svp in vitro; (2) PfaE can efficiently phosphopantetheinylate each of the PfaA-ACPs in vivo; (3) each of the tandem ACPs was functionally equivalent for PUFA biosynthesis, but the number of functional ACPs correlated directly to PUFA titers; and (4) PUFA product distributions were regulated in an ACP-independent fashion ( Jiang et al., 2008). Taken together, these findings provide new insight into the role of ACPs in PUFA synthases, as well as experimental evidence supporting the essential role of PfaE as a PPTase in PUFA biosynthesis. This chapter describes methods and protocols developed for characterizing the tandem ACPs of the Pfa PUFA synthase from S. japonica. They include: (1) production of PUFAs in E. coli via expression of the PUFA synthase genes; (2) mapping of the tandem ACP active sites by site-directed mutagenesis; (3) overproduction of each of the tandem ACPs; (4) overproduction of PPTases; (5) in vivo and in vitro preparation of holo-ACPs; and (6) elucidation of the relationship between PUFA production and the number of active ACPs. The strategies and methods described here should be applicable to other PUFA synthases and are envisioned to help close the knowledge gap currently limiting microbial production of PUFAs via PUFA synthase engineering and heterologous expression.


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2. Methods 2.1. Production of PUFAs in E. coli by expressing the PUFAS genes The genes responsible for EPA and DPA biosynthesis from S. japonica have been previously cloned, sequenced, and analyzed (Weaver et al., 2007). A 39,669-bp DNA fragment harboring the five PUFA synthase genes pfaABCDE was cloned into SuperCos 1 (Stratagene, La Jolla, CA) to afford cosmid construct 3F3. Functional expression of pfaABCDE in E. coli was accomplished by adopting the three-plasmid-expression system based on three compatible vectors (Novagen, Madison, WI) of pETDuet-1 (for pfaA/pfaE), pACYCDuet-1 (for pfaB/pfaD), and pCOLADuet-1 (for pfaC). 1. Design and synthesize PCR primers for the amplification of DNA encoding the target genes. Add restriction sites to the primers to facilitate directional cloning. Use high-fidelity DNA polymerases, and sequence the PCR products to verify amplification fidelity. For large PCR fragments, use PCR to introduce cloning sites but replace the central portion of the target gene with genomic DNA to minimize PCR error. 2. Amplify the pfaE gene by PCR, and clone the PCR product as an NdeI-XhoI fragment into the same sites of pETDuet-1. 3. Amplify the 50 and 30 ends of pfaA by PCR, replace the central portion of the PCR product with an EcoRV-PflMI fragment of the genomic pfaA, and clone the entire pfaA gene as a BbsI-NotI fragment into the NcoI-NotI sites of pETDuet-1 that already contains pfaE (Step 2) to afford the expression plasmid pREZ67 (for pfaA/pfaE). 4. Amplify the pfaB and pfaD genes by PCR, and clone the PCR products as an NcoI-NotI and an NdeI-XhoI fragment, respectively, into the same sites of pACYCDuet-1 to afford the expression plasmid pREZ65 (for pfaB/pfaD). 5. Amplify the 50 and 30 ends of pfaC, replace the central portion of the PCR product with an AvrII fragment of the genomic pfaC, and clone the entire pfaC gene as an NdeI-XhoI fragment into the same sites of pCOLADuet-1 to afford the expression plasmid pREZ71 (for pfaC). 6. Co-introduce the three expression plasmids pREZ67, pREZ65, and pREZ71 into host strain E. coli BLR(DE3) by transformation. Select for transformants on LB agar supplemented with ampicillin (100 mg/ml), chloramphenicol (25 mg/ml), and kanamycin (50 mg/ml). 7. Pick a single colony from the plate, grow a seed inoculum in 3 ml of LB at 37 , transfer 0.5 ml of the seed inoculum to 50 ml of fresh LB medium, and incubate at 20 until the OD600 reaches 0.5. LB medium used for both seed and production culture is supplemented with ampicillin (100 mg/ml), chloramphenicol (25 mg/ml), and kanamycin (50 mg/ml).

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8. Induce PUFA synthase gene expression by adding 1 mM of isopropyl b-D-1-thiogalactopyranoside (IPTG), and incubate the culture at 20 for an additional 12 h. Collect the cell pellet by centrifugation for fatty acid methyl ester (FAME) analysis. 9. Add 2 ml of 1.5 M anhydrous HCl in methanol and 1 ml of toluene to the cell pellet from Step 8, stir the sample at 100 for 2 h, and cool it to room temperature. Collect the organic layer, wash it with 1 ml of saturated brine, and concentrate the sample to dryness by removing the solvent under reduced pressure. Identify and quantify the resulting FAMEs by GC and GC-MS analysis with authentic FAME standards.

2.2. Mapping the active sites of PfaA-ACPs by site-directed mutagenesis A general strategy to investigate the role of each of the ACP domains in PUFA synthase is to mutate, via site-directed mutagenesis, the active site Ser to Ala. The Ser residue of ACP is phosphopantatheinylated by the PPTase and subsequently used to tether the growing PUFA intermediate to the PUFA synthase in PUFA biosynthesis. Directed by bioinformatics, the Ser residue for each of the ACPs can be predicted and mutated to map their roles in PUFA synthase and PUFA biosynthesis. To systematically mutate each of the ACPs individually or combinatorially, two common cloning vectors pETm and pUCm, were first generated by inserting the pET-linker into the BglII-XhoI sites of pET28a and the pUC-linker into the EcoRI-HindIII sites of pUC18, respectively, to facilitate subclonings (Fig. 4.4). Additional modifications to the multicloning sites of pETm and pUCm may be necessary to accommodate the cloning needs for all individual ACPs. Boundaries of individual ACP domains can be determined by sequence analysis. DNA fragments encoding each ACP were cloned from pREZ67 (see Step 3, Section 1) into pETm or pUCm via several subcloning steps. Ser-to-Ala point mutation to each of the ACPs was generated by following the QuikChangeÒ site-directed mutagenesis protocol (Stratagene, La Jolla, CA) with primers summarized in Table 4.1. The resultant ACP mutants were then cloned back into pRE67 to replace the wild-type ACPs via several subcloning steps to afford 16 pREZ67-derived

Figure 4.4 DNA sequences of the pET-linker (A) and pUC-linker (B) in pETm and pUCm for constructing Ser-to-Ala point mutations in each of the PfaA-ACPs.

Table 4.1 PCR primers used for introducing a Ser-to-Ala point mutation into each of the PfaA-ACPs by QuikChange1 site-directed mutagenesis

a b c d

Primer

Sequencea

ACP1mu5b ACP1mu3b ACP5mu5c ACP5mu3c ACP6mu5d ACP6mu3d

50 -GCCGATTTAGGCATCGATGCAATTAAACGCGTTGAAATATTAGGTACT-30 50 -AGTACCTAATATTTCAACGCGTTTAATTGCATCGATGCCTAAATCGGC-30 50 -GCGGATTTAGGCATCGATGCAATTAAACGCGTTGAGATCTACTGACTT-30 50 -AAGTCAGTAGATCTCAACGCGTTTAATTGCATCGATGCCTAAATCCGC-30 50 -GCGGATTTAGGCATCGATGCAATTAAACGCGTTGAAATTTTAGGGACG-30 50 -CGTCCCTAAAATTTCAACGCGTTTAATTGCATCGATGCCTAAATCCGC-30

Codons designed for the Ser-to-Ala mutation are underlined. This pair of primers was used for mutagenizing the ACP1, ACP2, ACP3, and ACP4 domains of PfaA. This pair of primers was used for mutagenizing the ACP5 domain of PfaA. This pair of primers was used for mutagenizing the ACP6 domain of PfaA.

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expression plasmids carrying 16 combinations of Ser-to-Ala point mutations to each of the ACP domains of the PfaA subunit. Representative protocols used to produce the full panel of Ser-to-Ala mutants of ACPs are presented for plasmids pREZ67-1m and pREZ67-2m, in which a Ser-to-Ala point mutation is introduced specifically for ACP1 and ACP2 domains of PfaA, respectively. 2.2.1. Construction of pREZ67-1m [for pfaA-ACP1(S1300A)/pfaE] 1. Clone the 2.3-kb KpnI-BglII fragment of pREZ67 into the same sites of pETm to generate pACP1-5. Clone the 1.1-kb KpnI-EagI fragment of pACP1-5 into the same sites of pETm to generate pACP1. 2. Generate the PfaA-ACP1 (S1300A) mutation by following the QuikChange site-directed mutagenesis protocol using pACP1 as the template and ACP1mu5 and ACP1mu3 as primers (Table 4.1). 3. Add 10 units of DpnI directly to the PCR reaction mixture and digest at 37 for 1 h. Introduce 10 ml of the DpnI digested reaction mixture into E. coli DH5a by transformation. Select for transformants on LB agar supplemented with kanamycin (50 mg/ml). Pick a single colony to confirm the desired mutant construct (pACP1m) by sequencing. 4. Clone the 1.1-kb KpnI-EagI fragment of pACP1m back into the same sites of pACP1-5 to generate pACP1-5-1m. 5. Clone the 2.3-kb KpnI-BglII fragment of pACP1-5-1m back into the same sites of pREZ67 to generate pREZ67-1m. 2.2.2. Construction of pREZ67-2m [for pfaA-ACP2(S1406A)/pfaE] 1. Clone the 1.2-kb EagI-BglII fragment of pACP1-5 into the same sites of pETm to generate pACP2-5. Clone the 0.98-kb EagI-HindIII fragment of pACP2-5 into the same sites of pUCm to generate pUCACP2-4. Clone the 0.28-kb EagI-DraIII fragment of pUCACP2-4 into the same sites of pUCm to generate pUCACP2. 2. Generate the PfaA-ACP2 (S1406A) mutation by following the QuikChange site-directed mutagenesis protocol using pUCACP2 as the template and ACP1mu5 and ACP1mu3 as primers (Table 4.1). 3. Add 10 units of DpnI directly to the PCR reaction mixture and digest at 37 for 1 h. Introduce 10 ml of the DpnI-digested reaction mixture into E. coli DH5a by transformation. Select for transformants on LB agar supplemented with ampicillin (100 mg/ml). Pick a single colony to confirm the desired mutant construct (pUCACP2m) by sequencing. 4. Clone the 0.28-kb EagI-DraIII fragment of pUCACP2m back into the same sites of pUCACP2-4 to generate pUCACP2-4-2m. 5. Clone the 0.98-kb EagI-HindIII fragment of pUCACP2-4-2m back into the same sites of pACP2-5 to generate pACP2-5-2m.

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6. Clone the 1.2-kb EagI-BglII fragment of pACP2-5-2m back into the same sites of pACP1-5 to generate pACP1-5-2m. 7. Clone the 2.3-kb KpnI and BglII fragment of pACP1-5-2m back into the same sites of pREZ67 to generate pREZ67-2m.

2.3. Overproduction of each of the PfaA-ACPs 1. Determine ACP boundaries by DNA sequence analysis. Design and synthesize PCR primers for the amplification of DNA encoding the target ACP domains. Incorporate the BamHI and NotI sites into the primers to facilitate directional cloning of the PCR products into pETDuet-1. 2. Amplify the DNAs encoding the six PfaA-ACPs from cosmid 3F3 using the primers summarized in Table 4.2. Clone the PCR products as BamHI-NotI fragments into the same sites of pETDuet-1 to afford pETDuet-1-based expression constructs. Sequence the constructs to verify PCR amplification fidelity. 3. Alternatively, amplify the DNA encoding PfaA-ACP1 from cosmid 3F3 using primers summarized in Table 4.2. Clone the PCR product as an NdeI-XhoI fragment into the same sites of pET28a to afford the pET28a-based expression constructs. Sequence the constructs to verify PCR amplification fidelity. 4. Introduce the expression constructs into host strain E. coli BL21(DE3) by transformation. Select for transformants on LB agar supplemented with ampicillin (100 mg/ml) for pETDuet-1 based expression constructs or kanamycin (50 mg/ml) for the pET28a based expression construct. 5. Pick a single colony from the plate, inoculate 3 ml of LB medium supplemented with ampicillin (100 mg/ml) or kanamycin (50 mg/ml), and incubate at 37 for 12 h. 6. Transfer 0.5 ml of the above culture to 50 ml of LB medium supplemented with ampicillin (100 mg/ml) or kanamycin (50 mg/ml), and grow cells at 37 until the OD600 reaches 0.5. Induce ACP expression by adding 1 mM IPTG, and continue incubation at 37 for 4 h. 7. Pellet cells by centrifugation at 4 and 9200g for 15 min, resuspend the cell pellet in 15 ml of buffer (50 mM Tris HCl, pH 8.0, 300 mM NaCl, 10 mM imidazole), add 1 mg of lysozyme, and keep cells on ice for 1 h. 8. Lyse cells by sonication on ice, centrifuge at 15,000 rpm for 1 h at 4 in a Beckman J2-HS centrifuge equipped with a JA-25.50 rotor (Beckman, Fullerton, CA), and collect the supernatant. 9. Add 2 ml of Ni-NTA agarose (Qiagen, Valencia, CA) to the supernatant, gently shake on ice for 1 h to allow the His6-tagged PfaA-ACPs to bind the Ni-NTA agarose, and load the Ni-NTA agarose-containing

Table 4.2 Primers used for PCR amplification of each of the pfaA-ACPs as well as pfaE from cosmid 3F3 Primer

Forward

Reverse

a b

Sequence

ACP1a ACP1b ACP2 ACP3 ACP4 ACP5 ACP6 pfaE ACP1a ACP1b ACP2 ACP3 ACP4 ACP5 ACP6 pfaE

For amplifying pfaA-ACP1 in Step 2 (Section 3). For amplifying pfaA-ACP1 in Step 3 (Section 3).

50 -GCGGGATCCAACAGCCCTGAGCTCACAAAA-30 50 -AGCCATATGACAGCCCTGAGCTCACAA-30 50 -GCGGGATCCATCAGGTCTTAGCGCAGAAAC-30 50 -GCGGGATCCATCTGGCCTTAGCGCTGAAAC-30 50 -GCGGGATCCATCTGGTCTTAGCGCAGAAAC-30 50 -GCGGGATCCATCAGGTTTAAGTGCGGAACA-30 50 -GCGGGATCCAACAGCCCTGAGCGCTGAGCA-30 50 -CAAGCGCATATGTCTTATTGCTATTATAAA-30 50 -CGCGCGGCCGCTTAGCCTGCGGCCGGTAGTTTA-30 50 -GGCCTCGAGTTATGCATTTGCAGTGTCGCT-30 50 -CGCGCGGCCGCTTAGCCTGCGGCGGGTAGTTTA-30 50 -CGCGCGGCCGCTTAACCAGCAGCGGGTAGCTTA-30 50 -CGCGCGGCCGCTTAGCCTGCGGCGGGTAGTTTA-30 50 -CGCGCGGCCGCTTAGCCTGCGGCGGGTAGTTTA-30 50 -CGCGCGGCCGCTTAGCCTGCGGCTGGCAGTTTA-30 50 -AGACTCGAGTCAGTTGGTTTTTATGAACATTT-30

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solution onto a column. Wash the Ni-NTA agarose column with 30 ml of the buffer (50 mM Tris HCl, pH 8.0, 300 mM NaCl, 20 mM imidazole). 10. Elute the His6-tagged PfaA-ACPs with 2 ml of buffer (50 mM Tris HCl, pH 8.0, 300 mM NaCl, 300 mM imidazole), and dialyze ACP fractions against 250 ml of buffer (25 mM Tris HCl, pH 8.0, 25 mM NaCl, 10% glycerol). 11. Assess purity of the purified PfaA-ACPs by 15% SDS-PAGE (Fig. 4.5A). 12. Characterize the resultant PfaA-ACPs in apo-form by reverse phase HPLC analysis using a Jupiter C-18 column (5 mm, 300A˚, 250 4.6 mm, Phenomenex, Belmont, CA), and a gradient from 15 to 90% CH3CN in 0.1% TFA-H2O over 30 min at a flow rate of 1.0 ml/min, and UV detection at 220 nm (Fig. 4.5B). 13. Collect individual protein peaks from HPLC, lyophilize and characterize the resultant PfaA-ACPs in apo-form by ESI-MS analysis (Agilent 1000 HPLC-MSD SL instrument, Palo Alto, CA) (Table 4.3).

2.4. Overproduction of PfaE and Svp PPTases 1. Amplify the pfaE gene by PCR from cosmid 3F3 using the primers summarized in Table 4.2. Clone the PCR product as an NdeI-XhoI fragment into the same sites of pET28a to afford the expression construct. A kDa 50 40 30 25 20 15 10

Lane 1 2

B mAU at 220 nm 800 0

800

0

800

C I

kDa 50 40

II

30

III

20

0 10

12 14 Retention time (min)

1

2

Lane 3

4

15

Figure 4.5 Analysis of PfaA-ACPs and PfaE PPTase, and HPLC analysis of in vivo and in vitro phosphopantatheinylation of PfaA-ACP1. (A) 15% SDS-PAGE analysis of purified PfaA-ACP1: lane 1, protein markers, and lane 2, PfaA-ACP1 (100 amino acids, calculated mass: 10,896 Da). (B) HPLC analysis of PfaA-ACP1: (I) PfaA-ACP1 overproduced in E. coli; (II) PfaA-ACP1 overproduced in E. coli coexpressing pfaE; and (III) apo-PfaA-ACP1 incubated with Svp in vitro. (C) 15% SDS-PAGE analysis of purified PfaE: lane 1, protein markers; lane 2, total PfaE; lane 3, insoluble PfaE; and lane 4, soluble PfaE. ○, apo-PfaA-ACP1, , holo-PfaA-ACP1.

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Table 4.3 ESI-MS analysis of apo- and holo-forms of PfaA-ACPs examined Apo-form

ACP (# amino acids) c

ACP1 (116) ACP1 (100)d ACP2 (100) ACP3 (100) ACP4 (100) ACP5 (100) ACP6 (100) a b c d

Holo-form

Calculated

Found

Calculated

Found upon in vitro assaya

12454 10896 10781 10882 10869 10799 10797

12450 10894 10778 10879 10872 10802 10795

12794 11236 11121 11222 11207 11139 11137

12790 ND ND ND ND ND ND

Found upon in vivo assayb

ND 11232 11117 11218 11205 11134 11133

Phosphopantetheinylation of PfaA–ACP1 catalyzed by Svp PPTase in vitro. Phosphopantetheinylation of PfaA–ACPs by coexpression of pfaA-ACPs with pfaE in vivo. PfaA–ACP1 overproduced from the expression construct in Step 3 (Section 3). PfaA–ACP1 overproduced from the expression construct in Step 2 (Section 3).

2. Follow Steps 4 to 8 in Section 2.3, and vary fermentation temperature between 18 and 37 and IPTG concentration from 1 mM and 1 mM to optimize production of soluble PfaE. Follow Steps 9 to 11 in Section 2.3 to purify PfaE. Under the conditions examined, the His6-tagged PfaE was overproduced predominantly as an insoluble protein (Fig. 4.5C). 3. Follow the literature procedure to overproduce Svp as a soluble protein, purify it to homogeneity and use it as a promiscuous PPTase for in vitro phosphopantetheinylation of apo-PfaA-ACPs to generate holo-PfaAACPs (Sanchez et al., 2001).

2.5. In vivo and in vitro preparation of the holo-form of PfaA-ACPs 2.5.1. In vivo phosphopantetheinylation of PfaA-ACPs from apo- into holo-form by coexpression of pfaA-ACPs with pfaE 1. Clone pfaE as an NdeI-XhoI fragment (Step 2, Section 2.1) into the same sites of the pETDuet-1–based pfaA-ACP expression constructs (Step 2, Section 2.3) to afford expression constructs for each of the pfaA-ACPs coexpressed with pfaE. 2. Follow Steps 4 to 13 in Section 2.3 for expression, overproduction, purification, HPLC (Fig. 4.5C) and ESI-MS analysis (Table 4.3) of the resultant PfaA-ACPs in holoform.

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2.5.2. In vitro phosphopantetheinylation of PfaA-ACP1 from apo-into holo-form by Svp 1. Prepare a typical in vitro phosphopantetheinylation reaction in 100 ml containing 100 mM apo-ACP, 500 mM CoA, 5 mM Svp, 12.5 mM MgCl2, and 2.5 mM DTT in 100 mM Tris HCl, pH 7.5. Incubate reactions at 25 for 30 min. 2. Analyze the conversion of apo- to holo-ACP by HPLC (Fig. 4.5B) and ESI-MS analysis (Table 4.3) as described in Steps 12 and 13 (Section 2.3).

2.6. Elucidation of the relationship between PUFA production and the number of active ACPs 1. Co-introduce each of the 16 pREZ67-derived expression plasmids (Section 2) with pREZ65 (Step 4, Section 2.1) and pREZ71 (Step 5, Section 2.1) into host strain E. coli BLR(DE3) by transformation to investigate PUFA titers resulting from different combinations of Ser-to-Ala point mutations to each of the ACP domains of the PfaA subunit (Section 2.2). Select for transformants on LB agar supplemented with ampicillin (100 mg/ml), chloramphenicol (25 mg/ml), and kanamycin (50 mg/ml). 2. Follow Steps 7 to 9 (Section 2.1) for pfa gene expression, PUFA production, and FAME preparation and analysis to correlate PUFA titers to number and relative location of active ACP domains within the PfaA mutant (Section 2.2). Representative data are shown in Table 4.4 and Fig. 4.6.

3. Conclusion Traditional sources of PUFAs (predominantly oceanic fish oils) are now widely recognized as being unsustainable due to global warming, pollution of the marine environment, overfishing, and a rapidly growing global population, among other problems (Graham et al., 2007). The discovery of PUFA production and subsequent cloning and sequencing of gene clusters encoding PUFA biosynthesis in a wide array of marine organisms have caused significant excitement for PUFA production via PUFA synthase expression and engineering (Wallis et al., 2002; Kaulmann and Hertweck, 2002; Berge´ and Barnathan, 2005). Key advantages for such an alternative source of PUFAs include (1) simpler PUFA profile (often one predominant product from microbial sources) rather than a complex

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Table 4.4 PUFA titers from recombinant strains harboring pfa PUFA synthase gene expression constructs that carry varying numbers of Ser-to-Ala point mutations in the ACP domains of the PfaA subunit

Wildtype PfaA Mutations to selected one of the six ACPs Mutations to selected two of the six ACPs Mutations to selected three of the six ACPs Mutations to selected four of the six ACPs Mutations to selected five of the six ACPs Mutations to all six ACPs a

No. of functional ACPs

% EPAa

% DPAa

6 5

8.9 0.3 7.7 0.8

1.4 0.1 1.3 0.2

4

8.2 0.3

1.2 0.1

3

6.6 0.3

0.7 0.1

2

5.4 0.2

0.6 0.1

1

3.3 0.5

0.3 0.1

0

0.0

0.0

Percent EPA and percent DPA values are percentages relative to total fatty acid content as determined by the FAME method.

% EPA production

8.0

6 4

6.0

3 2

2

5

0

1

3

4

4.0 1

2.0

0.0

0

0

5

Active ACP Inactive ACP

6

1

2 3 4 5 Number of active ACPs

6

Figure 4.6 Relationship between percent EPA yield (relative to total amount of fatty acids) and number of active ACP domains within PfaA. The horizontal axis indicates number of active, nonmutagenized, ACPs. The vertical axis indicates percent EPA production observed.


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mixture currently obtained from fish or algal oils, thereby significantly reducing the expense associated with preparative PUFA purification, and (2) biotechnological opportunities to investigate and exploit, via heterologous expression and engineering strategies, the PUFA synthase genes for large-scale production by fermentation, titer improvement, and engineered production of specific PUFAs. The past few years have witnessed exponential growth in discovering PUFA biosynthetic gene clusters (Okuyama et al., 2007). Although it is now well accepted that PUFA synthase-catalyzed de novo biosynthesis of PUFAs from acyl CoA precursors is widespread, our understanding of the enzymology and biochemistry of PUFA synthases remains rudimentary, requiring the development and application of new research methods for mechanistic characterization. The potential of engineering PUFA synthase genes for PUFA production remains, to date, largely untapped. The methods and protocols described here have been instrumental in elucidating key elements of PUFA biosynthesis in S. japonica, a microbial EPA and DPA producer that we have studied as a model for PUFA synthase and PUFA biosynthesis. These studies have allowed us to conclude that each of the bioinformatically predicted tandem ACPs is phosphopantetheinylated and functionally equivalent but that the number of functional ACPs correlates directly to PUFA yields ( Jiang et al., 2008). We have further demonstrated that our finding on the roles of tandem ACPs from PfaA most likely are general for other PUFA synthases by constructing a hybrid PUFA synthase with subunits from both the S. japonica (an EPA and DPA producer) and M. marina (a DHA producer) PUFA synthases (Fig. 4.3). Examination of the PUFA profiles from the recombinant E. coli strains carrying the hybrid PUFA synthase led us to the conclusion that, while the overall PUFA titer directly depends on the total number of active ACPs, the determinant(s) of the final PUFA products is in contrast ACP-independent ( Jiang et al., 2008). These studies set the stage to interrogate other domains and subunits of PUFA synthase for their roles in controlling the final PUFA products and to engineer the PUFA biosynthetic machinery for improving production. Central to future engineering efforts is a continued and rigorous dissection of PUFA synthases from a broader selection of microbial PUFA producers. The ubiquity of tandem ACP domains among PUFA synthases as shown by bioinformatics, combined with the clear need to develop new PUFA sources, hints at the importance of enzymology and biochemistry experiments sure to benefit from the methods and protocols described herein.

ACKNOWLEDGEMENT This work was supported in part by a grant from Martek Biosciences Co., Columbia, MD.

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REFERENCES Berge´, J.-P., and Barnathan, G. (2005). Fatty acids from lipids of marine organisms: Molecular biodiversity, roles as biomarkers, biologically active compounds, and economical aspects. Adv. Biochem. Eng. Biotechnol. 96, 49–125. Graham, I. A., Larson, T., and Napier, J. A. (2007). Rational metabolic engineering of transgenic plants for biosynthesis of omega-3-polyunsaturates. Curr. Opinion Biotechnol. 18, 142–147. Hauvermale, A., Kuner, J., Rosenzweig, B., Guerra, D., Diltz, S., and Metz, J. G. (2006). Fatty acid production in Schizochytrium sp.: IInvolvement of a polyunsaturated fatty acid synthase and a type I fatty acid synthase. Lipids 41, 739–747. Huang, Y.-S., Pereira, S. L., and Leonard, A. E. (2004). Enzymes for transgenic biosynthesis of long-chain polyunsaturated fatty acids. Biochimie 86, 793–798. Jiang, H., Zirkle, R., Metz, J. G., Braun, L., Richter, L., Van Lanen, S. G., and Shen, B. (2008). The role of tandem acyl carrier protein domains in polyunsaturated fatty acid biosynthesis. J. Am. Chem. Soc. 130, 6336–6337. Kaulmann, U., and Hertweck, C. (2002). Biosynthesis of polyunsaturated fatty acids by polyketide synthases. Angew. Chem. Int. Ed. Engl. 41, 1866–1869. Metz, J. G., Roessler, P., Facciotti, D., Levering, C., Dittrich, F., Lassner, M., Valentine, R., Lardizabal, K., Domergue, F., Yamada, A., Yazawa, K., Knauf, V., et al. (2001). Production of polyunsaturated fatty acids by polyketide synthases in both prokaryotes and eukaryotes. Science 293, 290–293. Muskiet, F. A., and Kemperman, R. F. (2006). Folate and long-chain polyunsaturated fatty acids in psychiatric disease. J. Nutr. Biochem. 17, 717–727. Okuyama, H., Orikasa, Y., Nishida, T., Watanabe, K., and Morita, N. (2007). Bacterial genes responsible for the biosynthesis of eicosapentaenoic and docosahexaenoic acids and their heterologous expression. Appl. Environ. Microbiol. 73, 665–670. Orikasa, Y., Nishida, T., Hase, A., Watanabe, K., Morita, N., and Okuyama, H. (2006). A phosphopantetheinyl transferase gene essential for biosynthesis of n–3 polyunsaturated fatty acids from Moritella marina strain MP-1. FEBS Lett. 580, 4423–4429. Pereira, S. L., Leonard, A. E., and Mukerji, P. (2003). Recent advances in the study of fatty acid desaturases from animals and lower eukaryotes. Prostaglandins Leukot. Essent. Fatty Acids 68, 97–106. Sanchez, E. L., Du, L. C., Edwards, D. J., Toney, M. D., and Shen, B. (2001). Cloning and characterization of a phosphopantetheinyl transferase from Streptomyces verticillus ATCC15003, the producer of the hybrid peptide-polyketide antitumor drug bleomycin. Chem. Biol. 8, 725–738. Sugihara, S., Orikasa, Y., and Okuyama, H. (2008). An EntD-like phosphopantetheinyl transferase gene from Photobacterium profundum SS9 complements pfa genes of Moritella marina strain MP-1 involved in biosynthesis of docosahexaenoic acid. Biotechnol. Lett. 30, 411–414. Uttaro, A. D. (2006). Biosynthesis of polyunsaturated fatty acids in lower eukaryotes. IUBMG Life 58, 563–571. Wallis, J. G., Watts, J. L., and Browse, J. (2002). Polyunsaturated fatty acid synthesis: What will they think of next? Trends Biochem. Sci. 27, 467–473. Warude, D., Joshi, K., and Harsulkar, A. (2006). Polyunsaturated fatty acids: Biotechnology. Crit. Rev. Biotechnol. 26, 83–93. Weaver, C. A., Zirkle, R., and Metz, J. G. (2007). PUFA polyketide synthase systems and uses thereof. U.S. Patent 7217856.

C H A P T E R

F I V E

Iterative Type I Polyketide Synthases for Enediyne Core Biosynthesis Geoffrey P. Horsman,* Steven G. Van Lanen,† and Ben Shen*,‡ Contents 1. Introduction 2. Methods 2.1. PCR amplification of PKSE cassettes for predictive classification of new enediynes 2.2. Heterologous expression and overproduction of PKSE proteins 2.3. Production and isolation of the polyene intermediate from 9-membered PKSEs 2.4. Production of apo-ACPs from PKSE for in vitro functional analyses 2.5. In vitro preparation of holo-ACPs 3. Conclusion Acknowledgment References

98 101 101 103 105 107 108 109 111 111

Abstract Enediyne natural products are extremely potent antitumor antibiotics with a remarkable core structure consisting of two acetylenic groups conjugated to a double bond within either a 9- or 10-membered ring. Biosynthesis of this fascinating scaffold is catalyzed in part by an unusual iterative type I polyketide synthase, PKSE, that is shared among all enediyne biosynthetic pathways whose gene clusters have been sequenced to date. The PKSE is unusual in two main respects: (1) it contains an acyl carrier protein (ACP) domain with no sequence homology to any known proteins, and (2) it is self-phosphopantetheinylated by an integrated phosphopantetheinyl transferase (PPTase) domain. The unusual domain architecture and biochemistry of the PKSE hold promise both for the rapid identification of new enediyne natural products and for obtaining fundamental catalytic insights into enediyne biosynthesis. This chapter describes methods for rapid PCR-based classification of conserved enediyne biosynthetic genes, * {

{

Division of Pharmaceutical Sciences, University of Wisconsin-Madison, Madison, Wisconsin, USA Division of Pharmaceutical Sciences, College of Pharmacy, University of Kentucky, Lexington, Kentucky, USA Department of Chemistry, University of Wisconsin-Madison, Madison, Wisconsin, USA


#


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heterologous production of 9-membered PKSE proteins and isolation of the resulting polyene product, and in vitro characterization of the PKSE ACP domain.

1. Introduction The enediyne family of natural products are among the most potent anticancer drugs ever discovered (Wang and Xie, 1999). A neocarzinostatinpolymer conjugate SMANCSÒ (Maeda, 2001) and calicheamicin-antibody conjugate MylotargÒ (Sievers and Linenberger, 2001) are used clinically in Japan and the United States, respectively. While their extraordinary biological activity generates interest among clinicians, the unprecedented molecular architecture of the enediynes motivates biochemical studies to decipher the biosynthetic logic of their assembly. The structurally remarkable enediyne core, or so-called ‘‘warhead,’’ consists of two acetylenic groups conjugated to a double bond within either a 9- or 10-membered ring exemplified by C-1027 and calicheamicin, respectively (Fig. 5.1). Cytotoxicity arises from environmentally triggered cyclization of the enediyne core to yield a benzenoid diradical capable of abstracting hydrogen from DNA (Galm et al., 2005). Although elucidation of the biosynthetic pathways for the peripheral moieties has progressed rapidly in recent years, little is known about the assembly of the enediyne core itself, and fascinating enzymology surely awaits discovery (Van Lanen and Shen, 2008). Such detailed biochemical knowledge is required to generate improved enediyne analogues via rational manipulation of the biosynthetic machinery (Kennedy et al., 2007a,b). A de novo biosynthetic pathway involving a dedicated polyketide synthase (PKS) has only recently been shown to be responsible for production of the enediyne core. Although early studies employing isotope-labeling experiments established acetate as a precursor (Hensens et al., 1989; Lam et al., 1993; OCH3

O

SSSCH3 O

N H

O

O H3C (CH3)2N

O

CH3 OH OH

HO

O

O

O

O O O

O

O

NHCO2 CH3 CH3 N O OH H

OH O

CH3 S

OCH3

H3C

OCH3 NHEt

OCH3

I

OH O

O H3C HO

Cl

CH3O

NH2

C-1027

O

O OH

Calicheamicin

Figure 5.1 Structures of representative members of the 9-membered (C-1027) and 10-membered (calicheamicin) enediyne natural products.

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Tokiwa et al., 1992), these experiments could not distinguish whether the enediyne core originated from a dedicated PKS or from the degradation of a fatty acid precursor. However, five enediyne biosynthetic gene clusters— those for C-1027 (Liu et al., 2002), neocarzinostatin (Liu et al., 2005), maduropeptin (Van Lanen et al., 2007), calicheamicin (Ahlert et al., 2002), and dynemicin (Gao and Thorson, 2008)—have now been sequenced and each has been shown to possess a unique iterative type I PKS (PKSE) sharing significant sequence homology and domain architecture. The essential role of the PKSE was established when C-1027 production was abolished by disruption of the sgcE gene (i.e., the pksE gene from the C-1027 cluster) and restored upon complementation of the △sgcE mutation by expressing a functional copy of sgcE in trans (Liu et al., 2002). Similar results were obtained using the PKSE from the neocarzinostatin cluster, NcsE (Liu et al., 2005), and the maduropeptin cluster, MdpE (Van Lanen et al., 2007). Together, these results definitively establish the polyketide origin of the enediyne core. The unique, enediyne-specific pksE gene cassettes may be exploited to identify new enediyne biosynthetic gene clusters. Indeed, a universal PCR-based method has been developed whereby degenerate primers from conserved PKSE regions are used to rapidly amplify and clone additional pksE genes (Liu et al., 2003). Sequencing and phylogenetic analysis of the resulting minimal enediyne cassettes has revealed a clear genotypic distinction between 9- and 10-member-specific PKSEs. This distinction significantly aids prediction of unknown enediyne core structures directly from the pksE gene sequence. Thus, rapid PCR-based enediyne genotyping of new enediyne-producing isolates may direct the discovery of new members of the enediyne family of natural products. The availability of multiple pksE sequences has enabled bioinformatics analysis, which unambiguously identified four domains: a ketosynthase (KS), acyltransferase (AT), ketoreductase (KR), and dehydratase (DH). In both domain organization and sequence homology, PKSEs are most closely related to the polyunsaturated fatty acid (PUFA) synthases involved in the biosynthesis of docosahexaenoic acid in Moritella marina and eicosapentaenoic acid in Shewanella japonica ( Jiang et al., 2008; Metz et al., 2001) (Fig. 5.2A). (See Chapter 4 in this volume.) However, two unusual features distinguish the PKSEs as an unusual PKS family unique to enediyne biosynthesis. First, a region between the AT and KR domains was proposed to be an acyl carrier protein (ACP) based on identical architecture to the PUFA synthases (Fig. 5.2A), even though this region has no homology to any known proteins. Second, the C-terminal region was predicted to be a phosphopantetheinyl transferase (PPTase) capable of loading the phosphopantetheine (Ppant) cofactor onto the ACP (Zazopoulos et al., 2003). In contrast to typical ACP loading by a discrete PPTase, such selfphosphopantetheinylation is extremely rare (Fichtlscherer et al., 2000; Weissman et al., 2004). The above-described in vivo complementation

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A PUFA synthase (∼2500 aa)

KS

AT

PKSE (∼1900 aa)

KS

AT

Multiple ACPs

ACP

KR

DH

KR

PPTase

9-membered specific accessory enzymes

B

DH

9-membered enediynes

O SCoA

O

ACP

PKSE

+

S

O

−

SCoA

O (7x)

–CO2 (7x)

R O

polyketide intermediate

10-membered enediynes

10-membered specific accessory enzymes

C

E10 (TE)

ACP

S

R

−

O O

OH CO2 + H2O

O Polyketide intermediate

1

3

5

7

9

11

13

15

Figure 5.2 PKSE domain organization and role in enediyne core biosynthesis. (A) Domain organization of PKSE is similar to that of the PUFA synthases. (B) Enediyne core biosynthesis carried out by PKSE and accessory enzymes is responsible for divergence of the polyketide intermediate into either 9- or 10-membered enediyne families of natural products. (C) Structure of the polyene product 1,3,5,7,9,11,13-pentadecaheptanene isolated upon coexpression of 9-membered pksE and pksE10 exemplified by sgcE/sgcE10 and ncsE/ncsE10.

system was used to demonstrate that Ser974Ala and Asp1827Ala variants of SgcE failed to restore C-1027 production in the △sgcE mutant (Zhang et al., 2008), consistent with the proposed roles of Ser974 as a site for Ppant modification on the ACP and Asp1827 as a PPTase catalytic residue. Although rapid genotyping and in vivo complementation revealed a domain architecture that is unique to the PKSEs, in vitro functional characterization is required to unambiguously identify the ACP and PPTase domains. Recent methods have been developed to biochemically characterize PKSEs, providing fascinating insights into their function (Zhang et al., 2008). For instance, the development of a heterologous expression system for 9-membered PKSEs led to the isolation of a polyene compound as the first

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isolable intermediate en route to enediyne core biosynthesis (Fig. 5.2B and C), and this intermediate could not be isolated from the aforementioned Ser974Ala or Asp1827Ala mutants of the ACP or PPTase, respectively. The roles of the ACP and PPTase domains were then confirmed by two crucial in vitro biochemical experiments employing both the wildtype SgcE and the respective Ser974Ala and Asp1827Ala site-directed mutants. First, peptide mapping of SgcE by Fourier-transform mass spectrometry clearly identified a þ340 amu mass shift associated with Ser974 of the ACP domain. Consistent with Ppant modification, the mass shift disappeared upon mutation of either the Ser974 of the ACP or the Asp1827 of the PPTase. Second, phosphopantetheinylation was definitively demonstrated by incubation of the ACP domain with the exogenous PPTase Svp (Sanchez et al., 2001). The conversion of apo-ACP to holo-ACP could be monitored by HPLC, and the substrate and product of this reaction could be isolated and identified by mass spectrometry. Moreover, the PPTase from the calicheamicin pathway was expressed and in vitro phosphopantetheinylation of several carrier proteins was demonstrated (Murugan and Liang, 2008). Together, these results establish PKSE as an iterative type I PKS of unusual domain architecture that self-phosphopantetheinylates a novel ACP domain. This chapter describes the protocols for identification and characterization of the iterative type I PKS for enediyne core biosynthesis (PKSEs): (1) PCR amplification of pksE cassettes for predictive classification of new enediynes; (2) heterologous expression and purification of 9-memberedspecific PKSE proteins in Escherichia coli; (3) production and isolation of the polyene intermediate produced by the 9-membered PKSEs; (4) heterologous expression and purification of 9-membered-specific ACP domains; and (5) their in vitro phosphopantetheinylation.

2. Methods 2.1. PCR amplification of PKSE cassettes for predictive classification of new enediynes 2.1.1. Primer design 1. Conserved regions of shared genes involved in enediyne core biosynthesis (including the PKSE) from the loci for C-1027, neocarzinostatin, and calicheamicin were used to construct degenerate primers (Fig. 5.3 and Table 5.1) (Liu et al., 2003). Primer pairs A–E and A–F, respectively, amplify 1.4- and 3.8-kb fragments of the N-terminal PKSE gene, whereas primer pairs C-H and D-H respectively amplify 2.9- and 0.9-kb fragments of the C-terminal region. The B–G primer pair is

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A

A

C

B E

0

1

2

3

4

5

F 6

7

NcsE3

NcsE4

NcsE5

NcsE

SgcE3

SgcE4

SgcE5

SgcE

CalU5

CalU4

CalT3

CalE8

D H

G 8

9

10

11

12

13 kb

NcsE10

NCS SgcE10

C-1027 CalE7

CAL

B

Figure 5.3 Degenerate primer design for PCR-based amplification of PKSE fragments. (A) Location of primers in relation to the pksE and associated genes that together constitute a minimal pksE cassette from the neocarzinostatin (NCS), C-1027, and calicheamicin (CAL) biosynthetic clusters. (B) The conserved amino acid motifs used to design degenerate primers A to H. Numbers between motifs represent amino acid distance, and the noncoding regions between open reading frames (108 CFU/ml) in a 250-ml baffled shakeflask, and incubate for 14 to 16 h at 28 with shaking at 250 rpm.

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2. Use 3 ml of the resulting culture to inoculate 100 ml of semidefined SAO-23 medium (Sekurova et al., 1999) in a 500-ml baffled shake-flask and incubate at 28 for 120 h with shaking (200 rpm, orbital movement amplitude 2.5 cm). To obtain optimal morphology, add 3 g of 2-mm glass beads to each flask. 3. Extract 1ml culture with 10 ml dimethyl sulfoxide (DMSO). 4. Perform LC-MS analysis of the DMSO extract on an Agilent 1100 HPLC connected to an Agilent MSD TOF mass spectrometer with electrospray ionization (ESI) in both positive and negative ion mode using the HPLC column Waters NovaPak C18 (150 2.1 mm) at a flow rate of 300 ml/min. The mobile phase is 10 mM ammonium acetate (pH 4.0) and acetonitrile (ACN). 5. Run the LC with a linear gradient from 30 to 70% ACN in 10 mM ammonium acetate (pH 4.0) for the first 10 min and then keep at 70% ACN for 5 min (Bruheim et al., 2004).

4.2. Scaled-up production of nystatin analogues 1. Fermentations are performed in 3-l fermentors with an initial medium (SAO-23) volume of 1 to 1.5 l, and an inoculum of 3% (v/v) grown in liquid 0.5 TSB medium in shake-flasks at 28 for 18 h. 2. Fermentations are run at 28 with constant aeration (0.25 VVM) and agitation is controlled to maintain the dissolved oxygen level higher than 30% saturation. The pH is maintained at 6.5 to 7.0 by addition of 2 M NaOH. 3. Polyene production is assayed by DMSO extraction of cultures and subsequent analysis by LC-MS on an Agilent 1100 HPLC system with an online diode array detector (DAD)—and a TOF mass-sensitive detector and electrospray ionization in the negative mode, as described in Bruheim et al., 2004.

4.3. Preparative LC-MS purification of nystatin analogues 1. For preparative isolation, polyenes are extracted from mycelium with 5 ml of MeOH per gram of cell pellet. The extract can be subjected to preparative HPLC either directly, or after concentration. Concentration is performed by adding 2.5 ml water to 5 ml extract, removing the liquid by vacuum evaporation until precipitation occurs, drying the precipitated analogue, and dissolving it in DMSO to a concentration of 1 to 2 mg/ml. 2. Subject the samples to preparative HPLC on the Agilent 1100 system with an online diode array detector (DAD)–Agilent MSD Trap mass spectrometer monitoring with flow split to the MSD and a fraction collector. A preparative Waters NovaPak C18 column (30 300 mm)

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is used at 30 with a flow rate of 50 to 60 ml/min, using 0.5- to 2-ml injection volume. 3. Run the LC with a linear gradient from 20 to 70% MeOH or ACN in 10 mM ammonium acetate pH 4.0 for the first 10 to 15 min and then keep at 70 to 80% for 2 to 5 min (Borgos et al., 2006b; Bruheim et al., 2004). 4. The collected fractions are pooled, and an aliquot is reanalyzed by analytical LC-MS to determine the purity of the isolated compound. The purity and concentration of each engineered polyene are determined by reference to USP standards of nystatin and amphotericin B for tetraenes and heptaenes, respectively, assuming that molar extinction coefficients in the spectral regions of interest are unaltered. Peak UV absorptions at 309 nm and 386 nm, are used for tetraenes and heptaenes, respectively. Typically, 95 to 97% purity is achieved. 5. The rest of the sample is dried in a Speed-Vac centrifuge after the mobile phase was exchanged for a water-MeOH mixture (10 to 90%) by solidphase extraction (Oasis HLB SPE kit; Waters).

5. Conclusion Genetic manipulation of S. noursei and S. nodosus has yielded insights into polyene biosynthesis. This work has yielded analogues that may eventually become medically important. The ability to engineer genes in these organisms will be important in future mining of their genomes. This work may also assist basic research on homologous recombination in streptomycetes.

ACKNOWLEDGMENTS We thank Ha˚vard Sletta for providing details on production and purification of nystatin analogues.

REFERENCES Allmansberger, R., Brau, B., and Piepersberg, W. (1985). Genes for gentamicin-(3)N-acetyltransferases III and IV. II. Nucleotide sequences of three AAC(3)-III genes and evolutionary aspects. Mol. Gen. Genet. 198, 514–520. Aparicio, J. F., Caffrey, P., Gil, J., and Zotchev, S. B. (2003). Polyene antibiotic biosynthesis gene clusters. Appl. Microbiol. Biotechnol. 61, 179–188. Arendrup, M., Lundgren, B., Mller Jensen, I., Snder Hansen, B., and Frimodt-Mller, N. (2001). Comparison of Etest and a tablet diffusion test with the NCCLS broth microdilution method for fluconazole and amphotericin B susceptibility testing of Candida isolates. J. Antimicrob. Chemother. 47, 521–526.

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C H A P T E R

T W E LV E

Polyketide Versatility in the Biosynthesis of Complex Mycobacterial Cell Wall Lipids Tarun Chopra and Rajesh S. Gokhale Contents 1. 2. 3. 4.

Introduction Acetate and Propionate Feeding Studies Genome Sequencing and Identification of Polyketide Synthases Mycobacterial Polyketide Synthases 4.1. Biosynthesis of dimycocerosate esters (DIMs) by PKS15/1, PpsABCDE, and MAS 4.2. PKS2 is involved in biosynthesis of sulfolipids 4.3. PKS12 uses a novel ‘‘modularly iterative’’ mechanism for biosynthesis of mannosyl-b-1-phosphomycoketides 4.4. PKS13 catalyzes condensation of fatty-acyl chains during biosynthesis of mycolic acids 4.5. PKS3/4 is involved in the biosynthesis of phthenoic acids 4.6. PKS10, PKS7, PKS8, PKS17, PKS9, and PKS11 constitute an unusual PKS cluster 4.7. PKS18 is involved in biosynthesis of long-chain pyrones 4.8. MbtC and MbtD are involved in biosynthesis of iron-chelating siderophores from Mtb 4.9. PKS5 and PKS6 5. Techniques to Characterize PKS Enzymes 5.1. Product formation assay to study activity of PKS enzymes 5.2. Characterization of PKS derived saturated fatty acids References

260 265 266 267 267 270 272 273 277 278 279 280 281 282 282 282 284

Abstract Genome sequencing of Mycobacterium tuberculosis (Mtb) has revealed a large number of open reading frames homologous to polyketide synthases (PKSs). Since Mtb is not known to produce secondary metabolites, their presence in the

Chemical Biology Laboratory, National Institute of Immunology, New Delhi, India Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04612-6

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Mtb genome was rather surprising. Research over the last decade has demonstrated that these PKSs are involved in the biosynthesis and assembly of complex lipids. The catalytic as well as mechanistic versatility of PKSs in producing acyl chains of Mtb lipidic metabolites are discussed here. We refer to the literature for most bench-level details, but include protocols for generally useful methods for analyzing the products of mycobacterial PKSs.

1. Introduction Mycobacterium tuberculosis (Mtb) is the etiologic agent of the human disease tuberculosis and remains a major cause of morbidity and mortality worldwide. Mtb bacilli are encapsulated by a complex cell envelope known to actively contribute toward virulence. This waxy barrier also provides protection to the bacterium from various therapeutic agents (Draper, 1998; Jarlier and Nikaido, 1994). Much of the early work on the chemical nature of the cell wall of Mtb was carried out by R.J. Anderson and T.B. Johnson at Yale University, where they focused on characterization of fat-like material present in the tubercle bacillus. While the initial instinct was for the presence of sterols, examination of both aqueous and solvent fractions did not yield any sterol-containing compounds. Purification and careful analysis of these fractions revealed many new and interesting chemical substances not usually found in the plant and animal kingdoms. These studies provided early leads for the presence of trehalose-containing compounds, phthiocerols, mycolic acids, and so on in mycobacteria (Anderson, 1941; Goren, 1972; Minnikin, 1982). Complete chemical characterization of these complex metabolites was carried out in subsequent years with the advent of new technologies and methodologies. It is now established that these chemical compounds are embedded as a highly complex network of sugars, proteins and lipids in the mycobacterial cell wall (Daffe and Draper, 1998; Lederer et al., 1975). The base of the mycobacterial cell wall consists of a plasma membrane that can be resolved into a thick outer layer and a thin inner layer using electron microscopy, as depicted in Fig. 12.1 (Paul and Beveridge, 1992; Zuber et al., 2008). The thickness of the outer layer is associated with the presence of carbohydrates and phospholipids, including phosphatidylinositol mannosides (PIMs). PIMs are involved in anchoring polysaccharides like lipoarabinomannan (LAM) and lipomannan (LM) in the cell wall (Brennan, 1988; Daffe´ and Lemassu, 2000; Guerardel et al., 2002; Pitarque et al., 2008). The plasma membrane also hosts a variety of other compounds like carotenoids, menaquinones and various glycosylphosphopolyprenols (Brennan and Nikaido, 1995; Daffe´ and Lemassu, 2000). Outside the plasma membrane, a complex polymer of peptidoglycan surrounds the mycobacteria and acts as a scaffold to which arabinogalactan moieties are connected by L-rhamnose-D-Nacetylglucosamine. The C-1 of N-acetylglucosamine is phosphodiesterified

261

PKS-Mediated Biosynthesis of Mycobacterial Lipids

Mycobactin

MPM

Porin HO HO O

O O P O– OH OH O

R

n O

O

OCH3

PDIMs

O O

R HO n N O O O N –OH NH O O R

R

O HN O N HO

Mycolic acids

LM portion of LAM Galactan Arabinose sugar Rhamanose Peptidoglycan PIMs Plasma membrane

Figure 12.1 Schematic representation of the mycobacterial cell envelope. MPM, mannosyl-b-1-phosphomycoketides; PDIM, phthiocerol dimycocerosates; PIM, phosphatidylinositol mannosides; LM, lipomannan; LAM, lipoarabinomannan.

to the hydroxyl moiety of C-6 of the muramic units of the peptidoglycan layer. The rhamnose moiety is connected to the galactan of the arabinoglactan layer, generating a bridge between the arabinogalactan and the peptidoglycan layer. Mycolic acids are esterified to these distal arabinose sugars of the arabinogalactan layer (Brennan, 2003; Brennan and Nikaido, 1995; McNeil et al., 1991; Misaki et al., 1974). Mycolic acids are also present as free mycolates or as esters of trehalose sugars called trehalose monomycolates (TMM) and trehalose dimycolates (TDM) (Bloch and Noll, 1955; Noll et al., 1956; Takayama et al., 2005). A number of other surface-exposed lipids like sulfolipids (SL), polyacyltrehaloses (PAT), phthiocerol dimycocerosates (PDIM), mannosyl-b-1-phosphomycoketides (MPM) and diacyl trehaloses (DAT) intercalate into the cell wall. This lipid-rich cell wall also harbors a number of proteins, including the antigen 85 complex and porins, which constitute the complex cell-wall assembly (Asselineau and Laneelle, 1998; Brennan and Nikaido, 1995; Draper, 1998; Gokhale et al., 2007b). In this article we describe the enzymology of the various components of the mycobacterial envelope, focusing on the role played by polyketide synthases (PKSs) in their biosynthesis. It is not feasible to provide standard protocols for studying all these varied processes. At the end of the article we give two protocols that we have found useful for product analysis, which can be adapted for assaying a wide range of compounds. Various techniques involved in analysis and characterization of mycobacterial enzymes are summarized in Tables 12.1 and 12.2.

262

Table 12.1 Techniques used to study biosynthetic pathways in mycobacteria Technique

Comments

References

Gene inactivation techniques

Gene inactivation along with mouse infection studies and biochemical analysis of mutant strains enables identification of gene function and its importance for virulence Random mutagenesis technique that enables generation of several gene mutants at a time Random mutagenesis technique that enables identification of conditionally essential genes Mutagenesis technique targeted towards a single gene

(Camacho et al., 1999; Hensel et al., 1995) (Murry et al., 2008; Sassetti et al., 2001) (Parish and Stoker, 2000)

1. Signature tagged mutagenesis (STM) 2. Transposon-site hybridization (TraSH) 3. Allelic exchange using suicide vectors 4. Allelic exchange using conditionally replicating vectors

5. Inducible expression of antisense RNA 6. Regulated gene expression by tetracycline inducible promoters Lipid analysis

Targeted mutagenesis technique that possesses high recombination efficiency. Utilizes temperaturesensitive plasmid or plasmid incompatibility delivery systems Conditional mutagenesis technique that enables reduction of target gene expression Conditional mutagenesis technique that enables a tighter regulation of target gene expression

(Guilhot et al., 1992; Pashley et al., 2003)

1. Useful in investigating the importance of the mutant gene in lipid biosynthesis 2. Analysis techniques include TLC and mass spectrometry 3. Different lipids can be analyzed by varying the solvent system used for TLC analysis

(Slayden and Barry, 2000) (Bhatt et al., 2007c; Constant et al., 2002; Kumar et al., 2007; Lea-Smith et al., 2007; Mougous et al., 2002; Waddell et al., 2005)

(Parish and Stoker, 1997) (Carroll et al., 2005)

Table 12.2 Biochemical assays used to characterize enzymes involved in lipid biosynthesis Enzymatic assays

Comments

1. PKS and FAS assays

1. PKSs and FASs are implicated in biosynthesis of most mycobacterial lipids 2. Product characterization using a combination of radio- TLC, radio-HPLC and mass spectrometric approaches

Type I PKSs/FASs Type II PKSs/FASs

Type III PKSs/FASs 2. FAAL/FACL assays

1. FAALs activate starter fatty acids as acyl adenylates for PKS proteins 2. Product characterization using radio-TLC, radio-HPLC and mass spectrometric approaches

3. Polyketide-associated proteins (Pap) A assays

1. PapA proteins demonstrate esterase activity 2. Product characterization using radio-TLC, radio-HPLC and mass spectrometric approaches

References

This article, (Chopra et al., 2008; Trivedi et al., 2005) (Gurvitz et al., 2008a; Kremer et al., 2001; Marrakchi et al., 2002; Sacco et al., 2007; Schaeffer et al., 2001a) (Funa et al., 2006; Ghosh et al., 2008; Saxena et al., 2003) (Arora et al., 2005; Ferreras et al., 2008; Trivedi et al., 2004)

(Kumar et al., 2007; Onwueme et al., 2004; Trivedi et al., 2005)

263

(continued)

Table 12.2 (continued) Enzymatic assays

Comments

References

4. Sulfotransferase (Stf ) assays

1. Stfs are responsible for transfer of a sulfuryl group from PAPS to a variety of substrates 2. Product characterization using radio-TLC, radio-HPLC and mass spectrometric approaches

(Mougous et al., 2004; Rivera-Marrero et al., 2002)

5. Acyl CoA carboxylase (ACC) assays

1. ACCs convert acetate and propionate units to malonyl CoA and methylmalonyl CoA which is utilized by the FAS and PKS enzymes 2. Product characterization using liquid scintillation counting and HPLC analysis

(Daniel et al., 2007; Gande et al., 2004; Oh et al., 2006; Rainwater and Kolattukudy, 1982)

6. Glycosyl-transferase (Gtf ) assays

1. Gtfs catalyze transfer of sugars onto various lipids 2. Product characterization using liquid scintillation counting, radio-TLC and mass spectrometric approaches

(Birch et al., 2008; Cooper et al., 2002; Pathak et al., 2002; Seidel et al., 2007)

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2. Acetate and Propionate Feeding Studies Initial analysis of wax components from Mtb revealed a mixture of fatty acids ranging from saturated palmitic acid to a wide variety of methylbranched fatty acids. These included tuberculostearic acid (10-Methyl C18 fatty acid), dextrorotatory fatty acids analogous to phthienoic acid (trimethyl unsaturated C27 acid) and levorotatory fatty acids called mycocerosic acids (tetramethyl branched C28 to C32 acids) (Anderson, 1941; Asselineau, 1966; Asselineau and Laneelle, 1998; Brennan, 1988; Goren, 1972). While the structure and chemical nature of these compounds was elucidated using conventional chemical characterization techniques, the enzymology and biochemistry of their in vivo biosynthesis was not well understood. Key insights into the biosynthesis of branched-chain fatty acids were provided by precursor feeding experiments in Mtb (Fig. 12.2) (Gastambide Odier et al., 1963; Narumi et al., 1973; Yano and Kusunose, 1966). Radioactive 14C-propionate/14C-acetate units are converted to methylmalonyl CoA/ malonyl CoA, respectively, by biotin-dependent propionyl/acetyl CoA carboxylases (Rainwater and Kolattukudy, 1982; Rawlings, 1997; Savvi et al., 2008). These methylmalonate and malonate units are used by the fatty acid synthase (FAS) and PKS enzymatic machineries for biosynthesis of various O

R′

FATTY ACID (R′ = H, CH3)

∗ SCoA

HO R′

R′

Extender unit

AT

Exterior

O

O

R

R

SCoA

KS

∗ S

HO KS

CO 2

O

O FAS

KR DH

O ∗ S

KR DH

KR DH ER

ACP

O

ts

∗ S

R

OH O

KR

ACP

R′

duc

ro dp rate

u

Sat

∗ S

R

ER

R′

ACP

R′

Interior

R

O

O S

Starter unit

R′ = H

ACC/ PCC

∗ SCoA

∗ OH

R′

O

O

O

O

∗ OH

S PK

∗ S

R′

ACP

d

ate

r satu /un ted s ura oduct t a S pr

R′

ACP

R′

O R ACP

∗ S

R

R′ = H, CH3

Figure 12.2 Overview of the acetate/propionate feeding experiments with their channeling to FASs and PKSs. The radioactive carbon is marked with an asterisk (*). ACC/ PCC, acyl/propionyl CoA carboxylase; KS, ketosynthase; AT, acyl transferase; ACP, acyl carrier protein.

266


metabolites. Lipid analysis of Mtb grown in the presence of 14C-propionate as a tracer revealed incorporation of radioactive label into the branched end of mycocerosic acids (Gastambide Odier et al., 1963). However, the fatty acid synthase responsible for the biosynthesis of these methylated acids could not be identified and it took another 20 years before Kolattukudy and coworkers demonstrated the existence of another elongation system. This elongation system, referred to as mycocerosic acid synthase (MAS), resembled the FASs in all respects, except for exhibiting specificity for methylmalonyl CoA (Gastambide Odier et al., 1963; Rainwater and Kolattukudy, 1983). Cloning and expression of the mas gene and construction of a genetic knockout strain of Mycobacterium bovis BCG facilitated investigation of the in vivo role of MAS in mycobacterial biology (Azad et al., 1996; Mathur and Kolattukudy, 1992). 14Cpropionate feeding experiments with this knockout strain established a role of MAS in biosynthesis of mycosides (phenolic glycolipids), and this methodology provided a basis for analyzing the in vivo role of mycobacterial genes in the biosynthesis of various cell wall lipids (Azad et al., 1996). Moreover, the presence of other methylated fatty acids in mycobacteria suggested the existence of multiple mas-like genes (Azad et al., 1996; Kolattukudy et al., 1997; Mathur and Kolattukudy, 1992).

3. Genome Sequencing and Identification of Polyketide Synthases In agreement with the presence of a number of lipid metabolites unique to Mtb, genome sequencing revealed many genes involved in lipid metabolism (Cole et al., 1998; Natarajan et al., 2008). Apart from the type I and type II FAS systems involved in fatty acid biosynthesis, several gene clusters homologous to PKSs were identified. Sequence homology studies suggested mycobacteria to contain examples of all three polyketide biosynthetic systems. (Table 12.3) (Cole et al., 1998; Natarajan et al., 2008). Since PKSs from Streptomyces are involved in the biosynthesis of polyketide natural products (Hopwood, 1997; Katz and Donadio, 1993; O’Hagan, 1992; Sanchez et al., 2008), the existence of these homologues indicated the presence of polyketide metabolites in Mtb. Prior to the genome availability, sequencing around the mas gene cluster had indicated the presence of modular PKSs (Azad et al., 1997; Kolattukudy et al., 1997). Research over the last decade has given some fascinating insights into the role of these enzymes in mycobacterial biology, where they have been implicated in the biosynthesis of various virulence lipids (Chhabra and Gokhale, 2009; Gokhale et al., 2007a,b; Jackson et al., 2007). In the following sections, we summarize the advances made toward understanding the roles of these PKSs in mycobacterial lipid biosynthesis.


267

Table 12.3 List of mycobacterial PKSs in the H37Rv strain of Mtb along with their domain organizations

Gene name

Accession number

pks15 pks1 ppsA ppsB ppsC ppsD ppsE mas pks2 pks12

Rv2947c Rv2946c Rv2931 Rv2932 Rv2933 Rv2934 Rv2935 Rv2940c Rv3825c Rv2048c

pks13 pks6 pks3 pks4 pks10 pks7 pks8 pks17 pks9 pks11 pks18 pks5 MbtC MbtD pks14 pks16

Rv3800c Rv0405 Rv1180 Rv1181 Rv1660 Rv1661 Rv1662 Rv1663 Rv1664 Rv1665 Rv1372 Rv1527c Rv2382c Rv2381c Rv1342c Rv1013

Domain organization

KS AT-DH-ER-KR-ACP ACP-KS-AT-KR-ACP KS-AT-KR-ACP KS-AT-DH-ER-KR-ACP KS-AT-DH-KR-ACP KS-AT-ACP-C KS-AT-DH-ER-KR-ACP KS-AT-DH-ER-KR-ACP KS-AT-DH-ER-KR-ACP-KS-AT-DHER-KR-ACP ACP-KS-AT-ACP-TE ACP-KS-AT-ACP-TE KS AT-DH-ER-KR-ACP KS (Type III) KS-AT-DH-ER-KR-ACP KS-AT-DH-ER KR-ACP KSQ-AT-ACP KS (Type III) KS (Type III) KS-AT-DH-ER-KR-ACP KS AT-KR-ACP ErroneouslyWrongly annotated ErroneouslyWrongly annotated

4. Mycobacterial Polyketide Synthases 4.1. Biosynthesis of dimycocerosate esters (DIMs) by PKS15/1, PpsABCDE, and MAS Analysis of lipid extracts suggested a number of hydroxy compounds in mycobacteria (Asselineau, 1966). Among these, the methoxyglycols were termed as phthiocerols (3-methoxy, 4-methyl, 9,11-dihydroxy glycols) and were found to be esterified with mycocerosic acids. Interestingly, bovine strains of mycobacteria were found to contain a glycolipid called mycoside B,

268


which on hydrolysis yielded mycocerosic acids and a variant of phthiocerol called phenolthiocerol (p-glycosylated phenylglycol). Phenolthiocerol possesses a phthiocerol-like chain but ends with a phenolic moiety glycosylated with various sugars (Asselineau, 1966; Onwueme et al., 2005a). Together, the phthiocerol esters and the phenolthiocerol esters are referred to as DIMs. These esters are present on the cell surface of Mtb and have been implicated in its virulence (Camacho et al., 1999; Cox et al., 1999; Reed et al., 2004). Knockout studies provided early insights into the biosynthesis of these compounds. While the mas-disrupted mutant of M. bovis BCG was incapable of synthesizing mycocerosic acids (Azad et al., 1996), the ppsdisrupted mutant lacked PDIMs and phenolic glycolipids (PGLs) (Azad et al., 1997). Genome sequencing revealed that the genes involved in biosynthesis of these metabolites are clustered in a large 73-kb operon (Cole et al., 1998; Onwueme et al., 2005a). A combination of genetic and biochemical studies has now provided a comprehensive picture for biosynthesis of these compounds (Fig. 12.3). The biosynthesis of PGLs and PDIMs can be dissected into four steps: (1) priming of PpsA with appropriate PGLor PDIM-specific starter unit; (2) extension of the primer unit by PpsABCDE, leading to the generation of the diol; (3) biosynthesis of mycocerosic acids by MAS; and (4) esterification and final assembly. The biosynthesis of PGLs is initiated by utilizing a common metabolic intermediate, chorismate, which is converted to p-hydroxybenzoic acid (pHB) by a chorismate pyruvate-lyase, Rv2949c (Stadthagen et al., 2005). pHB is then activated and transferred to a type I iterative PKS15/1enzyme by FAAL22 (Ferreras et al., 2008). FAAL22 is a member of a newly discovered family of fatty acyl AMP ligases (FAALs) that activate fatty acids as acyl adenylates instead of acyl coenzyme A (Ferreras et al., 2008; Trivedi et al., 2004). PKS15/1 exhibits extender unit specificity for malonyl CoA and is believed to extend pHB to p-hydroxyphenylalkanoate (Fig. 12.3A). The H37Rv strain of Mtb is devoid of PGLs due to a frameshift mutation in this gene (Constant et al., 2002; Reed et al., 2004). The Beijing family of Mtb strains, as well as M. bovis BCG and Mycobacterium leprae, possess a functional copy of this gene and make PGLs (Daffe and Laneelle, 1988; Reed et al., 2004; Tsenova et al., 2005). The p-hydroxyphenylalkanoate chain is then transferred to the PpsA starter for PGL biosynthesis. The starter n-fatty acyl units for PDIM synthesis are provided by FAAL26, which also uses the novel acyl adenylate activation mechanism (Trivedi et al., 2004). From this stage onward, the biosynthesis of PGLs and PDIMs follows the same biosynthetic route (Fig. 12.3B). PpsA catalyzes extension of the starter units with malonyl CoA, which results in the formation of a monohydroxy fatty acid. This is due to the presence of a single ketoreductase (KR) auxillary domain in PpsA. The acyl chain thus generated is then transferred to PpsB, which catalyzes formation of a diol through another two-carbon condensation, followed by ketoreduction. PpsC adds a malonyl unit to the growing chain and also

269


PKS15/1 KS

COOH

COOH

OH Chorismate

OH pHB

ATP

S

S

O

FAAL22

CH2 Rv2949c O COOH

AT DH ER KR ACP

O

n

A

PPi AMP OH OH

B

AMP PPi

PpsA

PpsB

PpsC

PpsD

PpsE

PCP KS AT KR ACP

KS AT KR ACP

KS AT DH ER KR ACP

KS AT DH ER KR ACP

KS AT ACP C

SH

S

FAAL26 O

S

S

O

ATP

S

O

O

O

HO

HO

HO

HO

HO

S

S

S

O HO

O

S

O

O

S

S

O

O

OH

HO

HO

HO

HO

R2= –H, –CH3 HO

HO

HO

HO

HO HO

R1

–

R1=–(CH2)3–7 CH3

R1

R1

or

R1

OH

R1

R2

O

R1 R1

R1 R1

R1

C

R1

MAS

MAS KS AT DH ER KR ACP O

S

S O

PpsE

KS AT DH ER KR ACP SH

O

KS AT ACP C

S

S

S O R

2

O

PapA5 R2 R1 R3

O O O O

O

R5

R4

HO HO

R4 = –CH2CH3or –CH3 R5 = –OCH3or =O

R3

R3 = –(CH2)3-7 –CH 3

Mycocerosic acids

R3 R1

Figure 12.3 (A) Biosynthesis of pHB and loading onto PKS15/1. (B) Biosynthesis of diol component of PGLs and PDIMs by modular PpsABCDE machinery. (C) Biosynthesis of mycocerosic acids by iterative MAS and PapA5 mediated condensation with diol for the formation of PGLs and PDIMs. The ER domain in PpsD is a trans ER, which is not a part of the type I architecture of PpsD. For simplicity, it has been shown within the PpsD domain organization.

catalyzes complete reduction to a methylene group. PpsD, in conjunction with a trans-acting enoyl-reductase (ER), Rv2953, extends this chain further with a methylmalonyl moiety (Simeone et al., 2007; Trivedi et al., 2005). The final extension to a phenolphthiocerol or phthiocerol chain is performed by PpsE, which can utilize either malonyl CoA or methylmalonyl CoA extender units (Simeone et al., 2007; Trivedi et al., 2005). MAS possesses all three auxiliary domains (KR, dehydratase [DH] and ER) necessary for complete reduction of newly-generated b-carbonyl acyl chain. MAS carries out iterative condensation of multibranched fatty acids

270


by using medium- to long-chain fatty acyl CoA starters with methylmalonyl CoA extender units (Mathur and Kolattukudy, 1992; Onwueme et al., 2004; Trivedi et al., 2005). Polyketide-associated protein A5 (PapA5) interacts with MAS and brings about trans-esterification of mycocerosic acids onto the diol component of phthiocerol/phenolpthtiocerol (Fig. 12.3C) (Mathur and Kolattukudy, 1992; Onwueme et al., 2004; Trivedi et al., 2005). Final processing and transport of DIMs requires other proteins like Rv2951c and Mtf2, which bring about reduction of the 3-keto group and subsequent Omethylation of this hydroxyl group. The glycosylation of the phenyl ring in PGLs is carried out by glycosyl transferases, Rv2962c, Rv2958c, and Rv2957c, which modify the phenyl ring with addition of two rhamnose and one fucose sugars. The fucose ring is further modified by action of methyl transferases, which complete the assembly of tri-O-methyl-dirhamnosyl-phenolphthiocerol dimycocerosates (Onwueme et al., 2005b; Perez et al., 2004a,b). The transport of the fully assembled DIMs to the cell wall is proposed to be mediated by a transmembrane protein, MmpL7, which is thought to couple synthesis with transport by specifically interacting with PpsE ( Jain and Cox, 2005). DrrC and LppX are other accessory proteins that mediate the transport of DIMs to the periphery of the cell wall ( Jain and Cox, 2005; Onwueme et al., 2005a; Sulzenbacher et al., 2006).

4.2. PKS2 is involved in biosynthesis of sulfolipids Sulfolipids (SLs) were identified in the late 1950s from Mtb, while studying a sulfur-containing material capable of fixing the cationic dye neutral red (Dubos and Middlebrook, 1948; Middlebrook et al., 1959). Subsequent analysis of this material by Goren and coworkers revealed a mixture of highly related compounds, with the most abundant being sulfolipid-I (SL-I). Chemical analysis revealed a trehalose-2-sulfate (T2S) core, tetra-acylated with fatty acids. While one of the fatty acid substituents is a straight-chain fatty acid (primarily palmitate or stearate), the other three are long-chain methylated fatty acids called phthioceronic acid (PA) or hydroxy phthioceronic acids (HPA) (Goren, 1970a,b, 1971, 1976). Phthioceronic acids differ from mycocerosic acids in having an absolute configuration of S- (also referred to as L- based in older nomenclature) for the methyl-branched carbon, as compared to R- (or D-) in the case of mycocerosic acids (Asselineau, 1966). Gene inactivation and complementation studies clearly indicated an essential role for PKS2 in the biosynthesis of these unusual acids (Sirakova et al., 2001). PKS2 possesses a complete set of active sites to add a completely reduced ketide unit to the starter chain. By a mechanism analogous to MAS, PKS2 catalyzes the biosynthesis of long-chain branched fatty acids by iterative utilization of methylmalonyl CoA (Chopra and Gokhale, unpublished results). The hydroxyl-modification of PA to produce HPA could in principle involve a hydroxylase, which remains to be identified.

271


Other biosynthetic steps have been deciphered by generating mutants of Mtb lacking various enzymes involved in SL biosynthesis (Fig. 12.4). The biosynthesis is initiated by a sulfotransferase, Stf0, which transfers a sulfuryl group from 30 -phosphoadenosine-50 -phosphosulfate (PAPS) onto trehalose, thereby generating trehalose-2-sulfate (T2S) (Mougous et al., 2004). T2S is acylated either by a palmitate or a stearate unit at the 20 -position by PapA2 to produce monoacylated SL. This is followed by PapA1-mediated transfer of the phthioceranoyl group from PKS2 to the palmitoyl-/stearoyl-T2S, leading to the formation of the diacylated intermediate (Bhatt et al., 2007c; Kumar et al., 2007). FAAL23 is responsible for activation and loading of starter fatty-acyl chains onto PKS2 (Chopra and Gokhale, unpublished results) (Lynett and Stokes, 2007). Since PKS2 also lacks an appended TE domain for chain release, one would expect PapA1 to interact with and sequester the chain from PKS2 for its transesterification onto monoacylated sulfolipid. A transmembrane protein called MmpL8 is also present in the SL cluster and is believed to play an important role in transport of SLs across the cell wall. Disruption of MmpL8 by two independent groups led to the accumulation of diacylated intermediates, SL1278 or SL-N (Converse et al., 2003; Domenech et al., 2004). The conversion of di-acylated sulfolipid to

OH

OH O

HO OH

stf0

AMP PPi O

S

SH

S

O

Trehalose

OSO– O

?

O

3

OH OH

O O 16–18

R1

OH

OH

O

HO OH

OH

R1

PapA2

OH

OH OH

O HO

KS AT DH ER KR ACP

S

O

FAAL23 O

OH OH

O HO

KS AT DH ER KR ACP

ATP

OH O

PAP PAPS

OSO3– O

T2S

PKS2

PKS2

O HO OH

OH

R1

Monoacylated SL

PapA1 R1= –(CH2)3–7 –CH3

R1

OH

Phthioceronic acids

O

HO OH

OSO3–

O

? OH

O

4–9

O

Diacylated SL OH

O O

14,16

O

14,16

O

OSO3–

O O OH

O O 16–18

SL1

OH OH

O

4–9

O HO OH

O

O 16–18

4–9

14,16

O 4–9

14,16

O

Figure 12.4 Assembly of SL-1. Phthioceronic acids are biosynthesized by an iterative PKS2 and are utilized for SL-1 production.

272


the final tetra-acylated form requires further investigation. The exact biological function of SL-1 in mycobacterial biology is not clear and Mtb mutants of various enzymes involved in sulfolipid biosynthesis have not provided clear correlation to its virulence (Bertozzi and Schelle, 2008). Interestingly, a sulfolipid-deficient PKS2 knockout strain of Mtb retains its ability to stain with neutral red (Andreu et al., 2004; Cardona et al., 2006).

4.3. PKS12 uses a novel ‘‘modularly iterative’’ mechanism for biosynthesis of mannosyl-b-1-phosphomycoketides Mannosyl-b-1-phosphomycoketides (MPMs) are a group of antigenic phospholipids that were recently isolated from pathogenic species of mycobacteria (Moody et al., 2000). Chemically, MPMs are similar to the mammalian mannosyl-b-1-phosphodolichols and possess an identical mannose-phosphate head group. However, the two molecules differ in their alkyl chain, which probably contributes to antigenicity of MPMs (Moody, 2001). Initial studies had suggested an isoprenoid mode of biosynthesis for this alkyl segment (now referred to as the mycoketide) (Moody et al., 2000). Careful mass spectrometric analysis of MPMs, followed by gene inactivation of pks12, revealed a polyketide origin for this chain (Matsunaga et al., 2004). Pks12 is the largest open reading frame in the Mtb genome and codes for a bimodular PKS (Cole et al., 1998; Natarajan et al., 2008). The two modules of PKS12 were predicted to exhibit methylmalonate and malonate extender unit specificity and thus mycoketide synthesis was proposed to involve five alternate condensations of methylmalonyl and malonyl units by using an iterative mechanism of biosynthesis (Matsunaga et al., 2004). Such a mechanism would require transfer of the growing chain from the ACP of the second module to the KS active site of the first module. This transfer would require covalent transfer of acyl chains over very large distances between the catalytic domains of module 2 and module 1. Based on the threedimensional organization of FASs and PKSs this seems unlikely (Chopra et al., 2008; Khosla et al., 2007; Maier et al., 2006; Sherman and Smith, 2006) and presented an interesting challenge to the intramolecular paradigm of iterative catalysis. In order to dissect the programming by which PKS12 biosynthesizes mycoketide, a systematic analysis with purified 431 kDa protein was carried out. Since PKS modules are reported to ‘‘stutter’’ and ‘‘skip’’ during biosynthesis (Thomas et al., 2002; Wenzel et al., 2005; Wilkinson et al., 2000), it was important to ascertain the specificity of the two AT domains. The AT domain specificity was investigated by engineering a single module PKS12D1 protein. These studies demonstrated stringent methylmalonate and malonate specificity for module 1 and module 2, respectively, making it mandatory to use these two modules alternately. Biochemical analysis and


273

ultracentrifugation studies interestingly suggested formation of a large supramolecular assembly for these proteins. Since PKS12 was found to be active under these conditions, it was speculated that this could be the functionally competent form of the protein. Modular PKSs are known to form specific interactions through the linker proteins and investigation of these linker sequences by swapping or by mutagenesis emphasized their role in formation of this assembly. Further proof for this organization was provided by the demonstration of specific transfer of the acyl chain from (n–1) protein to the next protein (n) through interacting C- and N-terminus linkers. Thus, catalysis proceeded by formation of a specific self-organizing assembly involving multiple molecules of PKS12 and the mechanism was referred to as ‘‘modularly iterative’’. PKS12 provides a novel example of intermolecular iterative catalysis for PKSs (Chopra et al., 2008). A complete picture for biosynthesis of MPM could be drawn from analysis of the pks12 cluster in the mycobacterial genome. The release of the mycoketide chain from PKS12 could occur via two alternative mechanisms. LipT possesses an esterase motif and could carry out hydrolytic release of the chain, which undergoes reduction and processing to form mycoketide. The other possibility is a direct reductive release of mycoketide alcohol by Rv2047c, which resembles reductive domains from multifunctional non–ribosomal peptide synthetases. A swivel domain commonly found in phosphoenolpyruvate (PEP)—utilizing enzymes is also present in Rv2047c and could play a role in phosphorylation of the reduced mycoketide chain. Finally, the mannosylation could be carried out by a polyprenol monophosphomannose synthase, Ppm1, encoded in the cluster (Fig. 12.5). Although the exact role of MPM in the biology of mycobacteria is not known, being an activated form of mannose sugar, MPMs could play an important role in transfer of sugars across the cell membrane.

4.4. PKS13 catalyzes condensation of fatty-acyl chains during biosynthesis of mycolic acids Mycolic acids are the most abundant lipids found in the mycobacterial cell wall and are responsible for the ‘‘acid-fast’’ nature of mycobacteria. Initial characterization by Anderson in the late 1930s suggested a general formula of C88H176O4, and a characteristic property to yield normal hexacosanoic acid on pyrolytic distillation under vacuum (Asselineau, 1966; Asselineau and Laneelle, 1998). Detailed chemical characterization over the years has revealed them to be a-alkyl-b-hydroxy fatty acids, which are present either as mycolyl-esters or as free fatty acids in the cell wall (Asselineau and Lederer, 1950; Brennan and Nikaido, 1995; Goren, 1972). The mycolate structure can be broken down to a saturated alkyl chain condensed to a longer meromycolate chain, which may carry various modifications. These modifications on the meromycolate chain classify mycolic acids into alpha-,

274


PKS12

PKS12 KS AT DH ER KR ACP KS AT DH ER KR ACP

KS AT DH ER KR ACP KS AT DH ER KR ACP S O

R

O

S

O

R

S

S O

PKS12

S

S

O

O

S

O

KS AT DH ER KR ACP KS AT DH ER KR ACP

S

S

O

O

R R

R = H or C –C 1 4

R R

R

?

R

O R

HO

Mycoketide acid

R

? HO

R

Mycoketide alcohol O –O P O O–

HO HO HO

HO O

? R

Phospho-mycoketide

? O O P O O–

R

Mannosyl-b-1-Phospho mycoketide (MPM)

Figure 12.5 PKS12 follows a ‘‘modularly-iterative’’ mechanism for the biosynthesis of mycoketide, which is further phosphorylated and glycosylated for the formation of MPMs.

keto-, and methoxy-subgroups. a-mycolic acids contain two cyclo-propane rings on the meromycolate chain and are the major type of mycolic acids in most mycobacterial species. Keto- and methoxy-mycolic acids carry additional oxygen functionalities in the meromycolate chain and are often termed oxygenated mycolates (Barry et al., 1998; Minnikin and Polgar, 1967; Takayama et al., 2005; Toubiana et al., 1979). Mycolic acids have also been found to contain unsaturation in the meromycolate chain. TLC and mass spectrometric approaches for mycolic acid characterization now provide a means to discriminate between many closely related mycobacterial species (Asselineau and Laneelle, 1998; Marrakchi et al., 2008; Minnikin et al., 1984; Takayama et al., 2005). The biosynthetic pathway for mycolic acids uses both type I FAS (all enzymatic domains on a single polypeptide) and type II FAS (enzymatic domains on separate polypeptides) machinery for the synthesis of the chains, and a PKS called PKS13 for their condensation (Fig. 12.6). The biosynthesis can be described in three steps: (1) type I FAS-mediated biosynthesis of both the alpha-alkyl chain and the meromycolate precursor, (2) extension and modification of the meromycolate chain by type II FAS assembly, and (3) condensation of the two chains and export to the cell wall. Rv2524c is the type I FAS that utilizes acetyl CoA as the starter and carries out repetitive decarboxylative condensations with malonyl CoA to biosynthesize the hexacosanoyl CoA (alpha chain) and the meromycolate

275


KS AT DH ER KR ACP TE

KS AT DH ER KR ACP TE S

SH

S

O

O

S

SH

S

O

O

n

O

n

O

KS AT DH ER KR ACP TE

Type I FAS Rv2524c

KS AT DH ER KR ACP TE S

HO

n

O

n

SH

O– KS AT DH ER KR ACP TE

O

AcpM

n+2

S

SH

O O –O

O

SCoA

SCoA

α-chain AccD4, AccD5

O SCoA COO–

FabD

O –O

2-Carboxy hexaxosanoyl-CoA

SCoA

O O AcpM

S

x X = 11

FabH

PKS13 AcpM

O O

O S

x

AcpM

MabA

Type II FAS S

AMP PPi

ATP

SH S O

PKS13

S –OOC

ACP KS AT KR ACP TE

O

SH SH

FAAL32

O

S

O

KasA/ KasB

OH O x

S

ACP KS AT KR ACP TE

AcpM

AcpM dehydratase

O S X+2

AcpM

InhA O x

S

AcpM

C

ar

bo Rv ny 25 l r 09 ed uc tio

n

Meromycolate chain

Mature mycolic acids

Figure 12.6 Biosynthesis of mycolic acids requires formation of meromycolate-chain and a-chain through the type I FAS/type II FAS systems.This is followed by condensation by PKS13.

precursor (Bloch, 1977; Smith et al., 2003). The ketosynthase, FabH (Rv0533c), links the type I and II FAS pathways and catalyzes condensation of type I FAS-derived meromycolate precursor with malonyl units present on AcpM (Choi et al., 2000; Schaeffer et al., 2001a). The generation of malonyl-AcpM is catalyzed by the enzyme FabD (Kremer et al., 2001). Condensation of meromycolate precursor with malonyl CoA leads to chain extension by two units and is subjected to a cycle of keto-reduction, dehydration and enoyl-reduction, catalyzed by MabA (FabG1, Rv1483)

276


(Marrakchi et al., 2002), AcpM dehydratase (Cronan et al., 1988; Gurvitz et al., 2008b; Sacco et al., 2007), and InhA (Rv1484) (Gurvitz et al., 2008a), respectively (Fig. 12.6). InhA is inhibited by the antituberculosis drug isoniazid via formation of a covalent adduct with NAD+ (Cronan et al., 1988; Dessen et al., 1995; Gurvitz et al., 2008a,b; Marrakchi et al., 2000, 2002; Quemard et al., 1995). The extended chain is transferred to KasA (Rv2245) and KasB (Rv2246), which catalyze further extension using the same sets of enzymes (Bhatt et al., 2005, 2007a; Kremer et al., 2002; Schaeffer et al., 2001b; Slayden and Barry, 2002). It is proposed that while FabH catalyzes the initial condensation, KasA carries out extension to an intermediate stage, followed by extension to full-length meromycolate by KasB. The modifications in the meromycolate chain are brought about by various cyclopropane synthases and methyl transferases during the type II FAS-mediated chain extension cycles (Bhatt et al., 2007b; Marrakchi et al., 2008; Takayama et al., 2005). The condensation of the meromycolate and the alpha chain is brought about by PKS13, which has domain architecture of ACP-KS-AT-ACP-TE (Gokhale et al., 2007b; Portevin et al., 2004). The meromycolate chain from meromycolyl-AcpM is activated by FAAL32 and transferred to the KS domain of PKS13 through the N-terminal ACP domain (Trivedi et al., 2004). It is not clear whether FAAL32 directly interacts with AcpM and sequesters the chain or the chain is released prior to activation by FAAL32. Hexacosanoyl CoA, derived from the type I FAS pathway, is acted upon by two acyl CoA carboxylases, AccD4 and AccD5, leading to formation of 2-carboxy-hexacosanoyl CoA (Gande et al., 2004, 2007). Through the AT domain of PKS13, the 2-carboxy-hexacosanoyl CoA is transferred to the ACP domain where it undergoes a decarboxylative Claisen condensation with the meromycolate chain. Rv2509 is believed to carry out the final reduction of the b-keto group to a secondary alcohol for the formation of mature mycolates (Fig. 12.6) (Bhatt et al., 2008; Lea-Smith et al., 2007). It is proposed that the mature mycolic acids are transferred from the ACP domain of PKS13 to mannopyranosyl-1-phosphoheptaprenol (PL), which transfers the mycolyl-group further to trehalose-6-phosphate to yield trehalose-monomycolyl (TMM) phosphate. Dephosphorylation of TMMphosphate leads to formation of TMM, which is exported out to the cell wall. Enzymes responsible for transfer of mycolates from PKS13 to the cell wall have not been very well characterized. The extracellular mycolyl transferases called the Ag85 complex are proposed to catalyze the formation of TDM and arabinogalactan-mycolate from TMM (Bhatt et al., 2007b; Marrakchi et al., 2008; Takayama et al., 2005). Recent studies suggest new possible modes of lipid biosynthesis involving formation of long-chain fatty acids, including mycobacteric acids, by degradation of mycolic acids (Rafidinarivo et al., 2008).

277


4.5. PKS3/4 is involved in the biosynthesis of phthenoic acids One of the components of the Mtb wax fraction analyzed by Anderson and coworkers was dextrorotatory fatty acids called ‘‘phthioic acids’’ esterified to a sugar. These acids were initially thought to be completely saturated trimethylated fatty acids. Detailed chemical characterization by two independent research groups revealed their chemical nature as trimethylated a,b-unsaturated acids or 2,4,6-trimethyl tetracos-2-enoic acids. These acids were independently referred to as phthienoic acids or mycolipenic acids by the two groups (Fig. 12.7) (Asselineau, 1966; Asselineau et al., 1972). During the late 1980s, analysis of mycobacterial glycolipids revealed a penta-acylated compound where four phthienoyl-groups and one palmitoyl- or stearoyl-group were found to occupy the 2, 20 , 30 , 4, and 60 positions of trehalose sugars (Daffe et al., 1988). These compounds were termed polyacyltrehaloses (PATs) and were found in virulent human and bovine strains of mycobacteria (Fig. 12.7). Similarly, a number of triacylated trehalose (TATs) and diacylated trehalose (DATs) compounds containing various different acylations were identified (Besra et al., 1992; Cruaud et al., 1990; Lemassu et al., 1991; Munoz et al., 1997a,b). Biochemical analysis of a PKS3/4 mutant strain of Mtb suggested involvement of PKS3/4 in the biosynthesis of phthienoic acids. Interestingly, H37Rv genome sequencing had suggested pks3 and pks4 to be independent open reading frames. Subsequent analysis identified an error in sequencing and showed PKS3/4 to be a single protein with KS-AT-DH-ER-KR-ACP domain organization. The absence of PATs from the Mtb strain caused cells to stick to each other as a clump without affecting the overall growth rate (Dubey et al., 2002). This suggested PATs to be localized on the outer surface of the cell wall. In another study, a pks3/4 mutant Mtb strain showed improved efficiency of binding to the host cells (Rousseau et al., 2003a). However, this property did not affect the overall replication and persistence of the bacillus in the host cells.

O

O HO

O

OH O O

O

C17H35

O O

O O

C17H35

OH O O

C17H35 O

C17H35 C17H35

Figure 12.7 Structure of polyacyltrehaloses. One of the phthienoic acids is highlighted in the box.

278


The biochemical pathway involved in assembly of PATs has not been investigated. Analysis of the pks3 and pks4 genetic locus reveals genes which could participate in the complete assembly and transport of these lipids. FAAL21 could activate and load fatty acyl chains onto PKS3/4 for the biosynthesis of phthienoic acids. Also encoded in the cluster is a PKSassociated protein, PapA3, which could mediate trans-esterification of phthienoic acids onto trehalose sugars and MmpL10 for export of PATs to the outer cell wall.

4.6. PKS10, PKS7, PKS8, PKS17, PKS9, and PKS11 constitute an unusual PKS cluster The H37Rv genome encodes three genes homologous to type III PKSs. Interestingly, two of these genes, pks10 and pks11, are present on either side of four type I PKSs, constituting a pks cluster (pks10-pks7-pks8-pks17-pks9pks11) (Cole et al., 1998). PKS7 is 31% identical to MAS and contains all the three auxiliary domains that could completely reduce a ketide unit. PKS8 contains KS, AT, DH and ER domains and PKS17 contains KR and ACP domains. Together, PKS8 and PKS17 would form one complete module. PKS9 resembles loading modules of modular PKSs and comprises KS, AT and ACP domains. In PKS9, the active site cysteine of KS is mutated to glutamine and is referred to as a KSQ domain. This type of KSQ domain has been previously reported for certain modular PKSs, where they function as chain initiation factors similar to the chain length factor (CLF) of aromatic PKSs (Bisang et al., 1999). The genome of M. bovis has also revealed an identical genomic organization of the pks10-pks11 cluster, with the putative proteins sharing 98 to 100% sequence identity with the M. tuberculosis homologues (Garnier et al., 2003). However, M. avium subsp paratuberculosis shows slight variations in the genomic organization and the broken module is missing from the cluster (Fig. 12.8) (Li et al., 2005). The functional importance of the pks10-pks11 genomic cluster is yet to be established in Mtb and M. avium subsp paratuberculosis. Gene inactivation studies suggest a possible role of pks7 and pks11 in the biosynthesis of PDIMs (Rousseau et al., 2003b; Waddell et al., 2005). This could also be due to spontaneous loss of PDIMs from the mutant strains (Domenech et al., 2004). Another report suggested a role for pks8 and pks17 in the biosynthesis of methyl-branched unsaturated fatty acids that are esterified to acyltrehaloses and sulfated acyltrehaloses as minor constituents (Dubey et al., 2003). Biochemical characterization of PKS11 suggests that it may be able to produce resorcinolic metabolites, which are known for their involvement in cellular physiology and membrane chemistry in other organisms (Saxena and Gokhale, unpublished results) (Kozubek and Tyman, 1999). These amphiphilic molecules possess diverse biological functions and are active antimicrobial and antiparasitic compounds. They are also known to

279


PKS10

M. tuberculosis

KS (type III)

PKS10

M. bovis

KS (type III)

PKS10

M. avium ssp. paratuberculosis

Figure 12.8 species.

KS (type III)

PKS7 KS AT DH ER KR ACP



PKS8 KS AT DH ER

PKS8 KS AT DH ER

PKS9 KSQ AT ACP

PKS17

PKS9

PKS11

KR ACP

KSQ AT ACP

KS (type III)

PKS17

PKS9

PKS11

KR ACP

KSQ AT ACP

KS (type III)

PKS11 KS (type III)

Organization of the PKS10-PKS11 cluster in various mycobacterial

modulate oxidation of liposomal membranes and fatty acids (Gubernator et al., 1999). Interestingly, early analysis of unsaponifiable fraction of fats from M. leprae has demonstrated the presence of methoxylated resorcinolic metabolites called a- and b-leprosols (Asselineau, 1966; Bu’Lock and Hudson, 1969). Similar molecules have been recently shown to be essential for formation of metabolically dormant cysts in Azotobacter vinelandii (Funa et al., 2006). It is tempting to speculate that these metabolites may be produced under specific conditions in mycobacteria and may have a role to play in the onset of dormancy.

4.7. PKS18 is involved in biosynthesis of long-chain pyrones The third type III pks gene, pks18, is not flanked by PKS-related genes and shows 40 to 45% sequence homology with bacterial and plant type III PKSs. Sequence analysis of PKS18 shows conservation of the catalytic and key active site residues of this class of proteins (Cole et al., 1998). While no physiological role has been assigned to pks18, biochemical investigation of its product revealed remarkable specificity for long-chain aliphatic CoA analogues. This unusual substrate specificity is unprecedented in the chalcone synthase super-family of type III PKSs and has added a new functional relevance to these proteins. PKS18 efficiently produces long-chain a-pyrones when primed with the long fatty acyl CoAs (Rukmini et al., 2004; Sankaranarayanan et al., 2004; Saxena et al., 2003). Since the biochemical characterization of PKS18, a number of plant and bacterial homologues have been shown to utilize long-chain precursor molecules for the synthesis of acyl pyrones. Such acyl pyrones have been recently identified in the cell envelope of Azotobacter sp. (Abe et al., 2004, 2005; Austin et al., 2004; Funa et al., 2006; Zha et al., 2006).

280


4.8. MbtC and MbtD are involved in biosynthesis of iron-chelating siderophores from Mtb Siderophores are iron-chelating compounds (see Chapter 17 of volume 458) that were discovered in mycobacteria in the late 1950s while searching for factors important for the growth of M. paratuberculosis (Snow, 1970). Subsequent research has revealed two types of siderophores which collectively scavenge iron in the Fe (III) form from the host organism. Mycobactins or intracellular siderophores are found within the cell envelope of the mycobacteria and are believed to play a role in controlled release of iron inside the cell. Extracellular siderophores vary in composition and are called carboxymycobactin or exochelins depending on whether the organism is pathogenic or saprophytic (Ratledge, 2004; Ratledge and Marshall, 1972; Rodriguez, 2006). Chemically, mycobactin and carboxymycobactin have a central lysine core which is modified at both the a- and e-amino termini with a hydroxyaryloxazoline group and an alkyl group, respectively (Fig. 12.9). The alkyl group varies from C10 to C21 in the case of mycobactin and sometimes contains a cis-double bond. However, it is shorter in the case of carboxymycobactin and carries a free carboxy group at the end. It is this alkyl group that differentiates carboxymycobactin from mycobactin. On the carboxylend, lysine is modified with a polyketide-derived b-hydroxy butyrate group, which is further linked to another N-hydroxylated and cyclized lysine. All three modifications on lysine together constitute the iron-coordinating framework of mycobactins (Ratledge, 2004). Although the biochemical reconstitution of mycobactin assembly has not been carried out, two gene clusters—mbt1 and mbt2—are proposed to be involved in its biosynthesis (Quadri et al., 1998). Expression of these two clusters is believed to be mbt-2 L

M

N K

mbt-1 I

J

A

B

KS

AT KR ACP

C

D

MbtK MbtL MbtM

O

N

E

F

G H

FadE14

R n

MbtG

OH MbtA

OH

MbtE

N MbtB

O

O

H N

O N H

O

O

MbtF

O OH N MbtJ

MbtG

MbtC/MbtD

Mycobactin: n = 17, 19; R = CH3 Carboxymycobactin: n = 2–9; R = COOH

Figure 12.9 The mbt locus involved in the biosynthesis of mycobactins.The structures of mycobactin/carboxymycobactin are shown.The core lysine is shown in bold.


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regulated by an iron-dependent IdeR repressor (Rodriguez, 2006; Rodriguez and Smith, 2003). mbt1 codes for MbtA to MbtJ, which are believed to participate in the assembly of the polyketide-peptide core (Quadri et al., 1998). mbt2 codes for enzymes responsible for modification of this core to the final metabolite (Krithika et al., 2006). The predicted pathway for assembly of mycobactin starts with the conversion of iso-chorismic acid to salicylic acid, which is activated by MbtA, a salicylate-AMP ligase, and loaded onto MbtB. MbtB is a non–ribosomal peptide synthetase and is expected to attach a Ser or Thr onto the salicylic acid core. It is this amino acid which is cyclized to an oxazoline ring, thus finishing the covalent assembly of the a-amino cap of lysine. MbtE is another NRPS and is thought to catalyzes the addition of the core lysine onto this cap (De Voss et al., 1999; Marshall and Ratledge, 1972; Snow, 1970). MbtC and MbtD are the two polyketide synthase subunits present in the cluster and code for the KS and AT-KR-ACP domains respectively. Together, they constitute a PKS enzyme believed to carry out the biosynthesis of the b-hydroxy butyrate group using acetyl and malonyl CoA units. Another NRPS called MbtF is proposed to transfer the final lysine onto the b-hydroxy butyrate group. The cyclization of this N-hydroxylated lysine group to a seven-membered lactam ring is proposed to be catalyzed by MbtJ (De Voss et al., 1999; Quadri et al., 1998). Modification of the e-amino termini of the core lysine is brought about by the mbt2 cluster, which codes for an N-acyl transferase (MbtK), an acyl carrier protein (MbtL), a fatty acyl AMP ligase (FAAL33 or MbtM), and an acyl CoA dehydrogenase (FadE14 or MbtN). FAAL33 activates and loads long-chain fatty acids onto MbtL, which are then transferred by MbtK onto the e-amino group of the core lysine. FadE14 is the enzyme responsible for the a,b-unsaturation present on the acyl chain. The N-hydroxylation of the lysine e-amino group has been shown to be catalyzed by the N6-hydroxylase, MbtG, from the mbt1 locus by using substrate mimics (Krithika et al., 2006). This enzyme belongs to a class of flavoprotein monooxygenase and uses molecular oxygen for hydroxylation. Although the biosynthesis of mycobactin has been dissected out in detail, the complete sequence of events that leads to its assembly has not been elucidated.

4.9. PKS5 and PKS6 While PKS5 is a type I PKS with a domain organization of KS-AT-DHER-KR-ACP, PKS6 is a type I PKS with domain architecture ACP-KSAT-ACP-TE, similar to PKS13 (Yadav et al., 2003). PKS5 is 66% identical to MAS, and biochemical analysis of a PKS5 mutant of Mtb reveals that its cell envelope composition is identical to that of the wildtype strain (Rousseau et al., 2003b). PKS5 could be involved in the biosynthesis of an unknown lipid which is a minor constituent of the cell wall. PKS6, on the

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other hand, is implicated in the biosynthesis of an unknown polar metabolite (Waddell et al., 2005), and FAAL30 has been shown to be involved in the activation and transfer of starter fatty-acyl chains onto PKS6 (Trivedi et al., 2004; Waddell et al., 2005). Interestingly, an Mtb mutant of pks6 was impaired for growth in the lungs of Balb/c mice, suggesting that the PKS6derived metabolite may play an important role in mycobacterial survival and virulence (Camacho et al., 1999). PKS14 and PKS16 are wrongly annotated as PKSs in the H37Rv genome. PKS14 is a 120–amino acid protein with no conserved domains and PKS16 is a 544–amino acid protein belonging to the acyl activating super family of enzymes (Cole et al., 1998).

5. Techniques to Characterize PKS Enzymes 5.1. Product formation assay to study activity of PKS enzymes A protocol to assay mycobacterial PKSs involved in biosynthesis of acyl chains of virulent lipids is described below. 1. Purify the PKS in 100 mM phosphate buffer containing 10% glycerol and 1mM Tris (2-carboxy ethyl) phosphine hydrochloride (TCEP). The protein can be stored at –80 after snap-freezing in liquid nitrogen. 2. Set up an assay with 100 mM phosphate buffer (pH 7.2), 150 mM starter acyl NACs, 75 mM extender acyl CoA, 7.2 mM 14C- extender acyl CoA, 4 mM NADPH, 10% glycerol, 2 mM TCEP, and 2.5–4.5 mM protein in a 100-ml reaction volume. 3. Incubate the reactions at 30 C for 6 to 12 h. If the PKS has an appended TE domain or follows any other chain release mechanism, proceed to Step 5. 4. Carry out alkaline hydrolysis of the products by addition of a half-volume 45% KOH and heat at 70 for 10 min. 5. Acidify the reactions with 50% HCl and extract with freshly distilled ethyl acetate (300 ml 2). Pool the two extractions. 6. Vacuum dry the extractions and resuspend in a small volume of ethyl acetate (20 ml). This contains the reaction products and can be directly analyzed by various techniques like TLC, HPLC or mass spectrometry.

5.2. Characterization of PKS derived saturated fatty acids 1. Spot the extracted products after alkali hydrolysis on a silica gel TLC plate and run in 40:60:1 hexanes: ether: acetic acid. Expose the TLC to an imaging plate for 6 to 12 h and analyze radiolabeled products using a PhosphorImager. The fatty acid products run at an Rf of 0.5 (Fig. 12.10A).

A Solvent front

n C io ct N A in ea S te l R C 6ro ro h it -P ont W c

PKS12Δ1 products

Origin

1

2

Radioactive counts (volts) Radioactive counts (volts) Radioactive counts (volts)

B C12

1.0

0.5

Standard fatty acids

C6 C16

0.0

C18

C

PKS12Δ1 + C6 starter

C8

C10

C12

C14

C16

C18

29.46

32.49

36.60

42.45

0.2

0.1

0.0 1.0

MAS + C6 starter

0.5

23.29

0.0 20

30 40 Elution time in minutes

50

26.64

Elution time in minutes

Figure 12.10 (A) TLC-based characterization of PKS products. Lane1: PKS reaction primed with hexanoyl-starter. Lane 2: Control reaction in the absence of protein (Chopra et al., 2008). (B) HPLC analysis of PKS products eluted from TLC. The upper panel shows saturated fatty acids. In the middle panel are PKS12D1 products after priming with hexanoyl-starter. The lower panel shows various iterative products of MAS proteins; with increasing time, mono-, di-, tri-, and tetra-branched mycocerosic acids elute. (C) TLC analysis of HPLC-eluted PKS12D1 products (Chopra et al., 2008).

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2. To carry out a radio-HPLC analysis of the product, set up fresh product formation assays with an increased 14C extender acyl CoA concentration of 25 mM while keeping the total extender acyl CoA concentration as 100 mM. Carry out alkali hydrolysis of the products and extract in ethyl acetate as discussed above. 3. Spot the extract on a TLC-plate and run in 40:60:1 hexanes: ether: acetic acid. Scrape off the band of interest (running at an Rf of 0.5) and carry out another extraction in ethyl acetate. Vacuum concentrate the products and inject on a C18 reverse-phase column on HPLC (gradient: 100% B in 20 min, 100% B in 40 min, 20% B in 60 min). Solvent A consists of H2O, and solvent B, 5% methanol in ACN with 0.1% formic acid. Alternatively, the extracted products from the reaction mix can be directly loaded on the HPLC. However, in this case the signal-to-noise ratio is lower in comparison and peaks are not as sharp as in the case of the eluted products. Analyze the HPLC eluate using an online radioactivity detector fitted to the HPLC in series with the UV detector. 4. Various iterative products from the PKS assay run as separate peaks on the HPLC and can be confirmed by running synthetic standards (Fig. 12.10B). Splitting the flow before the radioactive detector enables collection of these peaks and further TLC or mass spectrometric analysis (Fig. 12.10C).

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C H A P T E R

T H I R T E E N

Genetic Engineering to Produce Polyketide Analogues Christopher D. Reeves* and Eduardo Rodriguez† Contents 1. Introduction 2. AT Domain Replacement to Alter a-Carbon Substitution 3. Procedure for Engineering AT Replacements in the Chromosome 4. Engineering b-Carbon Processing 5. Engineering when only a Single Crossover Event is Possible 6. Heterologous Expression of Engineered PKS Genes 7. Chemobiosynthesis 8. Mutasynthesis 9. Gene Knockouts to Obtain Analogues References

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Abstract Polyketides are pharmaceutically important and structurally diverse natural products. Creating analogues for drug development can be done with chemistry, but this is generally restricted to a few accessible functional groups. Analogues can also be made by genetic engineering, which is particularly effective for polyketides synthesized by a modular polyketide synthase (PKS). Such a PKS displays colinearity, which means that the structural features along the polyketide chain are determined by the catalytic specificities in corresponding modules along a molecular assembly line. The assembly line can be genetically engineered through addition, deletion, or mutation of catalytic domains or the reorganization of whole modules. Chemically synthesized precursors also can be fed to engineered assembly lines to further expand the repertoire of analogues. These various methods are discussed with an aim of providing a guide to the strategies most likely to succeed in a given circumstance. Recent information that could be relevant to future polyketide engineering projects is also discussed.

* {

Amyris Biotechnologies, Inc., Emeryville, California, USA Instituto de Biologıá Molecular y Celular de Rosario (CONICET), Facultad de Ciencias Bioquı´micas y Farmaceúticas, Universidad Nacional de Rosario, Rosario, Argentina


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Christopher D. Reeves and Eduardo Rodriguez

1. Introduction Many polyketides have therapeutic value and some are well-known drugs. A small sample of the polyketides produced by assembly line enzymology include macrolide antibacterials, polyene antifungals, immunosuppressants related to rapamycin and FK506, ansamycin Hsp90 inhibitors, and epothilones, microtubule stabilizing anticancer agents. The drug properties of a polyketide, are not always ideal, but the process of developing a better polyketide drug is time consuming and expensive. Analogues must be generated to obtain a structure-activity relationship profile, but the complex structures of most polyketides restrict the use of medicinal chemistry for this purpose. Genetic engineering is an alternative way to make analogues, many of which would be difficult to make using chemistry. In the early 1990s, Donadio and Katz provided the first demonstration of modular PKS engineering with the erythromycin PKS, known as 6-deoxyerythronolide B synthase (DEBS) (Donadio and Katz, 1992; Donadio et al., 1991, 1992, 1993; Katz and Donadio, 1993). Subsequent work in several laboratories showed that DEBS was very pliable to a variety of engineering approaches. When two or three of the DEBS modules were used, various triketide lactones could be produced (Bedford et al., 1996; Cortes et al., 1995; McDaniel et al., 1997; Oliynyk et al., 1996) and when the complete DEBS was engineered, many 6-dEB or erythromycin analogues could be obtained (McDaniel et al., 1999; Ruan et al., 1997; Stassi et al., 1998; Xue et al., 1999). Individual catalytic domains or groups of domains were also studied in vitro with biochemical methods (Kumar et al., 2004). Detailed discussion of DEBS and the biosynthesis of erythromycin is found in Chapter 6 of this volume. A comprehensive description of assembly line enzymology is beyond the scope of this chapter. Various aspects are discussed both elsewhere in this volume and in many recent reviews (Fischbach and Walsh, 2006; Smith and Tsai, 2007; Van Lanen and Shen, 2008; Walsh, 2008; Weissman and Leadlay, 2005; Weissman and Muller, 2008). A representative diagram of assembly line enzymology is shown in Fig. 13.1, in this case synthesis of the microtubule stabilizing anticancer agent, epothilone, by a PKS composed of nine PKS modules and a non–ribosomal peptide synthetase (NRPS) module. NRPS modules are often found in PKS assembly lines and are also amenable to genetic engineering (see Chapter 20 of volume 458). Assembly line diagrams for other PKSs can be found in several other chapters of this volume. Although the epothilone PKS was one of the more difficult to engineer, inactivation of two of its KR domains did give some production of the expected products (Tang et al., 2005). The epothilone PKS is also a source of modules having unusual specificities that potentially can be used to

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9

13

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OH

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OH

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O 1 O

OH

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N

Figure 13.1 The PKS assembly line for epothilone biosynthesis. The six polypeptides of this megasynthase are shown at the top (EpoA to EpoF) as arrows from N- to C-terminus. The modules on these polypeptides are numbered below and the domain composition is shown schematically. The intermediate attached to the phosphopantetheine arm on the ACP domain is that expected just prior to its transfer to the downstream KS. This diagram illustrates various aspects of PKS engineering discussed in the text.

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construct novel PKSs (Menzella et al., 2005). To date there are well over a hundred modular PKS gene cluster sequences known, with more appearing nearly every week. In synthetic biology terms this represents a large collection of parts that could be used to engineer the production of novel polyketides. The number of polyketide structures that are theoretically possible from these assembly lines is huge, but this chapter is focused on engineering relative modest changes in a natural modular PKS in order to generate analogues for medicinal chemistry. The structure of each ‘‘ketide’’ unit of a polyketide is determined by the catalytic domains in each PKS module as shown in Fig. 13.2. Generating polyketide analogues involves deleting, inserting, mutating or replacing catalytic domains in a PKS module. Entire modules also can be rearranged to create a different assembly line. Feeding of chemically synthesized precursors to an assembly line engineered to accept them expands the possibilities still further. Some modular PKSs have been relatively pliable to engineering, DEBS being the prime example, but others have been difficult to engineer for reasons that are largely unknown. Unexpected results may be partly because a PKS does not always follow the strict correspondence shown in Fig. 13.2. Modules can contain domains that do not appear to be necessary based on the structure of the intermediate. For example, three modules of the epothilone PKS (L, 8, and 9) have dehydratase (DH) domains that appear unnecessary (Fig. 13.1). Module 8 also has an apparently unnecessary ketoreductase (KR) domain. The sequence of some of these apparently redundant domains suggests they are catalytically inactive, which may represent relics of a recent evolutionary event. On the other hand, domains that at first glance seem unnecessary might have a function. For example, ketoductase domains unable to catalyze ketoreduction can still epimerize the side chain on the a carbon (Keatinge-Clay, 2007). A catalytic activity that seems to be necessary also can be missing from a module. The 12,13 double bond of epothilone is predicted to arise by dehydration of the module 4 intermediate, yet module 4 contains no DH domain. Experiments have suggested that the DH in module 5 carries out this reaction (Tang et al., 2004). Occasionally, a PKS module catalyzes more than one condensation before passing its intermediate forward, a phenomenon referred to as ‘‘stuttering’’ and reviewed in Moss et al. (2004). An incomplete understanding of such deviations from colinearity or the correlation between domain organization and polyketide structure may have been the reason for the failure of some previous engineering attempts. An engineered PKS generally produces a lower titer of the analogue than the wildtype PKS produces of the original polyketide. This reduction in assembly line efficiency could be caused by disturbance of PKS structure or by the altered PKS intermediate being a poor substrate for the next module. When the downstream ketoacyl synthase (KS) domain acts as a gatekeeper, experiments have shown that catalytic efficiency can be increased by

AT specificity

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Additional Examples

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Figure 13.2 (Continued)

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replacing the KS with one having better tolerance (Chandran et al., 2006). Post-PKS reactions, such as hydroxylation, glycosylation and methylation are often critical for the potency and other drug properties of a polyketide. An analogue produced by an engineered PKS may not be processed efficiently by the post-PKS pathway, and this can present additional obstacles to be overcome. Strategies for PKS engineering fall in two broad categories. Either the sequence of the PKS genes is altered directly in the chromosome of the original producing organism or the genes are cloned, altered in vitro (or in Escherichia coli), and expressed in a suitable host. When the original polyketide producing organism can be genetically manipulated and produces reasonable titers of the natural polyketide, it is often best to do the engineering in that strain to obtain the analogue in sufficient quantity. However, expressing engineered PKS genes in heterologous hosts can have significant advantages, especially for making large libraries of polyketide analogues (McDaniel et al., 1999). Given the structural diversity of polyketides, the diverse mechanisms of their biosynthesis, and the phylogenetic diversity of organisms that express polyketide pathways, it is not possible to provide detailed PKS engineering protocols that would be useful. Instead this chapter will provide generic procedures and guidelines for increasing the chance of success.

2. AT Domain Replacement to Alter a-Carbon Substitution An a-carbon side-chain on a polyketide can be changed by replacing the AT domain in the corresponding module with a heterologous AT domain having a different substrate specificity. The methylmalonyl-CoA-specific AT Figure 13.2 The ‘‘ketide’’ unit structure determined by the domain organization of some PKS modules. AT domain substrate specificity is shown in the left column. The specific stereoisomer produced by a module having a KR, but no DH or ER, is determined by the particular KR domain. KR domain sequence motifs apparently correlated with stereochemical outcome have been identified. The example on the bottom left is a module with a methylmalonyl-CoA-specific AT and a C-methyltransferase domain, which produces an a-gem-dimethyl. The next example shows two of the four stereoisomers possible with an ethylmalonyl-CoA specific AT. The next example at the bottom shows a cis double bond arising from a module that has both DH and KR domains, the latter determining this outcome. The bottom right example shows the trapping of an enol ether by an O-methyltransferase domain. Engineering a change in ketide structure is achieved by engineering the corresponding change in the domain organization of the module.

Genetic Engineering to Produce Polyketide Analogues

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domains of the erythromycin PKS have been replaced with AT domains specific for malonyl-CoA (McDaniel et al., 1999; Petkovic et al., 2003; Ruan et al., 1997) and other substrates such as ethylmalonyl-CoA or methoxymalonyl-ACP (Kato et al., 2002; Stassi et al., 1998). The methoxymalonyl-ACP-specific AT domains of the ascomycin (FK-520) PKS of Streptomyces hygroscopicus have been replaced with malonyl and methylmalonyl transferases giving the predicted analogue (Reeves et al., 2002; Revill et al., 2002). After replacing six of the AT domains in the geldanamycin PKS with malonyl transferase domains, four gave the expected analogue (2-desmethyl, 6-desmethoxy, 8-desmethyl, and 14-desmethyl), although some also altered post-PKS reactions (Patel et al., 2004). The specificity of an AT domain, at least with respect to malonyl-CoA versus methylmalonylCoA substrates, is correlated with certain amino acid sequence motifs (Haydock et al., 1995; Smith and Tsai, 2007). In addition to AT replacement, specificity can be changed by mutating the sequence of these motifs (Reeves et al., 2001). Complete AT domain replacement is probably the strategy of choice unless it fails in a particular experiment, in which case site-specific mutagenesis provides a backup strategy.

3. Procedure for Engineering AT Replacements in the Chromosome 1. Design the AT replacement cassettes by first considering the amino acid sequence. Align the region around the AT to be replaced with heterologous AT domains having the desired substrate specificity. AT domain specificities in a given PKS are usually defined by applying the colinearity rule to the PKS sequence. Choose those AT domains that have sequences as similar as possible to the AT being replaced. An AT domain from another module of the same PKS may be used, but it is prudent to synthesize the encoding DNA with different codon choices to avoid undesired recombination events. Also use the sequence alignments to choose the junctions where the heterologous AT sequence and the native PKS sequence will be spliced together. Table 13.1 lists conserved sequence motifs found near the boundaries between PKS domains. Whether to include the sequence between the ketoacyl synthase (KS) and AT domains or the post-AT linker sequence in the replacement cannot be predicted and it is probably best to try the replacement both with and without these sequences. 2. Obtain DNA encoding the chosen heterologous AT domains either by PCR amplification or gene synthesis. Obtain the native PKS sequence upstream and downstream of the AT being replaced by PCR amplification of about 1000 base pair regions. Assembly of the AT replacement

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Table 13.1 Conserved sequence motifs between PKS domains

Region

Approx length (AA)

Boundary motifs

KS KS/AT linker AT Post-AT DH (1 and 2) KR structural ER KR ACP

430 100 330 25 240 190 320 250 80

(4–6)EPIAIV SGTNAHVIVE Not conserved VFVFPGQG GVAVDW(4–6) Not conserved YPFXXXXYWL (40–45)HXXXGXXXXP LPFAW(45–50) (6–8)RVNW EDQVAVR(14–16) (40–45)NFRDV GKIVLTXPX (4–6)LVTGG AWGXWA(75–90) (35–40)LGFDSL(35–40)

Notes: The actual sequence is that found in the erythromycin PKS. This table helps in locating these motifs in any given PKS. Some domain boundaries have no well-conserved motifs nearby, in which case the nearest motif that is conserved was chosen. The approximate number of amino acid residues between the motif and the actual boundary is given in parentheses. Precise identification of boundaries should be based on analysis of multiple PKS sequence alignments and three-dimensional homologue models.

cassettes from these three fragments (Fig. 13.3) can be done using several methods. Stitching by overlap extension (Heckman and Pease, 2007; Horton et al., 1993) is best, because it does not require the use of restriction sites at the junctions between the pieces. A proofreading polymerase, such as Phusion from Finnzymes, should be used for this method and the PCR primers should have 20-bp overlapping extensions. The ends of the cassette should have restriction sites that will facilitate cloning in the next step. 3. Clone the AT replacement cassettes into a vector that can be used for two-step double homologous recombination in the producing microorganism (Fig. 13.3). For many species of Streptomyces, either the plasmid pKC1139, which has a temperature-sensitive Streptomyces replicon, or the fC31 phage vector KC515 are excellent choices (Kieser et al., 2000). The simplest type of vector for this purpose, called a suicide vector, cannot be propagated in the polyketide producer unless it integrates into the chromosome by homologous recombination. Plasmids that contain an oriT site can often be introduced into the polyketide producing strain by conjugation from an E. coli strain. Many species of Streptomyces can only be transformed with DNA that is not methylated and the plasmid must be isolated or introduced by conjugation from a dam , dcm strain of E. coli. Site-directed mutagenesis may be required to eliminate interfering sites in the cassettes built in Step 2 before they can be cloned into the vector. 4. Introduce each AT replacement vector into the producing organism and select for colonies resistant to the marker expressed by the chosen vector.

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Vector SM US AT∗ DS or Chromosome

US AT DS

First crossover Marker resistant isolates

US AT∗ DS

SM US AT DS

US AT DS

SM US AT∗ DS

Second crossover Marker sensitive isolates

US AT DS

Original polyketide

US AT∗ DS

Polyketide analog

Figure 13.3 Engineering an AT replacement in the chromosome of a polyketide producing strain. See the text for details. The vector must have a selectable marker (SM) functional in the polyketide producing strain and can have sequences to facilitate its delivery. A cassette consisting of a heterologous AT flanked by upstream (US) and downstream (DS) homologues is built and cloned into the vector. The first and second crossovers can occur between either the US or DS homologues. If these sequences are the same size, each crossover has an equal probability of occurring at the US or DS sequence. Small paired arrows indicate where primers should bind for PCR verification that the desired crossover event has occurred.

The method used to introduce the vector DNA (transformation, conjugation or transfection) will depend on the strain and the vector as just discussed. For all commonly used actinomycete vectors, the methods are well-described in Practical Streptomyces Genetics (Kieser et al., 2000). For vectors with a temperature-sensitive replicon, the strain also must be grown at the nonpermissive temperature to ensure that chromosomal recombination is the only mechanism for antibiotic resistant colonies to arise. 5. Pick drug resistant colonies and streak for pure clones. For Streptomyces and other filamentous microorganisms that can sporulate, the spores themselves should be streaked or plated for single colonies. For organisms that grow as single cells, such as myxobacteria, cyanobacteria, or pseudomonads, this is not necessary. 6. Verify correct integration using colony PCR with primer pairs that will amplify either the upstream or downstream flanking region only when adjacent to the heterologous AT sequence and inserted at the correct chromosomal locus (Fig. 13.3). Southern blot hybridization can also be

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used for this purpose, which, although more time consuming, provides more detail about the locus following integration. 7. Grow the verified strains in liquid medium without selection for several generations to allow a second crossover event to loop the construct out of the chromosome (Fig. 13.3). For Streptomyces and other filamentous microorganisms that sporulate, a spore stock must be generated after culturing in liquid to ensure that the subsequent colonies are clonal. For bacteria that grow as single cells this should not be necessary. 8. Identify clones in which the construct has looped out. For some microorganisms counter-selectable markers can be included in the vector used in step 3, allowing direct selection for the second crossover event. However, there are few reliable counter-selectable markers for most polyketide-producing microorganisms and the usual method of finding the desired strains is by replicating colonies from plates without antibiotic onto plates with antibiotic and then hunting for the rare clones that have become sensitive. The frequency of such clones ranges between 0.1% and 1% for most actinomycetes (Kieser et al., 2000). The second crossover can occur between the sequences that recombined to give integration or between the homologues on the opposite side of the heterologous AT. Only the latter case will give the desired AT replacement. If the flanking homologues are the same size, crossing over with each occurs with equal probability and, statistically, half the antibiotic-sensitive clones will carry the AT replacement. For the myxobacteria, colonies are very difficult to replicate because the cells grow slowly and migrate on the agar surface. Since a reliable counterselectable marker has not yet been found for most myxobacteria, they are difficult to engineer by double homologous recombination. 9. Identify antibiotic-sensitive clones with the AT replacement using PCR with primers that bind just outside the upstream and downstream homologues used for recombination (Fig. 13.3). Southern blot hybridization can also be used for this. In addition, grow each isolate in production medium, extract the broth, and analyze the extracts by a liquid chromatography–mass spectrometry (LC-MS) method that will detect the original polyketide and related analogues. If the AT replacement results in a complete switch of substrate specificity, the original polyketide will be absent and one or more new compounds should be seen. The LC-MS data usually provides strong evidence that an isolate makes the predicted analogue. However, interpretation of LC-MS data can be complicated when the product of an engineered PKS is processed differently by the post-PKS tailoring enzymes (Patel et al., 2004; Reeves et al., 2004). 10. For unequivocal proof of analogue structure, the engineered strain should be grown in larger-scale cultures, the products purified (typically using C18 column chromatography), and each fully characterized using

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NMR and mass spectrometry. At this stage the purified analogues also should be evaluated for drug properties using in vitro assays. Preclinical testing in animals generally requires gram quantities of pure compound, which highlights the importance of doing the engineering in such a way as to obtain good titers of the polyketide analogues.

4. Engineering b-Carbon Processing The nascent b-ketone following a condensation is processed sequentially by a keto-reductase (KR), a dehydratase (DH) and an enoyl reductase (ER) domain, when present in the module. The first examples of PKS engineering were deletion of most of the KR domain of module 5 of the erythromycin PKS (using fortuitous restriction endonuclease sites), to obtain 5,6-dideoxy-5-oxoerythronolide B (Donadio et al., 1991) and mutation of two amino acids in the putative cofactor binding site of the module 4 ER domain to obtain D6,7-anhydroerythromycin C (Donadio et al., 1993). Subsequently the erythromycin PKS has been engineered in many different ways including replacement of sets of b-carbon processing domains with domains from heterologous PKS modules (McDaniel et al., 1999). The procedure for engineering different b-carbon processing in the chromosome of a polyketide producing strain will follow closely the steps already described for an AT replacement, except that sequences encoding b-carbon processing domains will be altered. Figure 13.4 gives examples of cassettes for ER deletion, insertion of a DH domain or replacement of one set of domains with another. Selecting the optimum points at which to join the native and heterologous sequences is aided by aligning the PKS sequences of interest and using Table 13.1 as a guide for locating domain

KS

AT

AT

ψKR

DH1 DH2

ψKR

KR

ER

KR

ACP

KS

ACP

DEBS module 1

DEBS module 4

Figure 13.4 Domain organization of PKS modules of different composition. Note where DH and ER domains are located when present. The smaller unlabeled black boxes represent flanking sequences on either side of the AT domain that are not part of the catalytic domain itself, but may have an important structural role. In the trans-AT PKSs lacking integral AT domains, these flanking sequences are still present.

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boundaries. However, not enough engineering has been done to specify the best junction points in a given situation. Engineering of b-carbon processing is complicated by the fact that the KR domain sets the chirality at both the a and b carbon atoms, with some KR domains epimerizing the a-carbon without reducing the b-ketone (KeatingeClay, 2007). Moreover, the stereochemistry set by the KR determines whether a cis or a trans double bond is formed during subsequent dehydration if there is an active DH domain in the module (Keatinge-Clay, 2007). When enoyl reduction occurs, the ER domain determines the stereochemistry at the a-carbon. A protocol for identifying the stereochemical outcome of a module from its organization and the sequence motifs in the KR domain has been presented (Keatinge-Clay, 2007). Most previous engineering of b-carbon processing was done well before this information was known, and future engineering that accounts for this aspect of assembly line enzymology should have better success at obtaining the desired analogues. Recent high-resolution crystal structure data for various portions of modular PKSs (Keatinge-Clay, 2007, 2008; Keatinge-Clay and Stroud, 2006; Khosla et al., 2007; Smith and Tsai, 2007; Tang et al., 2006, 2007), and for the entire mammalian fatty acid synthase (Maier et al., 2008) has better defined the boundaries between catalytic domains (Fig. 13.5 and Table 13.1). This structural data reveals extensive domain–domain interactions and we are just beginning to understand their importance for assembly line efficiency (Weissman and Muller, 2008). In future engineering efforts, in silico analysis of the relevant structures may allow the identification of residues in domain– domain interaction surfaces that, if mutated, might improve catalytic activity. An ER domain is inserted between two subdomains of the KR, one serving a structural role (cKR) and the other a catalytic role (KR) (Keatinge-Clay and Stroud, 2006). Although inactivation of an ER by site-specific mutagenesis was successful (Donadio et al., 1993), such a strategy could leave an ER able to bind its PKS intermediate and thus reduce assembly line efficiency. It may be preferable to delete ER domains at its boundaries, although this has yet to be tried. Insertion of an ER domain between the KR subdomains also seems likely to give the desired polyketide analogue, though it is important that the KR and DH domains generate the appropriate trans double-bond substrate for the inserted ER. The DH domain folds as two subdomains with the active site at the interface of those subdomains (Keatinge-Clay, 2008; Maier et al., 2008). Deletion of both DH subdomains at the proper boundaries should give an analogue with a hydroxyl in place of the double bond and with stereochemistry that may be predicted from the sequence of the KR (Keatinge-Clay, 2007). All known previous attempts to engineer a b-hydroxyl by inactivating a DH domain have resulted in a PKS that produced no product at all (unpublished results). However, this new information may allow such engineering to be done successfully in the future.

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ER deletion to convert a saturated bond to a double bond ψKR KR

ψKR ER

KR

ψKR KR

DH insertion to convert a hydroxyl to a double bond AT

DH ψKR

AT ψKR KR AT

DH ψKR KR

Domain set replacement to convert a carbonyl to a methylene AT

DH ψKR ER

KR ACP

AT ψKR KR ACP AT DH ψKR

ER

KR ACP

Figure 13.5 Examples of cassettes that could be used to engineer b-carbon processing. Any of these cassettes would be used as described for AT replacement (Fig. 13.2). The sequence to be deleted, inserted or replaced may be a single b-carbon processing domain or the complete set found in a given module. Insertion or replacement of several b-carbon processing domains might give a cassette too large to fit in some vectors, in which case another vector or a heterologous expression strategy would be alternatives. The symbols are defined in the text and the DH domain represents the two subdomains shown in Fig. 13.3.

Whether to delete a domain completely or to inactivate it using point mutations probably depends on the context and other factors. For example, deletion of the KR domain in DEBS module 6 gave the desired 3-keto 6-dEB analogue, but also relaxed the specificity of the AT domain in the module so that 2-desmethyl-3-keto 6-dEB was also produced (McDaniel et al., 1999). In subsequent work, various putative catalytic residues in KR6 were mutated, not only achieving production of 3-keto 6-dEB as the sole analogue, but also identifying key residues in the active site of the KR domains (Reid et al., 2003). Deletion of the KR6 domain in the geldanamycin PKS gave production of the expected product with no affect on the other catalytic domains (Vetcher et al., 2005).

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5. Engineering when only a Single Crossover Event is Possible DNA can be introduced into some myxobacteria by conjugation from E. coli and if the plasmid carries homologous sequence it can integrate into the chromosome ( Jaoua et al., 1992; Kopp et al., 2004; Xia et al., 2008). On the other hand, it is very difficult to achieve domain replacements because a good counter selectable marker is not available and colonies are difficult to replicate. Despite this, some PKS engineering may be possible. For example, a construct was devised to change the AT specificity of the loading module of the ambruticin PKS of Sorangium cellulosum So ce10 by a single recombination event (Rodriguez, unpublished work). The first ambruticin PKS gene is relatively short, encoding only a loading module consisting of KS, AT and ACP domains ( Julien et al., 2006). Thus, a plasmid was constructed that carried a cassette composed of a heterologous promoter from the epothilone PKS, an N-terminally truncated KS from the ambruticin loading module, a malonyl-CoA specific AT from a different ambruticin module, and finally the loading module ACP (Fig. 13.6). Upon transfer of this plasmid into Sorangium by conjugation from E. coli, the construct could integrate into the chromosome by recombination with the KS, the AT or the ACP sequence. By screening a sufficient number of strains, one was found in which crossover had occurred at the KS PhR E. coli vector w oriT

P KSq Sorangium chromosome

P

KSq

ATm

P

ACP

KSq

ATm

ACP

ATmm

ACP

PhR

P KSq ATmm

ACP

Figure 13.6 PKS engineering in Sorangium cellulosum using a single crossover event. The first PKS gene for ambruticin biosynthesis is shown on the Sorangium chromosome with the promoter (white P) that drives expression of the entire set of PKS genes. This first PKS gene encodes a KSq, a methylmalonyl-CoA specific AT (ATmm) and an ACP for the loading module. A construct was built such that if crossing over occurred via the KSq as shown, a functional loading module gene would be created that had its AT replaced with one specific for malonyl-CoA (ATm). The heterologous promoter ( gray P) thus would drive expression of the remaining PKS genes, but the first gene would not be translated because of an N-terminal truncation. The desired crossover event was sufficiently frequent that desired strains were identified by screening the phleomycin resistant (PhR) exconjugants of Sorangium with a PCR assay.


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sequence (Fig. 13.6) to give a strain in which the heterologous epo promoter drove transcription of the truncated loading module gene (which could not be translated into a functional protein) and the remaining downstream PKS genes, while the ambruticin promoter drove transcription of the engineered loading module gene carrying the malonyl-CoA-specific AT domain. As a result, the strain produced the ambruticin analogue predicted from the chromosomal engineering.

6. Heterologous Expression of Engineered PKS Genes If the original polyketide producing strain cannot be genetically manipulated, analogues may be obtained by expressing the engineered genes in a heterologous host. The use of a heterologous expression system also facilitates high-throughput PKS engineering, allowing many analogues to be produced in parallel. The first step is construction of cassettes to delete, insert or replace domains much as was described earlier for chromosomal engineering. In this case, however, the cassettes represent the first step in the construction of large plasmids designed for expression of the engineered PKS gene (Fig. 13.7). The remaining PKS genes, as well as genes for essential post-PKS reactions, or for synthesis of essential precursors, also must be expressed in the host to achieve production of an active analogue. The upstream and downstream sequences of the initial cassettes need not be the same length, but must have restriction sites at the ends that allow it to be inserted into the PKS gene and maintain the reading frame. The final expression constructs can be large and several methods are available to facilitate their construction. DNA encoding various sections of a PKS gene can be synthesized with desired codon usage and the required restriction sites. Cosmids can be used to increase the efficiency of introducing the large constructs into E. coli, although this requires that the construct be a size that can be packaged into the phage in vitro. The lRed/ET recombination system can be used to achieve insertion, deletion or replacement of domains directly in plasmids maintained in E. coli (Vetcher et al., 2005). Many different expression vectors are available, depending on the chosen host. The various PKS genes can be maintained on vectors that replicate autonomously or that integrate into the genome by homologous recombination or by site-specific integration. To keep the size of any one construct manageable, large PKS clusters can be expressed on separate vectors. Multiple plasmids have been used for combinatorial expression of engineered genes for the erythromycin PKS in Streptomyces coelicolor, generating a large library of 6-dEB analogues (Xue et al., 1999). Methods for the use of most actinomycete vectors are described in detail in Practical Streptomyces Genetics (Kieser et al., 2000) as well as in other chapters in this volume.

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Deleted sequence X

Y DS

US X

Y

or US

Heterologous sequence

X

DS

Y

Q

Z Engineered PKS gene Z

Q

Expression vector

Figure 13.7 Vector construction for heterologous expression of an engineered PKS gene. This diagram assumes that unique restriction enzyme sites (X, Y, Z, and Q) are conveniently located or can be introduced during stitching of the cassettes. The cassettes are made first (probably using PCR stitching by overlap extension) to join upstream (US) and downstream (DS) sequences to the heterologous sequence. Using sequential ligation reactions, the cassette is then built into the entire PKS gene and finally expressed in a chosen host.

E. coli has been used as a host to functionally express the full set of erythromycin biosynthetic genes from Saccharopolyspora erythraea (Peiru et al., 2005) and the full set of epothilone biosynthetic genes from S. cellulosum (Mutka et al., 2006). Although E. coli has the advantage of facile genetic manipulation, synthesis, and expression of large PKS genes is nevertheless quite challenging. Moreover, the level of polyketide production from E. coli has been relatively low, which limits the utility of an E. coli host for polyketide drug development. Production levels of polyketide analogues can be much higher when the host is phylogenetically close to the polyketide-producing strain. For example, several polyketides produced by various species of Streptomyces have been successfully made in host strains derived from S. coelicolor or S. lividans (Hu et al., 2005; McDaniel et al., 1999; Xue et al., 1999; Ziermann and Betlach, 1999). A strain of Streptomyces fradiae that produced almost 2 g/l of tylosin was found to be amenable to transformation by conjugation from E. coli DH5a


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and was engineered for production of 16-membered macrolide analogues (see Chapter 15 of this volume). The tylosin PKS genes (tylGI-tylGV) in the strain were deleted by double homologous recombination leaving the tylGI promoter for expression of downstream post-PKS tailoring genes (Rodriguez et al., 2003). Integration of the tylosin PKS genes at the fC31 phage attachment site restored tylosin production. Some of the 16-membered macrolide PKS genes to be expressed encoded a module that used methoxymalonyl-ACP as a precursor. Therefore, a cassette to express a set of genes for biosynthesis of this precursor was integrated at the pSAM2 att site (Rodriguez et al., 2004). Integration of the midecamycin PKS genes from Streptomyces mycarofaciens under control of the tylGI promoter into this strain gave production of midecamycin. Such engineered S. fradiae host strains were used for combinatorial expression of different 16-membered macrolide PKS genes (Reeves et al., 2004), which gave results instructive for PKS engineering in general. All 16-membered macrolide PKSs are composed of five polypeptides and each polypeptide shares significant sequence similarity with the corresponding polypeptide of the other PKSs (Fig. 13.8). Differences between the macrolactone products of these PKSs arise from differences in substrate specificities of the AT domains and a missing KR domain for the chalcomycin PKS. Presumably these differences arose by relatively recent gene conversion events. The first two genes for the chalcomycin PKS and the last three from either the spiramycin or tylosin PKS were expressed in an S. fradiae clean host. The total titer of polyketides derived from these hybrid PKSs was almost equal on a molar basis to the titer of tylosin produced by the original strain. Thus modules appear to be very tolerant of different intermediates and the low titers of an analogue after PKS engineering is most likely due to perturbation of PKS structure. Despite the subtle differences in the structures of the natural and hybrid macrolactones, however, the P450 hydroxylase encoded by tylH in the S. fradiae host would not accept the chalcolactone-platenolide hybrid macrolide as a substrate. Furthermore, the P450 hydroxylase encoded by tylI converted the ethyl side chain of this hybrid to a carboxylate, instead of the usual aldehyde. To circumvent this obstacle, chmH, a tylH homolog from the chalcomycin gene cluster was integrated at the fBT1 att site and expressed via the tylGI promoter. This gave production of a novel and biologically active tylosin analogue derived by post-PKS tailoring of the chalcolactone-platenolide hybrid macrolide.

7. Chemobiosynthesis Chemobiosynthesis is a specific type of PKS engineering coupled with feeding of chemically synthesized precursors, which are used by the PKS and incorporated at the starter end of the polyketide chain. It has

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Figure 13.8 Expressing combinations of different 16-membered macrolide PKS genes in a heterologous host. The five PKS polypeptides encoded by tylGI-V of S. fradiae, chmGI-V of S. bikiniensis and srmGI-V of S. ambofaciens are shown schematically with the differences in domains between them highlighted in black. The three natural macrolactone products, tylactone, chalcolactone and platenolide lead, respectively, to tylosin, chalcomycin, and spiramycin in the respective strains are shown below the PKS polypeptides. The chalcolactonetylactone and chalcolactone-platenolide hybrids formed by the first two chalcomycin PKS genes and the last three PKS genes for tylosin or spiramycin, respectively, are shown between the natural macrolactone structures.


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been quite successful for the erythromycin PKS ( Jacobsen et al., 1997; Ward et al., 2007) and for a 16-membered macrolide PKS (Kinoshita et al., 2001). It was also used to obtain production of epothilone C from E. coli (Boddy et al., 2004). The first module of a PKS assembly line captures a starter unit and loads it onto the KS of the first extender module. These loading modules use a variety of mechanisms. Many are minimal PKS modules with a KS domain in which the active site cysteine is replaced by another residue, often a glutamine (a so-called KSQ domain). The AT domain of this type of loading module transfers a carboxyacyl extender unit onto the phosphopantetheine arm and the KS domain catalyzes its decarboxylation, generating the starter unit to be passed downstream. The erythromycin PKS and some others have a loading module with no KS and the AT is specific for propionyl-CoA (or some other acyl-CoA). Yet another type of loading module is an adenylation-thiolation didomain, which can pick up a free carboxylic acid using ATP to adenylate the carboxylate before transferring it to the phosphopantetheine arm, as is seen for the rapamycin PKS (Goss et al., 2006). The chemobiosynthesis strategy involves the following steps: 1. The loading module or, more typically, the first extender module of the PKS is rendered nonfunctional by introducing null mutations in the KS active site, or by deleting the N-terminus of the PKS up to the beginning of the KS of the second extender module. 2. Synthetic acyl-thioester precursors (e.g., diketide thioesters of N-acetylcysteamine) are fed to the resulting mutant strain. 3. If the KS domain of the first or second extender module will accept the synthetic thioesters as polyketide intermediates, analogues are produced that have incorporated the synthetic structure.

8. Mutasynthesis Mutasynthesis is a potential strategy for generating analogues when the PKS assembly line incorporates an atypical precursor and the biosynthesis of that precursor can be blocked through gene disruption. The strategy involves the following steps: 1. The biosynthetic pathway to the precursor is deleted by introducing null mutations into one or more of the genes encoding enzymes in that pathway. Such mutations can be introduced by classical mutagenesis, though it is more common to delete genes by double homologous recombination using methods similar to those described earlier for AT replacement. The resulting strain should no longer produce the polyketide product.

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2. For the strategy to work, it must be possible to restore polyketide production by adding the precursor to the culture. This shows that the precursor can be taken up by the cell, is not degraded, and is a substrate for the PKS assembly line. 3. If step 2 is successful, then chemically synthesized precursor analogues can be fed to the mutant strain, some of which may be taken up and incorporated to give corresponding polyketide analogues. The range of precursor analogues that can be accepted as substrates and incorporated varies for different PKS assembly lines. A successful example of this strategy was the production of nonquinone ansamycin analogues of geldanamycin by engineering a strain of Streptomyces hygroscopicus (Kim et al., 2007). The first KS domain of the geldanamycin PKS is primed with the starter unit 3-amino-5-hydroxybenzoic acid (AHBA), which requires a special pathway for its synthesis. When genes encoding enzymes of this pathway were disrupted, geldanamycin production was abolished unless the strain was fed AHBA. The feeding of several AHBA analogues to this mutant gave production of corresponding nonquinone geldanamycins. Several other examples of mutasynthesis have been reviewed (Weissman, 2007).

9. Gene Knockouts to Obtain Analogues It is not always necessary to engineer the PKS in order to obtain useful polyketide analogues. Inactivation of genes for discrete enzymes in a polyketide pathway often gives analogues. In the case of the complex PKS assembly line for ambruticin biosynthesis in S. cellulosum, this not only gave analogues, but also provided insight into ambruticin biosynthesis ( Julien et al., 2006). The procedure resembles that described for AT replacements earlier, but is much simpler. 1. A region of a gene to be knocked out is amplified by PCR. The region should be just downstream of the start codon and about 40% upstream of the distance to the stop codon. The primers should introduce an inframe stop codon at both ends of the PCR product to ensure that upon integration neither copy will be functional. 2. The PCR products are cloned into a pUC19-derived plasmid that contains a marker for selection in the polyketide producing strain (a bleomycin resistance gene is usually best for myxobacteria, while genes for resistance to thiostrepton, kanamycin, or hygromycin can be used with many actinomycetes) and the R6K oriT sequence, if the plasmid will be delivered by conjugation.


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3. The resulting plasmid is introduced by conjugation or transformation and colonies resistant to the marker antibiotic are selected. These colonies usually represent the desired gene disruption, but should be verified by colony PCR. Some of the S. cellulosum So ce10 strains carrying disruptions generated by the above procedure produced novel ambruticin analogues that had bioactivity. One of the genes, ambM, encoded a discrete C-methyltransferase that acted in concert with a specific module in the assembly line as part of an unusual PKS reaction sequence. Knockout of this gene gave an analogue missing a C-methyl group known to be derived from the methyl carbon of methionine ( Julien et al., 2006).

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C H A P T E R

F O U R T E E N

Design and Synthesis of Pathway Genes for Polyketide Biosynthesis Salvador Peiru´, Hugo Gramajo, and Hugo G. Menzella Contents 1. Introduction 2. Redesign of PKS Genes to Accommodate Unique Restriction Sites Flanking Individual Components and for Efficient Expression in E. coli 3. Validation of Synthetic PKS Gene Design 4. A Rapid Assay to Identify Productive Combinations of PKS Modules 5. Assembly of Larger Polyketide Synthases Using Information Gained with the Bimodular Assay 6. Design and Construction of Synthetic Operons for the Expression of Sugar Pathway Genes References

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Abstract In this chapter we describe novel methods for the design and assembly of synthetic pathways for the synthesis of polyketides and tailoring sugars. First, a generic design for type I polyketide synthase genes is presented that allows their facile assembly for the expression of chimeric enzymes in an engineered Escherichia coli host. The sequences of the synthetic genes are based on naturally occurring polyketide synthase genes but they are redesigned by custom-made software to optimize codon usage to maximize expression in E. coli and to provide a standard set of restriction sites to allow combinatorial assembly into unnatural enzymes. The methodology has been validated by building a large number of bimodular mini-PKSs that make easily assayed triketide products. Learning from the successful bimodules, a conceptual advance was made by assembling genes encoding functional trimodular enzymes, capable of making tetraketide products. Second, methods for the rapid assembly and exchange of sugar pathway genes into functional operons

Instituto de Biologıá Molecular y Celular de Rosario (CONICET), Facultad de Ciencias Bioquı´micas y Farmaceúticas, Universidad Nacional de Rosario, Rosario, Argentina Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04614-X

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are described. The approach was validated by the assembly of the 15 genes for the synthesis of mycarose and desosamine in two operons, which yielded erythromycin C when coexpressed with the corresponding PKS genes. These methods are important enabling steps toward the goals of making designer drugs by polyketide synthase and sugar pathway engineering and, in the shorter term, producing by fermentation advanced intermediates for the synthesis of compounds that otherwise require large numbers of chemical steps.

1. Introduction Type I polyketide synthases (PKSs) that determine the biosynthesis of valuable natural products like erythromycin, epothilone, tacrolimus and many others consist of multiple catalytic domains forming a protein ‘‘assembly line.’’ The domains are associated in modules that determine the structure of a 2-carbon extender unit of the polyketide chain. Thus, the sequence of domains in the assembly line, directly reflecting the DNA sequence of the corresponding genes, determines the structure of the polyketide product. This simple logic led to genetic engineering of PKSs to make ‘‘unnatural natural products’’ (Walsh, 2003, 2004; Weissman and Leadlay, 2005) as shown in the early 1990s by research groups at Stanford and Cambridge (Cortez et al., 1995; Kao et al., 1994). They demonstrated that domains and modules could be reorganized to create novel enzymes capable of making a variety of unnatural compounds. The initial approaches in the field of genetic engineering of polyketides included adding, deleting or interchanging domains or whole modules in a PKS aimed to alter the number and structure of the 2-C units built into a polyketide. Whereas these might appear simple tasks for current genetic engineering technologies, they are encumbered by difficulties, and PKS engineering is far from being established in the high-throughput format required to provide the necessary flow of unnatural compounds in order to obtain ‘‘hits’’ with new biological properties. One problem is that most natural polyketide producers are not readily amenable to a complete range of genetic methodologies without considerable time-consuming effort. Most of them are slow-growing compared with commonly used laboratory strains, often requiring more than a month for a round of genetic manipulation, and many target organisms are so far uncultured. Therefore, there is a need for a ‘‘universal host’’ in which PKS components from different sources can be expressed and where combinatorial experiments can be performed rapidly and ideally in a high-throughput fashion (Pfeifer and Khosla, 2001).

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A second problem is that natural PKS gene clusters are extraordinarily long (up to 200 kb), usually with a high GþC content and with highly repetitive coding sequences for successive modules. Rarely are there conserved restriction sites to facilitate domain or module exchange. Thus, there is a need for molecular biological tools to accelerate the assembly of chimeric PKS genes in order to find productive combinations. Expressing PKS genes in a more tractable host can circumvent some of these difficulties. Recently, Escherichia coli has been successfully engineered to produce polyketides (Murli et al., 2003; Pfeifer and Khosla, 2001). Since its genetic manipulation is so straightforward, it was chosen as a host for developing new methods for combinatorial biosynthesis of polyketides in the work described here. To avoid the problems of manipulating natural PKS genes, a first step was to develop a technology for facile synthesis of error-free 5 kb sequences of DNA—the average for complete PKS modules (Kodumal et al., 2004; Reisinger et al., 2006). Next, a generic design of PKS modules was created with unique restriction sites flanking their components to allow exchange as building blocks in an approach that has been dubbed ‘‘Legoization’’ (Menzella et al., 2005; Sherman, 2005). Finally, methods were developed to rapidly assemble and express simple chimeric PKSs and to screen for their activity in order to identify productive combinations to be used in the construction of larger, designer enzymes (Menzella et al., 2005; Menzella and Reeves, 2007). Another important aspect to be considered during polyketide engineering is that, in order to be fully active, many of these molecules require the attachment of sugar moieties on their core structure. The presence of these sugars, as well as other post-PKS modifications, is often essential to impart or enhance the biological activity of the molecule. Thus, heterologous expression of PKSs also requires the incorporation of complete glycosylation pathways into the host, to achieve the synthesis of a fully decorated molecule. The possibility of altering the glycosylation pattern of polyketides through genetic engineering is another promising tool for diversification of these compounds. Sugar biosynthesis genes are normally located within the polyketide gene cluster (Peiru´ et al., 2005). The heterologous expression of these sugar pathways typically requires the cloning of individual genes and their assembly into operons in order to have a controlled expression system under inducible promoters. As with PKS modules, several issues need to be considered for the construction of sugar operons. Here we describe practical experience with these genetic engineering tools. The ultimate goal is to produce complex molecules by creating multidomain PKSs combined with sugar tailoring genes to be used directly as drugs or as lead compounds for chemical optimization. Meanwhile, at a more modest level, even the combination of a few PKS modules can

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produce molecules with multiple chiral centers (up to two per module) that are difficult to obtain by ordinary chemical synthesis (McDaniel et al., 2005). The production of such molecules made by fermentation to be used as intermediates for the synthesis of complex structures can avoid large numbers of chemical steps and significantly reduce manufacturing costs. Therefore, engineering synthetic polyketide synthases is a promising strategy to produce novel molecules for multiple applications.

2. Redesign of PKS Genes to Accommodate Unique Restriction Sites Flanking Individual Components and for Efficient Expression in E. coli The strategy involves the synthesis of redesigned genes containing a set of standard unique restriction sites at the borders of each PKS component, including domains, linkers and modules, to allow their facile interchange. To allocate these restriction sites, sequence conservation at domain edges was analyzed (Fig. 14.1) by aligning the amino acid sequences of more than 300 PKS modules. When the identified conserved sequences were reversetranslated to all possible DNA sequences, a conserved six-base-pair restriction site could be found in some cases. In modules where that sequence was not present, amino acids were changed to a sequence resembling one present in at least some PKSs or which was already known, from prior experimental evidence, not to disturb functionality. For the ketosynthase (KS) domain, an MfeI site was incorporated near the 50 edge by using bases 2 to 7 of nine bases coding for the amino acid sequence PIA at the beginning of this domain. Of the KS domains analyzed, 70% needed no change in amino acid sequence, and 30% required only conservative changes from V, L, or M. On the 30 edge of all KSs, the conserved GT can be encoded by the sequence for a KpnI site. At the 30 side of the acyl transferase (AT) domain, a PstI site was placed where PstI and XhoI have been used successfully by others (McDaniel et al., 1999; Oliynyk et al., 1996). A highly conserved sequence occurs in 90% of the modules as R-Y/F-W. The codons specifying the two amino acids C-terminal to the W of this motif were modified to introduce the PstI site coding for LQ. The dipeptide at this position is variable in the native sequence. However, in more than half the cases the first amino acid is L, and in 80% of these the second is hydrophilic or even charged. Thus, the LQ dipeptide was likely to be well tolerated. No constant sites were found allowing independent exchange of the reductive domains ketoreductase (KR), dehydratase (DH), enoyl reductase (ER). For modules containing a KR domain, an AgeI site could be

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–54

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Spe I eryM1 eryM2 eryM6 sorM6 epoM7 eryM5 gelM3 lepM10 rifM5 rapM2 epoM3 eryM3 rapM3 pikM6 Design

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Kpn I GT GT GT GT GT GT GT GT GT GT GT GT GT GT GT

Pst I RV W LE RF W LL RYW LA RF W LD RC W IE RYW LP HYW LT RF W LE HF W LS RYW LE RYW IE RYW LQ RY W LR S YW I S LQ

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XbaI/SpeI L A AE L G G L D AW L I G QQ I L R DS L L L TDV Y L ERL I R TEL I D GE L

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L R DE L G G L G EL I F T L L S QA L E L R AR L V G L - DE L F T I S DE L A E SS

Figure 14.1 Conserved amino acid sequences at domain borders and the chosen design to generate the desired restriction sites for domain assembly. Restriction site positions are referenced to module 4 of DEBS. Numbering begins at the first residue of the motif EPIAIVon the N-terminal edge of the KS domain. Sequence homologues for 14 of the 140 modules used for the generic module design are shown. eryM3 and rapM3 contain an inactive KR domain. ery, erythromycin; sor, soraphen; epo, epothilone; gel, geldanamycin; lep, leptomycin; rif, rifamycin; rap, rapamycin; pik, pikromycin. (From Menzella et al., 2005. Reproduced with permission from Nature Journals.)

incorporated at its 50 edge to encode the TG dipeptide found in 132 of 137 native KR domains analyzed, while in the remaining five this domain was predicted to be inactive. When required, DH/ER pairs could be added, deleted, or exchanged using the PstI and AgeI sites. Finally, since the 50 boundary of the acyl carrier protein (ACP) domain is poorly defined, no unique restriction site could be placed there, so domain exchange required the use of an upstream unique site. Except for the minimal module containing only AT, KS, and ACP, this required comobilization of the reductive domains. At the 30 end of the module, an XbaI site was incorporated at a well-defined position next to the carboxy side of the ACP. There are two conserved L residues at positions 36 and 40 downstream of the active site serine of ACPs. The codons of the two amino acids at positions 41 and 42 were changed to the XbaI sequence coding for SS, already known not to affect functionality (Gokhale et al., 1999). To facilitate gene expression in E. coli, all the PKS genes were codonoptimized. For the gene design, a software package called GeMS ( Jayaraj et al., 2005) was developed, comprising a set of programs to predict restriction sites, perform codon optimization for expression in any host, assign or

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eliminate restriction sites, separate long sequences into smaller fragments to facilitate synthesis, carry out Tm and stem-loop determinations, and design oligonucleotides to be used as raw material. GeMS accepts DNA or protein sequences and its output is a complete design report with a list of oligonucleotides to be used in the gene synthesis process. Using GeMS a complete collection of PKS components was designed and later synthesized, including 50 extender modules, 5 loading modules, 5 thioesterases, 30 intrapeptide linkers, and 20 pairs of docking domains, totaling 400 kb of synthetic DNA. All the building blocks contain the standard set of restriction sites to facilitate the assembly of hybrid PKSs. For codon optimization, a codon table was used representing the E. coli genome. For each amino acid in the target protein, the software randomly selects a codon in accordance with its distribution in the table. The optimized genes thus contain codons in the same proportion as in E. coli. Although all the gene design work described here was made using GeMS, many other free webbased programs like, for example, Gene Designer (http://www.dna20.com/ tools/genedesigner) and Optimizer (http://genomes.urv.es/OPTIMIZER) may serve well for this purpose.

3. Validation of Synthetic PKS Gene Design In the first attempt to validate the generic PKS gene design, the 31-kb gene cluster encoding the 6-deoxyerythronolide B (6dEB) synthase (DEBS) was assembled and expressed in an E. coli strain engineered for polyketide production (Menzella et al., 2006). In the redesigned cluster the total set of 15 restriction sites were accommodated flanking the individual domains (Fig. 14.1). For this, 20 amino acid substitutions were required to insert some of the MfeI and PstI sites and all of the SpeI/XbaI fusions. The PIA tripeptide needed for the MfeI site is naturally present in five of the six DEBS modules, and module 1 (Mod1) has a conserved PVA. The LQ dipeptide required for the PstI site is found naturally only in DEBS Mod3; four modules have L followed by E (Mod1), L (Mod2), P (Mod5) or A (Mod6), and Mod4 has an unusual PR sequence found in only 2% of PKS modules. The SpeI/ XbaI fusion used to connect all redesigned modules results in an SS dipeptide that is unnatural to all modules. Overall, in the final redesigned DEBS, 13 of the 20 amino acid changes accommodated the SpeI/XbaI junction, one accommodated the MfeI site of Mod1 and six accommodated the PstI sites in Mod 1, 2, 4, 5, and 6. For the rapid assembly of building blocks to create chimeric open reading frames a suitable method is described below. In the described gene design, NdeI and XbaI sites flank the coding region of the N-terminal PKS components (LM and docking domains); the coding regions of internal modules are flanked by SpeI and XbaI sites;


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and the coding region of C-terminal components (docking domains and TE) are flanked by SpeI and EcoRI sites. All the PKS components were synthesized and cloned into pUC-based vectors containing an additional NotI site upstream of the coding region. 1. Create a multicopy vector with a polylinker containing NdeI-NotISpeI-EcoRI sites. 2. Initiate the assembly by inserting the C-terminal component of the ORF into the SpeI-EcoRI sites of the vector. 3. Clone the internal components sequentially as NcoI-XbaI fragments into the NcoI-SpeI sites of the vector. This SpeI/XbaI ligation destroys both sites, allowing the repetitive use of these enzymes to insert as many components as required. 4. Finally, clone the N-terminal component as an NdeI-XbaI fragment into the NdeI-SpeI sites to complete the assembly of the ORFs, which can be mobilized as NdeI-EcoRI fragments into expression vectors or used to assemble operons. To determine whether these amino acid changes introduced to accommodate the standard set of restriction sites affected function of the enzymes, protein expression levels of the three synthetic DEBS ORFs as well as 6dEB production were compared with those of the native sequences using the following method: 1. Introduce expression vectors for synthetic PKSs by co-transformation into E. coli K207-3[BL21△prpBCD::T7prom prpE, T7prom accA1pccB, T7prom sfp]. This strain is a derivative of BL21(DE3) with the propionate utilization operon (prpRBCD) and the methylmalonyl-CoA decarboxylase gene (ygfG) deleted and with chromosomal copies of sfp (phosphopantetheine transferase from Bacillus subtilis), prpE (endogenous propionyl-CoA ligase), and accA1/pccB (propionyl-CoA carboxylase from Streptomyces coelicolor) under the control of T7 promoters (Murli et al., 2003). Feeding propionate to this strain results in the accumulation of methylmalonyl-CoA, a polyketide synthase substrate produced by wildtype E. coli at very low levels. 2. Grow the resulting colonies in 3 ml LB containing appropriate antibiotics for selection at 37 to an OD600 ¼ 0.5. 3. Induce with IPTG (0.5 mM), and a mixture of sodium glutamate (50 mM), sodium succinate (50 mM), and sodium propionate (5 mM). 4. Incubate the induced cultures at 22 for 72 h with agitation. 5. Centrifuge 10 min at 10,000 g; remove the supernatant for polyketide analysis and cell pellet for protein expression analysis. 6. Extract cell-free supernatants (1 ml) twice with an equal volume of ethyl acetate. Dry the extracts in vacuo and dissolve in 1 ml of methanol. Analyze for 6dEB production as described in Menzella et al. (2006).

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7. Re-suspend cell pellets in 1 ml 20-mM Tris, 150 mM NaCl, pH 7.5, containing 1 tablet of CompleteTM EDTA-free protease inhibitor cocktail (Roche) per 50 ml. 8. Disrupt by sonication, and centrifuge at 14,000g for 3 min. 9. Analyze soluble and insoluble fractions on NuPAGE Novex 3 to 8% Tris-Acetate gels (Invitrogen). Gels can be stained using Simply Blue SafeStain (Invitrogen) and analyzed using a UVP Bioimaging System with Labworks 4.0 software (UVP, Inc.). The synthetic components of DEBS ORFs were expressed using three compatible vectors (ColE1 for DEBS1, P15A for DEBS2, and RSF1010 for DEBS3), all under control of a T7 promoter. When individually expressed in the engineered E. coli strain K207-3, each codon-optimized ORF produced about fivefold more protein than its native counterpart, as did the complete set of three coexpressed synthetic ORFs. However, no polyketide product could be detected from the three coexpressed synthetic ORFs, while the wildtype PKS gene set produced 20 mg/l of 6dEB in shake-flask fermentations. To identify the defective module(s), chimeras of wildtype and synthetic ORFs were constructed by cotransformation of combinations using the three-plasmid system. Of the three possible chimeras containing one synthetic and two natural ORFs, only the combination containing synthetic DEBS2 produced no 6dEB. Since the synthetic DEBS2 contained the nonconserved PR-to-LQ mutation to accommodate the Mod4 PstI site, we converted the sequence back to PR. With this change, synthetic DEBS2 combined with DEBS1 and DEBS3 produced 6dEB, but at a level far below that obtained with the wildtype sequences. Further studies showed that expression of synthetic DEBS from the strong T7 promoters led to a significant reduction in the expression of the accessory genes accA and pccB required to produce methylmalonyl CoA in the E. coli K207-3 strain, and this could presumably account for the low levels of 6dEB observed. When expression of the three synthetic DEBS genes was driven by the lacUV5 promoter, three- to four-fold less PKS protein was observed than when expressed under control of the T7 promoter, but both the production levels of 6dEB and the expression of the accA and pccB genes were comparable to those observed with the wildtype sequences. In another validation example, the entire synthetic epothilone gene cluster—epoA, epoB, epoC, epoD, epoE, and epoF—and the C-terminal thioesterase were assembled and expressed in the E. coli K207-3 strain (Mutka et al., 2006). The expected products, epothilone C and D, were obtained, indicating that the 29 amino acid changes needed to insert the restriction sites into the redesigned genes could be tolerated to obtain a catalytically active PKS complex of approximately 2 MDa (monomeric mass). As in the case of DEBS, titers were improved when gene expression


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was down-regulated by replacing the strong T7 promoters by an alternative, in this case by the PBAD arabinose-inducible promoter. In summary, with the sole exception of the PstI site in DEBS module 4, all the amino acid changes introduced to accommodate the restriction sites in the more than 70 kb of PKS genes for DEBS and epothilone were well tolerated. Strong evidence was also obtained to suggest that weaker promoters are needed to balance expression of the codon-optimized genes with that of the accessory proteins required for polyketide production, while stronger promoters may be necessary to achieve balance when the high-GC native genes are used.

4. A Rapid Assay to Identify Productive Combinations of PKS Modules In spite of initial excitement, in more than 15 years of PKS engineering only a few successful attempts to create functional unnatural module–module interfaces using wildtype PKS sequences have been reported (Gokhale et al., 1999; Kim et al., 2002; McDaniel et al., 1997; Ranganathan et al., 1999; Watanabe et al., 2003). Since unsuccessful attempts to create hybrid PKSs are rarely published, it is hard to anticipate how often two unrelated PKS modules could be successfully connected to yield the expected polyketide. Thus the possibilities and limitations of the combinatorial approach remain unclear. In order to address this fundamental issue, an in vivo assay was developed to rapidly screen interactions between unrelated PKS modules (Menzella et al., 2005). Here, two expression vectors with compatible origins of replication and different selectable markers capable of expressing equivalent amounts of protein from arabinose-inducible promoters were used (Fig. 14.2A). The ‘‘donor’’ vector contains the loading module (LM) of DEBS and the C-terminal docking domain of DEBS1 (DCeryM2), separated by unique XbaI and SpeI sites, so any synthetic module can be inserted into this plasmid in one step to create a fusion ORF of the class LMery-Module-DCeryM2. The ‘‘acceptor’’ vector contains the DEBS2 N-terminal linker (LNeryM3) and the DEBS thioesterase (TE), separated by unique MfeI and SpeI sites. Any synthetic PKS module can be cloned into this plasmid to create a second type of fusion ORF designated LNeryM3-ModuleTEery. In this in vivo assay, the two fusion proteins are coexpressed and the rapid detection of the predicted triketide product by LC/MS/MS reveals productive module–module interactions. To validate the assay 154 mini-PKSs of this kind were assembled, introduced into E. coli K207-3 (Murli et al., 2003) and the resulting clones cultured and tested for protein expression and production of the expected compounds using the following method:

A LM

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1. Introduce the two expression vectors by co-transformation into E. coli K207-3. 2. Grow the resulting colonies in 3 ml LB containing antibiotics for selection at 37 to an OD600¼0.5. 3. Induce with IPTG (0.5 mM), arabinose (2 mg/ml) and a mixture of sodium glutamate (50 mM), sodium succinate (50 mM), and sodium propionate (5 mM). 4. Incubate the induced cultures at 22 for 72 h with agitation. 5. Centrifuge 10 min at 10,000g; remove the supernatant for polyketide analysis and cell pellet for protein expression analysis. 6. Acidify culture supernatants with phosphoric acid to pH 2.5. 7. Analyze for triketide production y LC/MS/MS as described in Ashley and Carney (2004). 8. Protein expression can be assessed following the same procedure described above for DEBS. Remarkably, almost half of the 154 bimodules produced the expected polyketide and each tested module functioned as a donor or acceptor in one or more contexts, indicating that all redesigned modules were catalytically competent, and further validating the generic PKS gene design. Interestingly, all the modules processed unnatural substrates, confirming their relaxed specificity (Gokhale et al., 1999; McDaniel et al., 1999; Ranganathan et al., 1999). Production of a triketide by the hybrid mini-PKSs must have involved overcoming several barriers. Potential obstacles to transfer of the propionyl moiety from the LM to the KS of each attached module include the foreign intrapeptide linkers, potential substrate intolerance by noncognate KS domains, or a requirement for possible ACP-KS interactions. Some of the unnatural fusions used the intrapeptide linker naturally associated with the module, and some others had the linker that normally separates LM from eryM1. Except for eryM1, none of the modules naturally accept and process a propionyl group, but all module fusions to LM produced the expected triketide in some bimodular combinations. The obtained results show that there is considerable flexibility in the intrapeptide linkers used to separate the LM and the first module, and the KS in all tested modules fused to the LM can elongate the propionyl moiety. Therefore, all donor modules form Figure 14.2 (A) The two classes of expression plasmid used to test bimodular interactions in E. coli: pAng vectors contain a CloDF13 replication origin, a streptomycin resistance selection marker and a PBAD promoter to drive expression of LM-Module-DCeryM2 ORFs. pBru vectors contain a ColE1 replication origin, a carbenicillin resistance selection marker and a PBAD promoter to drive expression of DNeryM3-Module-TE ORFs. (B) The two types of vectors used to express trimodular PKSs: pAngII vectors are similar to pAng, but DCeryM2 was replaced by DCeryM4. pCot plasmids are similar to pBru, but they express ORFs of the class DNeryM5-Mod-LI-Mod-TE. (From Menzella et al., 2007. Reproduced with permission from Elsevier.)

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an acyl-ACP at the C terminus that can donate the diketide to a subsequent module. Transfer of the diketide through the hybrid interface to the KS of the acceptor module requires physical proximity of the ACP and KS, and possible protein–protein interactions between them (Wu et al., 2002); and elongation requires substrate tolerance by and catalytic activity of the KS (Watanabe et al., 2003). Since about half of the bimodules produced the predicted polyketide, and the modules were always connected by the eryM2-eryM3 linkers, this pair serves well as generic linkers to connect unrelated modules. The results of this study provided a clear indication of the possibilities of the PKS combinatorial biosynthesis approach since, to the best of our knowledge, this is the first analysis of the productivity of a large number of random module–module combinations. Encouragingly, it seems that unrelated PKS modules can be successfully connected to create functional pairs in many cases. On the other hand, the yields of polyketides obtained from the hybrid PKSs are typically at least one order of magnitude lower than those obtained from the natural bimodular pairs (Menzella et al., 2005). Importantly, the chosen assay can be easily adapted to a high-throughput format to rapidly screen for the most productive unrelated combinations of modules. In the above tests, all the acceptor modules processed a particular diketide from some donor modules but not from others, so recognition and elongation of the incoming diketide by the acceptor KS was unlikely to explain the failures. We attributed them instead to the absence of an appropriate ACP-KS protein–protein interaction (Wu et al., 2002). To address this issue, KS domains were combinatorially changed to activate inactive module pairs and optimize the yield of a desired polyketide product (Chandran et al., 2006). The unique MfeI-KpnI restriction sites flanking the KS domain in all the synthetic modules made this a simple task (Fig. 14.2). The feasibility of the approach was validated by resuscitating nine previously inactive bimodular PKSs. Such activation of hybrid modular interfaces by KS replacements provides a powerful tool to engineer synthetic PKSs. Of particular interest is the use of this procedure to incorporate into assembly lines unusual modules with rare activities that cannot be replaced easily just by seeking a more adaptable natural module with the same activity.

5. Assembly of Larger Polyketide Synthases Using Information Gained with the Bimodular Assay Polyketides with biological activity are much more complex than the triketides produced by the bimodular PKSs described above, and biosynthesis of such molecules requires the assembly of synthetic enzymes


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containing multiple modules with several hybrid interfaces. Thus, it is imperative to employ the information gained with the simple test system to assemble larger PKSs. For this, a conceptual advance was made by developing the concept of ‘‘connectivity’’ to allow the overlapping of functional bimodules to build active trimodular PKSs. Connectivity describes the situation in which module B in a given bimodule extends the ketide offered by module A, and in another bimodule, module C extends the ketide offered by module B; then a trimodular PKS made by connecting module A to module C via module B should also be active. The library of bimodules previously described was constructed at random, and gave a success rate of 45%. Constructing trimodules by simply adding another module randomly should have had, at most, a 20% success rate. A recent publication (Menzella et al., 2007) showed that, using data from the bimodular library and the connectivity principle, an additional functional heterologous interface was constructed with a remarkably higher success rate. To prove the concept of connectivity, an expression system was developed to express trimodular PKSs (Fig. 14.2B). In this case a two-plasmid system was again used for the expression. The first plasmid encoded a fusion of the class LM-Mod-DC and the second plasmid a chimeric protein comprising DN-Mod-LI-Mod-TE. Given that the erythromycin LM and TE are functional in many heterologous bimodular PKS constructs, these domains were used for all the trimodular PKSs. The docking domains derived from eryM2 (DCeryM2) and eryM3 (DNeryM3) were successful at forming functional interactions between most single modules tested. However, we observed that the larger protein encoded by bimodular constructs in the pCot vector were better expressed when the N-terminal linker of eryM5 (DNeryM5) was used instead of DNeryM3. Using the unique restriction sites in our generic design, we examined several intrapeptide linkers to join the two modules in pCot, including the linker naturally associated with the first module and the one naturally present upstream of the N-terminus of the second module in this plasmid as well as foreign intrapeptide linkers (LIeryM6, LIeryM1) between the two modules, creating two unnatural junctions. The ability of bimodular constructs in pCot to interact and produce a polyketide was assessed and shown to be similar in all cases. Thus, it seems that all the tested linkers are equally efficient in connecting the modules. Several pairs of functional bimodules were overlapped to create 54 trimodular PKSs flanked by the DEBS LM and TE. Remarkably, 52 (96%) of the synthetic enzymes formed the expected tetraketide product. Twelve of the trimodules contain contiguous extender modules in the order in which they occur in their native PKS, but in 42 there are no naturally contiguous modules; nevertheless, 40 (95%) of these also produced the expected polyketide. Interestingly, in terms of substrate specificity these results suggest that the portion of the upcoming polyketide recognized by a

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given module is in most cases the extension added by the previous module regardless of the length and complexity of the polyketide chain. The rational assembly of trimodular PKSs represents an important step toward the goal of making bioactive molecules by genetic engineering, both complete polyketides that cannot be isolated in significant quantities, and novel, ‘‘designer’’ molecules. The rationale should, for example, be applicable to building unnatural PKSs from scratch by combining individual artificial modules, as described here, as well as to adding or replacing ketide units in preexisting chains. It also provides a rational approach for ‘‘stitching’’ two or more strings of naturally contiguous modules together to make novel or inaccessible large polyketides. However, since yields of polyketide products were seen to decrease as a function of the number of unnatural junctions introduced, the efficiency of product formation will probably continue to decrease when further modules and heterologous interfaces are added. This limitation might be circumvented by using only the highest yielding trimodules as components, or by incorporating strings of as many naturally contiguous modules as possible when designing long chimeric PKSs so as to reduce the number of unnatural interfaces

6. Design and Construction of Synthetic Operons for the Expression of Sugar Pathway Genes The increasing knowledge and technologies related to PKS engineering represent a significant advance toward the ultimate goal of synthesizing rationally designed polyketides in E. coli. However, as in many naturally occurring polyketides, such novel compounds might also require several post-PKS biosynthetic steps in order to become bioactive. These polyketide ‘‘decoration’’ steps, which generally involve complex glycosylation processes, are absent in E. coli and thus need to be incorporated in this heterologous host in combination with the PKSs to produce fully decorated polyketides. Glycosylation of polyketides has been validated in E. coli, through the production of the antibiotic erythromycin C (EryC) (Peiru´ et al. 2005). This process involves the attachment of the deoxysugars TDP-L-mycarose and TDP-D-desosamine to the aglycon 6dEB (firstly hydroxylated to EB), to yield erythromycin D (EryD), which is further hydroxylated at C12 to obtain EryC. Two operons were constructed for this purpose, in order to have separate control of the two sugar biosynthetic pathways. The complete conversion of 6dEB to EryC requires functional expression of 17 genes with a variety of catalytic activities, comprising TDP-sugar biosynthesis, glycosyltransferases and hydroxylases. In this seminal work, all the


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genes were individually amplified by PCR from Micromonospora megalomicea chromosomal DNA and a simple method was designed to allow sequential cloning of genes to build synthetic operons by adding suitable flanking restriction sites and translation signaling sequences (Fig. 14.3). NdeI and SpeI sites were introduced immediately 50 and 30 of each gene, respectively, to enable cloning into a special expression plasmid named pD2, containing XbaI-RBS-His5 coding sequence-NdeI-SpeI. The N-terminal His5 fusion, which can be omitted, introduces a short codon-optimized sequence that we have observed improves gene expression in most cases. It also provides a tag for further protein expression analysis and/or protein purification by means of anti-His5 antibodies. The flanking XbaI and SpeI sites that result from cloning the PCR-amplified ORFs into this plasmid permit the sequential cloning of genes as XbaI/SpeI fragments into the SpeI site of the acceptor vector, resulting in a growing string of genes separated by a minimal sequence. The obtained operons can be further mobilized as XbaI/SpeI fragments to vectors with different promoters, selection markers, or replication origins if required. A detailed method is described below. 1. Amplify by PCR the individual genes to be included in the synthetic operon. For the design of the upper primer, engineer an NdeI site overlapping the translational initiation codon (changing GTG start codons to ATG when necessary). For the design of the lower primer, engineer a SpeI site immediately downstream of the stop codon. 2. Create a special vector for the sequential cloning of the PCR products. For this purpose the pET24a expression vector (Novagen) can be modified by replacing the BamHI site in the polylinker by a SpeI site, and by deleting the 21-bp region between the XbaI site and the RBS. Alternatively, a His5 coding sequence can be added upstream of the NdeI site. 3. Clone each PCR-amplified sugar biosynthetic gene as an NdeI/SpeI fragment into the same sites of the modified pET24a vector. 4. Initiate the assembly by sequentially cloning each gene as XbaI/SpeI fragments into the SpeI site of the acceptor vector, making sure that the inserted gene is properly orientated. The intergenic XbaI/SpeI ligation destroys both sites, allowing the repetitive use of the remaining 30 SpeI site to insert as many components as required. 5. Finally, subclone the resulting operon as an XbaI/SpeI fragment into the expression vector with the appropriate promoter, selection marker and replication origins. Using this method, two operons were built: the ‘‘mycarose’’ operon, for the conversion of 6dEB up to 3-a-mycarosyl-EB (MEB), and the ‘‘desosamine’’ operon, for the conversion of MEB to EryC. The activity of both operons was found to be optimal in E. coli cultures grown at 22 , when induced in the presence of a plasmid for the overexpression of the E. coli chaperones GroES/GroEL, which improve solubility of the heterologous

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eI

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Figure 14.3 Cloning strategy for the construction of sugar biosynthesis operons. Sugar genes are PCR-amplified, with flanking NdeI and SpeI sites, and cloned into pD2. This expression vector contains a ColE1replication origin, a kanamycin resistance selection marker, and adds an RBS and a His5 N-terminal fusion to the cloned gene, whose expression is driven by a PT7 promoter. The acceptor vector (pSugar 1) is digested with SpeI, and the second gene in the nascent operon is introduced as an XbaI/SpeI fragment. The process can be repeated with the resulting pOperon plasmid in a recursive fashion, until the entire operon is constructed.


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proteins. Biosynthesis of EryC from propionate in K207-3 required cloning of the mycarose and desosamine operons into compatible vectors (RSF1030 and CloDF13, respectively), to allow coexpression with the three plasmids for the synthesis of DEBS and chaperones containing ColE1, RSF1010, and P15A origins of replication (Peiru´ et al., 2005). Codon-optimized versions of the genes involved in these pathways were not tested since the expression levels were satisfactory in all cases. Using a wildtype background the glycosylation yields achieved in the initial experiments were low (30 kb), to be integrated into the K24-1 strain with high fidelity. Insertion of the FC31 attB site is a valuable tool with broad application in actinomycetes and other microorganisms, including Pseudomonas putida (Martinez et al., 2004). Similarly, a clean-host version of the industrial S. fradiae strain, K159-1, was constructed by deleting the tylGI-V genes encoding the tylosin PKS and leaving the tylGI promoter for expression of the deoxysugar genes downstream of the tylGV locus. Introduction of an attB site into S. fradiae was unnecessary because it already contains one. For both strains, and in agreement with results with S. coelicolor, it was found that only the native PKS promoter was capable of sustaining high polyketide titers, suggesting that the expression level of PKS genes is the primary factor associated with the overproducing capability of both strains. Recently, it was demonstrated that in the Sac. erythraea overproducer strain this effect was correlated with a higher expression level of BldD, a recently identified positive regulator of the erythromycin biosynthetic genes (Chng et al., 2008; Lum et al., 2004). The potential for using a previously optimized industrial microorganism as a host for improved polyketide production capitalizes on the intrinsic overproduction properties of the strain and on the extensive process development work already done to achieve optimal production conditions. The advantage of using such hosts for engineered polyketide overproduction was demonstrated by the enhanced production (50-fold improvement in titer) of erythromycin analogues expressing genetically engineered erythromycin PKSs in Sac. erythraea K24-1 compared to the production in a nonoptimized host like S. lividans K4-114 (Rodriguez et al., 2003). In addition, Sac. erythraea provided the ability to generate biologically active glycosylated derivatives of the structurally modified 6-dEB aglycones, and is therefore especially useful for overproduction of valuable erythromycin analogues or other compounds requiring these deoxysugars. The potential for using S. fradiae 159-1 was demonstrated by expressing a vast array of native and hybrid 16-membered macrolide PKSs from the tylosin, chalcomycin, midecamycin, and spiromycin gene clusters, thanks to the expanded precursor supply capability of the engineered strain, as discussed below (Reeves et al., 2004; Rodriguez et al., 2003, 2004; Ward et al., 2004).

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5.3. Optimization of conjugation protocols for industrial strains 5.3.1. Sac. erythaea 1. Harvest spores from one or two plates, filter through cotton, spin down, resuspend in 1 ml of 20% sterile glycerol and freeze at –80 . 2. Inoculate 20 ml of spore suspension (107 to 108 UFC) into 5 ml 2 YT and incubate at 30 with shaking for 1 to 2 h. 3. Spin down cells (recipient culture) and resuspend in a final volume of 100 ml. 4. Transform competent E. coli ET12567/pUB307 cells with an oriTcontaining vector and select for the marker of the incoming plasmid. ET12567 is used as donor strain since Sac. erythraea has a lower efficiency of transformation with methylated DNA. 5. Inoculate a colony into 5 ml LB containing chloramphenicol, kanamycin, and the antibiotic used to select for the oriT-containing plasmid and grow overnight at 30 . 6. Use half of the culture to check the integrity of the plasmid by restriction digestion. Modular PKSs usually contain large DNA fragments that can be highly homologous, which have the chance to rearrange in a recombination-proficient (RecAþ) strain like ET12567. 7. Spin down the rest of the cells, wash in 5 ml LB, and spin down again (donor culture). 8. Resuspend the donor culture with 100 ml of the recipient culture. 9. Spread on R5 plates containing 50 mg/ml nalidixic acid and incubate at 37 for 20 h. 10. Overlay with 3 ml of Soft Nutrient Agar (SNA) or sterile H2O containing 1 mg nalidixic acid and 1.5 mg of apramacyn and continue incubation at 34 for 48 h. 11. Streak potential exconjugants into new plates of selective media. 5.3.2. Cross-streak method for E. coli–streptomyces conjugation 1. 1 109 spores are pregerminated by heating for 10 min at 65 in 2 YT and then spread on conjugation plates (SFM for S. coelicolor and AS1 for S. fradiae). 2. Individual colonies of E. coli (ET12567/pUB307 for S. coelicolor or DH5a/pUB307 for S. fradiae) transformed with an oriT-containing vector are streaked on top of the conjugation plates containing the pregerminated spores and incubated 16 to 24 h at 30 . 3. Overlay with 3 ml of soft nutrient agar (SNA) or sterile water containing 1 mg nalidixic acid and 1.5 mg of the appropriate antibiotic and continue incubation at 30 . Since some Streptomyces colonies adhere to agar plates by digesting the agar (only if they contain the DagA agarase of S. coelicolor

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A3(2)), E. coli colonies carrying different plasmids can be streaked on the same plate without mixing them after overlaying with the selective antibiotic. 4. Streak potential exconjugants into new plates of selective medium. This new conjugation protocol saves 1 day of work, since no overnight E. coli culture is necessary. Besides, this faster technique allows the conjugation of nearly 30 to 50 different colonies per plate, avoiding growing each E. coli clone individually before conjugation. The method is a potentially high-throughput technique that can be adapted for metagenomic research.

5.4. Optimizing polyketide precursors supply Identification of the precursors required for the biosynthesis of a particular polyketide, as well as the enzymatic pathways that generate them, are important steps that help to optimize the heterologous production of these natural compounds. Therefore, genetic manipulation of these pathways, either by overexpressing the native genes or by introducing heterologous genes for the reconstitution of new metabolic pathways, should increase the spectrum of PKS substrates in any particular host. Here we will describe the most important known polyketide precursor pathways and how these pathways were used for optimizing heterologous polyketide production. 5.4.1. Enhance precursor supply The classical extender units used for polyketide biosynthesis are malonylCoA and methylmalonyl-CoA, which generally come from primary metabolic pathways. The acyl-CoA carboxylase complexes (ACCase), characterized in S. coelicolor and other actinomycetes, are able to carboxylate acetyl- and propionyl-CoA to produce malonyl- and methylmalonyl-CoA, respectively (Diacovich et al., 2002; Rodriguez and Gramajo, 1999; Rodriguez et al., 2001). Alternatively, methylmalonyl-CoA can also derive from the Krebs intermediate succinyl-CoA through methylmalonyl-CoA mutase (MCM) (mutA y mutB) and methylmalonyl-CoA epimerase (meaB) (Korotkova and Lidstrom, 2004). Ethylmalonyl-CoA, used for the biosynthesis of several important 16-membered macrolide compounds like tylosin, can also be generated by carboxylation of butyryl-CoA by a substrate-tolerant ACCase complex (Diacovich et al., 2002). Crotonyl-CoA reductase encoded by the ccr genes is involved in the generation of butyryl-CoA from two acetate units, which has been shown to enhance ethylmalonyl-CoA incorporation and production of ethyl-substituted erythromycin derivative when the ccr gene from Streptomyces colinus was expressed in an engineered Sac. erythraea strain (Stassi et al., 1998). Other less common extender units like ethylmalonyl-CoA, methoxymalonyl-ACP, hydroxymalonyl-ACP, and

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aminomalonyl-ACP come from special pathways which sometimes are clustered with the polyketide biosynthetic genes (Wu et al., 2000). Knowledge from primary metabolic pathways can be used in order to enhance precursor supply. For example, over-expression of the acetyl-CoA carboxylase complex in S. coelicolor improves the concentration of malonylCoA and, as a consequence, increases the actinorhodin titer in laboratory conditions (Ryu et al., 2006). In Sac. erythraea, it was found that the methylmalonyl-CoA used for erythromycin biosynthesis comes from different pathways depending on the carbon source used (Reeves et al., 2006). In carbohydrate-based media methylmalonyl-CoA mutase acts as a drain of methylmalonyl-CoA, but in oil-based media MCM provides the methylmalonyl-CoA. This information was used to increase erythromycin production in an oil-based fermentation medium by engineering the methylmalonyl-CoA route through duplication of the methylmalonyl-CoA mutase operon (Reeves et al., 2007). 5.4.2. Incorporation of new precursor pathways Although S. coelicolor naturally produces malonyl-CoA–derived polyketides, it also has the capacity to produce several mg/l of 6-dEB, a propionyl-CoA/ methylmalonyl-CoA–derived polyketide. This is achieved by using, most probably, substrates generated by the endogenous ACCase complexes (Rodriguez and Gramajo, 1999). Moreover, Lombo et al. (2001) also demonstrated that by introducing into S. coelicolor CH999 a heterologous precursor pathway consisting of the MatB and MatC enzymes, a malonylCoA synthase, and a putative dicarboxylate transport protein able to convert exogenous malonate and methylmalonate into their corresponding CoA thioesters, the capacity of this strain to produce 6-dEB was improved threefold. In addition, the capacity of CH999 for precursor supply was expanded further by introducing a set of genes involved in the biosynthesis of the extender unit methoxymalonyl-ACP from the ascomycin gene cluster (Wu et al., 2000) and used for production of new 6-dEB analogues (Kato et al., 2002). Tylosin is a16-membered macrolide that requires malonyl-, methylmalonyl-, and ethylmalonyl-CoA precursors for its biosynthesis (Cundliffe et al., 2001). The polyketide production capacity of the industrial strain of S. fradiae mentioned above was also expanded by introducing the methoxymalonyl-ACP biosynthetic pathway of FK520 from Streptomyces hygroscopicus (Wu et al., 2000). The resulting strain can produce the four different precursors, malonyl-CoA, methylmalonyl-CoA, ethylmalonylCoA, and methoxymalonyl-ACP, and produced more than 2 g/l of a 16-membered macrolide through heterologous expression of native and hybrid PKSs from the tylosin, chalcomycin, spiromycin, and midecamycin gene clusters (Reeves et al., 2004; Rodriguez et al., 2003, 2004; Ward et al., 2004). These experiments not only allowed the production of large


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amounts of new 16-memberd macrolides but also helped to study the flexibility of tailoring enzymes toward alternative aglycones. The modifications included glycosylation by TylMII, which adds mycaminose to the 5-hydroxyl group (Gandecha et al., 1997), oxidation at C-20 by the cytochrome P450 TylI (Merson-Davies and Cundliffe, 1994), and attachment of mycarose by TylCV (Bate et al., 2000). To introduce the genes for the methoxymalonyl-ACP pathway into S. fradiae, a new integrative vector containing elements of the pSAM2integration system was developed. This new vector is compatible with FC31 integrative plasmids and integrates with high efficiency at a specific tRNA locus. A set of five genes, fkbG-K (homologues to asm13-17), from the FK520 gene cluster of S. hygroscopicus, was assembled in plasmid pKOS24417a, carrying a site-specific integration system (the integrase intpSAM2 gene and the attachment site attPpSAM2 from pSAM2), and a selection marker compatible with pSET152-derived plasmids. In this plasmid the intpSAM2 gene is under the control of the intFC31 promoter rather than its natural promoter. This plasmid is compatible with FC31-derived vectors, permitting integration of PKS genes and precursor pathway genes on separate plasmids. Thus, integration of the fkbG-K cluster and the full mdm PKS genes in this host produced 1 g/l of 3, 4-didespropionylmidecamycin A3 (Fig. 15.2). Using the same expression system a hybrid chalcomycin-spiramycin PKS operon in S. fradiae yielded a new series of hybrid 16-membered macrolides (Reeves et al., 2004).

6. Recent Developments for the Production of Polyketides in Nonactinomycete Bacteria 6.1. E. coli as heterologous host E. coli is, without doubt, the best-studied bacterium from the physiological point of view. Besides, this strain also possess the best developed molecular biology protocols, the most abundant genetic tool box, and well-optimized fermentation protocols, which allow it to be grown to very high optical density (>150 OD590 nm) in less than 48 h in a simple and inexpensive medium. Furthermore, E. coli also has the best developed plasmid systems and expression vectors, which have been extensively used for the efficient production of recombinant proteins for industrial and pharmaceutical applications. Finally, it has long been used as a model for genetic and metabolic engineering. All these factors made E. coli one of the best candidates to be developed into a surrogate host for the expression of heterologous biosynthetic pathways for the production of industrially or pharmaceutically relevant molecules.

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However, a drawback is that E. coli is not an abundant producer of secondary metabolites, and so it probably lacks the intracellular machinery needed to make a wide range of natural products. E. coli, however, does produce some natural products in the terpenoid and NRP families, but not polyketides (Altincicek et al., 2001). Therefore, to convert E. coli into a producer of heterologous polyketides several limitations had to be addressed and overcome. The most important issues to be considered where: (1) to have a good expression/vector system that could accommodate and transcribe large fragments of DNA, (2) to find conditions in which the transcribed genes were efficiently translated into active (properly folded) proteins, (3) to find and introduce a tolerant phosphopantetheinyltransferase (PPTase) for efficient post-translational modification of ACPs, (4) to introduce the biosynthetic pathways needed to generate the appropriate substrates for the PKSs introduced, and (5) to coexpress resistance mechanisms toward the compound to be made if this presented toxicity or antibacterial activity. The first polyketide to be made in E. coli was 6dEB (Pfeifer et al., 2001). To achieve this, a BL21-derived E. coli strain was extensively engineered. A PPTase gene from Bacillus subtilis, sfp, was inserted into the prp operon responsible for propionate catabolism in E. coli, deleting the prpRBCD genes but leaving prpE under the inducible T7 promoter. The PrpE protein is necessary to convert exogenous propionate into propionyl-CoA, the starter unit of 6dEB and a substrate of the propionyl-CoA carboxylase (PCC) that converts it into (2S)-methylmalonyl-CoA, used as extender unit by DEBS for 6dEB biosynthesis, but propionate catabolism was abolished, thus avoiding wastage of this building unit. The accA2-pccB genes, coding for the two subunits of the PCC of S. coelicolor (Rodriguez and Gramajo, 1999), and the three debs genes were introduced on selectable pET plasmids (Novagene), allowing the successful production of 6dEB in E. coli (Pfeifer et al., 2001). In order to identify and optimize parameters that affect polyketide production in engineered E. coli, three independent pathways were later investigated for supply of the extender unit (2S)-methylmalonyl-CoA. These studies showed that the PCC pathway was predominant, as indicated by greater flux of 13C-propionate into 6dEB through the PCC pathway, compared to the S. coelicolor malonyl/methylmalonyl-CoA ligase (matB) pathway or the Propionibacterium shermanii methylmalonyl-CoA mutase/ epimerase pathway (Murli et al., 2003). The high GC content of the DNA of actinomycete PKS genes leads to an inappropriate bias in codon usage that usually causes poor translation of the proteins. To overcome this issue, extra copies of the rare AGG and CCC codons can be helpful (Schenk et al., 1995). Alternatively, a method for the semisynthetic synthesis of long and accurate DNA sequences could be used (Kodumal et al., 2004). The method enables the manipulation of the


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codon bias and also deletion or insertion of restriction sites as desired without changing the encoded protein sequence. The applicability of this idea to natural product research was shown by the total synthesis of the complete debs genes cluster comprising 32 kb of contiguous synthetic DNA using conventional cloning methods. The synthetic gene cluster was finally expressed in E. coli and the corresponding product 6dEB was produced. This method is extremely powerful and provides the possibility to generate a tool box of several synthetic PKS modules from different biosynthetic gene clusters, which can then be combined to generate novel hybrid pathways to produce new compounds. In recent studies, coexpression of two biosynthetic gene clusters responsible for the production of deoxysugars, TDP-L-mycarose and TDP-D-desosamine, were also reconstituted in E. coli and when coexpressed with the debs genes, the P450 auxiliary proteins and the erythromycin resistance ermE gene, the potent antibiotic erythromycin C was produced (Peiru et al., 2005). Moreover, by carrying out metabolic engineering of three endogenous pathways that lead to the synthesis of TDP sugars in E. coli, we have greatly improved the intracellular levels of the common deoxysugar intermediate TDP-4-keto-6-deoxyglucose, resulting in increased production of the heterologous sugars TDP-L-mycarose and TDP-D-desosamine, both components of medically important polyketides. Bioconversion experiments carried out by feeding 6-deoxyerythronolide B (6-dEB) or 3-a-mycarosylerythronolide B (MEB) demonstrated that the genetically modified E. coli B strain was able to produce 60- and 25-fold more erythromycin D (EryD) than the original strain K207-3, respectively (Peiru et al., 2008). Several other natural compounds have now been successfully made in E. coli. One remarkable achievement was the production of epothilones C and D (Mutka et al., 2006). Here, several important genetic manipulations were made both to optimize expression of the substrate biosynthetic pathways and to enhance heterologous expression of the EpoD protein (765 kDa). For this, several genetic manipulations were done to split these proteins into two polypeptides that helped protein solubilization and allowed protein reconstitution. Also the use of pBAD promoters, inducible by arabinose, codon optimization and chaperone coexpression strategies were used to successfully produce the final compound. Other examples of heterologous biosynthesis of the products of PKS/ NRPS hybrids in E. coli, like yersiniabactin (Pfeifer et al., 2003) and the NRPS-like product echinomycin (Watanabe et al., 2006) have been reported.

6.2. Pseudomonas putida as heterologous host Unlike most secondary metabolite producing hosts, such as streptomycetes, pseudomonads grow easily and rapidly in culture and their metabolic versatility has transformed them into very useful organisms from the

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biotechnological point of view. As a heterologous host for the production of natural compounds, P. putida has several advantage compared with other systems. For example the high-GC genomic DNA content and the codon usage makes it more suitable than E. coli for the expression of actinomycete and myxobacterial genes. It also has a very promiscuous PPTase that allows it to activate PKSs and NRPSs from myxobacteria and Streptomyces spp. (Finking et al., 2002; Gross et al., 2005). Furthermore, because of the lack of intrinsic pathways for natural products, it is less likely that the expression of heterologous natural compound pathways could suffer substrate competition. P. putida has been successfully used for the production of myxochomide S, a compound produced by the myxobacterium Stigmatella aurantiaca (Wenzel et al., 2005) and for expression of a silent type III polyketide synthase, which was identified in the genome sequence of Sorangium cellulosum So ce56. The expression of this gene allowed the corresponding product flaviolin to be identified (Gross et al., 2006).

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C H A P T E R

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In Vitro Analysis of Type II Polyketide Synthase Wenjun Zhang and Yi Tang Contents 368 371 371 372 373 376 376 381

1. Introduction 2. Expression and Purification of Type II PKSs 2.1. Escherichia coli as a host for protein expression 2.2. ACP expression and modification 2.3. Streptomyces as a host for PKS expression 3. In Vitro Activity Assays 3.1. Minimal PKS activity assays 3.2. Starter unit synthesis and incorporation assays 3.3. Assays of tailoring enzymes responsible for aromatic polyketide scaffold formation: KR, ARO, and CYC 3.4. Assays of downstream tailoring enzymes Acknowledgments References

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Abstract Bacterial type II polyketide synthases (PKSs) are responsible for the biosynthesis of therapeutically important aromatic polyketides, including daunorubicin and tetracycline. Discovery and cloning of the type II, actinorhodin biosynthetic gene cluster from Streptomyces coelicolor marked the beginning of a new era in natural product biosynthesis. While genetics has played an indispensable role in understanding the programming rules of this unique family of natural product biosynthetic pathways, in vitro reconstitution and characterization of individual protein components have provided important insights into their mechanisms and specificities. In this chapter, we will provide an overview of in vitro analysis of type II PKSs in recent years. We will first describe the expression and purification of individual type II PKS components from different sources. This will be followed by a detailed description of the most commonly employed assays used to study different components of the type II PKSs

Department of Chemical and Biomolecular Engineering, University of California-Los Angeles, Los Angeles, California, USA Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04616-3

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Wenjun Zhang and Yi Tang

1. Introduction Aromatic polyketides (type II polyketides) constitute an important family of structurally diverse natural products (O’Hagan, 1991), including tetracyclines, anthracyclines (e.g., daunorubicin), aureolic acids (e.g., mithramycin), tetracenomycins, angucyclines (e.g., jadomycin and pradimicin), benzoisochromanequinones (e.g., actinorhodin and frenolicin), and so on (Fig. 16.1). Aromatic polyketides have a large range of medicinal applications, including the treatment of infectious diseases and cancer. For example, anthracyclines rank among the most important anticancer drugs in the market today (Batist, 2001); and tetracyclines and their semisynthetic derivatives are well-known broad-spectrum antibiotics with effective antimicrobial properties (Chopra and Roberts, 2001). Most known aromatic polyketides are produced by actinomycetes via type II polyketide synthases (PKSs) (Hertweck et al., 2007). The biosynthesis of type II polyketides has been studied extensively at the genetic level (Hopwood, 1997), starting with the cloning of the biosynthetic gene cluster of the model compound actinorhodin (Malpartida and Hopwood, 1984). Since then, genetic manipulations in both native producing hosts and in heterologous Streptomyces hosts (McDaniel et al., 1993), have led to the functional assignment of the enzymes involved in the many biosynthetic pathways. While genetics has played an indispensable role in revealing the programming rules of type II PKSs, a thorough understanding of the catalytic mechanisms and protein–protein interactions requires in vitro analysis. In contrast to type I PKSs in which catalytic domains are assembled linearly in multimodule megasynthases, type II PKSs are composed of dissociated enzymes. Each type II PKS gene cluster consists of genes encoding a minimal PKS and a collection of tailoring enzymes. The minimal PKS includes the ketosynthase (KS or KSa), the chain-length factor (CLF, alternatively referred to as KSb), and the acyl carrier protein (ACP) (Hertweck et al., 2007) (Fig. 16.2). The KS and CLF form a strongly associating heterodimer which is the site of the Claisen-like condensation of malonyl-ACP extender units. In addition, the malonyl-CoA:ACP acyltransferase (MAT) can be shared between fatty acid synthase (FAS) and PKS (Summers et al., 1995). The minimal PKS synthesizes a highly reactive polyb-ketone backbone, and the structural basis of the growing chain extruding in the active site of KS-CLF has been resolved (Keatinge-Clay et al., 2004). Interestingly, malonyl-CoA is used exclusively as the extender unit among all known type II PKSs, an invariant feature that is controlled by the highly stringent MAT (Koppisch and Khosla, 2003). Immediate tailoring enzymes, including ketoreductase (KR), cyclase (CYC), and aromatase (ARO), can regiospecifically reduce, cyclize, and dehydrate the polyketide chain,

O OH

OH

O

O O

O HO O

HO HO

O

O

O

OH O OH

O

OH

O NH2

H

H HO OH N Oxytetracycline O

OH

D O

OH

OH HN O

OH

O HO O

O

O O HN

OH O

HO HO

OH Daunorubicin

HO

OH

O

O

OH O

O

HO

OH OH Actinorhodin

O

COOH

O

R

O OH

O

O

N H

O

OH OH

O

Jadomycin B

Pradimicin

O

COOH

OH

HO OH

O OH

O

OH

O

Tetracenomycin

O

O NH2

O

OH O

O

O

O

OH O

OH O OH D

O

O

OH

O OH

HO

Mithramycin

O

O OH

O O

OH

OH

O

O

O

Frenolicin

O

R1128a: R=Me R1128b: R=Et R1128c: R=iPr R1128d: R=Pr

Figure 16.1 Examples of polyketides synthesized by type II PKSs.

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Minimal Type II PKS:

KS

CLF

MAT

ACP

Decarboxylative Priming O

SH

O

HO

S-CoA

ACP

O

S

HO

MAT

KS

O

CLF

O

-CO2

ACP

S−Enz

Chain Elongation =

ACP

O

O

-O

ACP

MAT

S-CoA +

O HS

O

ACP KS

O

S

O O

O-

KS

O

S

Growing Polyketide Chain

O

S

−

−S

O O

O

O

SH N N H H OH Phosphopantetheine arm

ACP

KS

S

S

KS

S O-

=

ACP

CO2 +

S+ HS−CoA

O O P O O-

ACP

+ O

HS

O

Figure 16.2 Minimal PKS catalyzed decarboxylative priming and chain elongation in poly-b-keto backbone formation of type II polyketides. KS, ketosynthase; CLF, chainlength factor; ACP, acyl carrier protein; MAT, malonyl-CoA:ACP acyltransferase.

respectively, to afford the polycyclic scaffolds of the aromatic polyketides. Subsequently, downstream tailoring enzymes such as methyltransferases, oxygenases, and glycosyltransferases can decorate the scaffold to yield a bioactive natural product (Rix et al., 2002). The structural diversities among aromatic natural products are derived from choices of starter units, variations in poly-b-ketone chain length, the regioselective cyclization steps and tailoring modifications. In vitro analysis of type II PKSs can be challenging for several reasons. First, a functional type II PKS requires simultaneous reconstitution of multiple enzymes in the same reaction, especially when working with the minimal PKS. Controlling the concentrations of the individual enzymes, their cofactors and precursors is therefore crucial to both product turnover and product structure. Determining the rate-limiting steps among the multitude of reactions is also important for analyzing the kinetics of the PKSs. Second, the poly-b-ketone backbone is highly unstable and spontaneous cyclization occurs rapidly under aqueous conditions. This makes resolving the enzymology of the minimal PKS and the immediate tailoring enzymes difficult. Third, many important cyclic intermediates of aromatic PKS pathways are inaccessible or highly insoluble, complicating biochemical characterization of downstream tailoring enzymes. Spontaneous oxidation of polycyclic phenols occurs rapidly to give quinone shunt products that are not substrates of the enzyme of interest. In the last 10 years, many of the challenges have been resolved and nearly all components of type II

In Vitro Analysis of Type II Polyketide Synthase

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PKS have been reconstituted in vitro. The ensuing biochemical analyses have provided significant insights into the mechanism and specificity of the enzymes. This article will first describe the expression and purification of individual type II PKS components. This will be followed by a list of the most commonly employed assays used to analyze and reconstitute the PKS enzymes.

2. Expression and Purification of Type II PKSs 2.1. Escherichia coli as a host for protein expression E. coli is the most widely used expression host for bacterial proteins. Commercially available BL21(DE3) strain, in combination with pET vectors, are routinely used to express proteins containing N-terminal, C-terminal or both N- and C-terminal hexahistidine tags. Cloning of the target PKS gene can be readily accomplished by using the AT-rich restriction sites, such as NdeI, EcoRI, and HindIII, in vectors such as pET28 or pET21. These sites are absent from most PKS genes, which are generally of high GC-content. To compensate for the difference in codon usage between E. coli and actinomycetes, BL21-derived Rosetta strains (Novagen) that overexpress tRNAs for some codons such as CCC and CGG that are rare in E. coli while abundant in actinomycetes can be used. E. coli has been successfully used for expression of many type II PKS components, including ACP and numerous tailoring enzymes. General protein expression and purification procedures in our laboratory are described here (Zhang et al., 2007). The pET vector containing the target gene is introduced into E. coli BL21(DE3) or BAP1 strain by electroporation. A single colony is picked and grown in 5 ml starter culture of LB plus antibiotics at 37 overnight. This is used to inoculate 500 ml of large-scale LB culture (1:500 inoculation ratio), and the cells are shaken at 250 rpm at 37 until an OD600 of 0.4 to 0.6 is reached. At this time the cells are incubated on ice for 10 min, and then induced with 0.1 mM isopropyl thio-b-D-galactoside (IPTG) for 16 h at 16 . The inoculation ratio, IPTG concentration, induction time and temperature can be optimized for different proteins. The cells are then harvested by centrifugation (3500 rpm, 10 min, 4 ), re-suspended in 30 ml lysis buffer (20 mM Tris-HCl pH ¼ 7.9, 0.5 M NaCl, 10 mM imidazole, 2 mM DTT) and lysed using sonication on ice. Cellular debris is removed by centrifugation (15,000 g, 1 h, 4 ). Ni-NTA agarose (Qiagen) resin is added to the supernatant (2 ml/l of culture), and the solution is stirred at 4 for at least 2 h. The protein resin mixture is loaded into a gravity flow column and proteins are purified with increasing concentration of imidazole (up to 250 mM) in buffer A (20 mM Tris-HCl pH ¼ 7.9, 0.5 M NaCl, 5 mM imidazole, 2 mM DTT). The cell

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debris pellet is re-suspended with 8 M urea, frozen in liquid nitrogen, thawed on ice for 5 min, and centrifuged and analyzed for the presence of insoluble proteins. All fractions are checked by SDS-PAGE. Purified proteins are concentrated and buffer exchanged into buffer B (50 mM TrisHCl, pH ¼ 7.9, 2 mM EDTA, 2 mM DTT, 10% glycerol) with Centriprep filter devices (Amicon Ultra, Millipore). Protein concentrations are determined by the Bradford assay (Biorad). The eluted protein from a Ni-NTA column is about 50 to 90% pure, depending on the expression levels of individual proteins. In addition to the hexahistidine tag, the maltosebinding-protein (MBP) affinity tag is occasionally used to enhance the solubility of proteins in E. coli (Piel et al., 2000). The pMAL vectors are used together with the E. coli K12 TB1 strain, and an amylose resin affinity column is used to purify the fusion protein using standard protocols (New England Biolabs). To further purify the recombinant PKS proteins, ion-exchange chromatography is normally the second column of choice in our laboratory. Proteins obtained after Ni-NTA and ion-exchange column can achieve greater than 95% purity. Alternatively, gel filtration column chromatography is adopted by some laboratories to perform the post Ni-NTA purification (Shen and Hutchinson, 1996; Zawada and Khosla, 1999). A typical elution condition for the Sepharose column is 25 mM Tris HCl buffer (pH 8.0), 150 mM NaCl, 1 mM DTT with a flow rate of 0.5 ml/min. For example, using gel-filtration chromatography, the gris ARO/CYC and tcm ARO/CYC (partial TcmN) were determined to be monomers, while full length TcmN and act KR eluted as dimers (Zawada and Khosla, 1999). Various PKS components can also be loaded together onto the column to check the possibility of complex formation.

2.2. ACP expression and modification The small ACP (10 to 12 kDa) is an integral part of the minimal PKS and its expression warrants a separate section. Most of ACPs produced in BL21 (DE3) are in the apo form since the PKS ACPs are not substrates of the E. coli holo-ACP synthase. A notable exception is ZhuN from the R1128 pathway, which is produced in the holo form in BL21(DE3) (Meadows and Khosla, 2001). Coexpression of Sfp during ACP expression has led to the direct recovery of holo-ACPs from E. coli (Khosla et al., 1999). In a metabolic engineering approach, Khosla’s group has modified the BL21 (DE3) strain to contain a chromosomal copy of sfp under control of the T7 promoter (Pfeifer et al., 2001). The resulting BAP1 strain is able to produce nearly all type II ACPs in the holo form. A caveat of the BAP1 system is that if an ACP is highly overexpressed (>20 mg/l), the coexpressed Sfp level can be insufficient to phosphopantetheinylate all of the apo-ACP, resulting in a mixture of apo and holo forms.


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The complete conversion of apo-ACP to holo-ACP can be carried out in vitro using Sfp (Lambalot et al., 1996). A typical reaction buffer contains 75 mM MES-acetate (pH 6.0), 10 mM MgCl2 and 5 mM DTT. The molar ratios of CoA, apo-ACP, and Sfp are maintained at 5:1:0.1. The reaction is incubated at 30 overnight and holo-ACP can be purified using anionexchange chromatography (type II holo-ACPs elute at 350 mM NaCl). Furthermore, Sfp can utilize various acyl-CoAs and synthesize acyl-ACPs from apo-ACPs. To prepare acyl-ACP, DTT is omitted from the reaction buffer and 5% tetrabutylammonium chloride can be added to increase the solubility of acyl-ACP when medium- to long-chain fatty acyl-CoA is used. The modified ACP can be confirmed by mass spectrometry (MALDI MS or ESMS). Radiolabeled acyl-CoA can also be used to convert apo-ACP into radiolabeled acyl-ACP, which can then be purified by a PD-10 desalting column and concentrated with a 5000 MWCO filter device. The final concentrate can be applied to a SDS-PAGE gel and the amount of radioactivity can be quantified by autoradiography.

2.3. Streptomyces as a host for PKS expression E. coli is usually the first choice as a host for the heterologous expression of proteins; however, many type II PKS components are completely insoluble or inactive when expressed in E. coli. The most glaring representative of this list is the KS-CLF heterodimer of the minimal PKS, which has always been present as 100% inclusion bodies in E. coli despite repeated attempts by numerous groups. Therefore, Streptomyces coelicolor and Streptomyces lividans are widely used hosts for functional expression of KS-CLF and other enzymes that are not accessible via E. coli. These strains are genetically and physiologically closely related to the native hosts of most type II PKS, and can be manipulated with a powerful collection of genetic tools (Hopwood et al., 1985; Kieser et al., 2000). DNA transformation in these hosts can be performed with relatively high efficiencies, and several selection markers are available. In addition, convenient E. coli–Streptomyces shuttle vectors have been developed, facilitating the DNA manipulation in E. coli and protein expression in Streptomyces (Fig. 16.3). One of the best known shuttle vectors is pRM5 (McDaniel et al., 1993), which has served as the parent vector for numerous PKS expression constructs. It contains the following essential features: it has a ColE1 replicon for replication in E. coli and a SCP2* replicon for propagation in Streptomyces; it has an E. coli selection marker, bla, and a Streptomyces selection marker, tsr (thiostrepton resistance); the gene of interest is cloned under the control of the actI promoter, which is regulated by actII-ORF4 present on the same plasmid. A high level of PKS gene expression is observed at the beginning of the stationary phase of Streptomyces growth. In pRM5, the gene of interest is usually cloned between unique

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PactI Pac I

Nsi I Xba I Eco RI b-lac

act KR actII-ORF4 fd term

colEI ori tsr

pRM5

∼14 kb

SCP2∗

SCP2∗ ori

RBS ermEp∗

Nde I Hin dIII Eco RI Bam HI b-lac

pIJ101 ori

pUWL201PW

pUC18 ori

∼ 6.4 kb

ter

tsr Figure 16.3 Examples of E. coli^Streptomyces shuttle vectors. b-lac, Amp resistance gene; tsr, thiostrepton resistance; RBS, ribosome-binding site.

sites PacI (downstream of the actI promoter and preferred for the 50 end of the gene) and EcoRI (or NsiI, XbaI). A ribosome-binding site (RBS) is inserted upstream of the gene of interest via the 50 PCR primer (Zhang et al., 2006). The commonly used sequence of RBS is GGAGGAGCCCATATG (RBS is italicized). The shuttle vector portion of pRM5 is 14 kb in size, which may bring challenges for efficient molecular cloning.


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Digested vector mix is usually purified by ethanol precipitation without gel purification, and vortexing and harsh pipetting are avoided. The pRM5derived protein expression vector can be introduced into the expression strain (either S. coelicolor CH999 or S. lividans K4-114) (McDaniel et al., 1995; Ziermann and Betlach, 1999) using PEG-mediated protoplast transformation. Smaller Streptomyces expression vectors have been reported, such as pUWL201PW (6.4 kb) and pPWW50-Gen Poly (6.5 kb) (Doumith et al., 2000). pUWL201PW has bla and tsr selection markers, and ori derived from pUC18 and pIJ101 for E. coli and Streptomyces, respectively. Gene expression is driven by the constitutive promoter ermEp*, and the RBS and multiple cloning sites (MCSs) lie immediately downstream of the promoter, which simplifies molecular cloning. Vector pPWW50-Gen Poly resembles pUWL201PW in all these aspects, with a preinstalled N-terminal hexahistidine tag followed by MCSs. These vectors have been used in various Streptomyces hosts, such as S. lividans TK64, with good success. Other notable examples are pIJ6021 (7.8 kb) and pIJ4123 (9.2 kb), both inducible, high-copy-number expression vectors (Takano et al., 1995). The thiostrepton inducible strong promoter PtipA is used in these vectors, followed by RBS and MCSs. pIJ4123 is a derivative of pIJ6021 with a unique NdeI site positioned downstream from the hexahistidine sequence and thrombin cleavage site. Streptomyces hosts have been used to express and purify KS-CLF heterodimers of different type II PKSs, including those from act (Tang et al., 2003a), tcm (Tang et al., 2003a), oxy (Zhang et al., 2007), whiE (Tang, Y., unpublished data), sch (Lee et al., 2005), pms (Lee et al., 2005), and ent (Izumikawa et al., 2006) gene clusters. Typical culturing and initial purification procedures for Streptomyces/pRM5-derived systems are as follows: spore suspensions of the expression strain collected from a well-sporulated R2YE plate are used to inoculate 50 ml of R2YE liquid seed culture (supplemented with 50 mg/l thiostrepton) at 250 rpm and 30 for 2 to 3 days. This is then used to inoculate 4 500 ml of R2YE medium containing 50 mg/l thiostrepton (1:50 inoculation ratio). Mycelium from the stationary phase cultures (3 days) is collected by centrifugation and resuspended in 60 ml of disruption buffer (D-buffer: 250 mM sodium phosphate [pH 7.1], 0.3 mM sodium chloride, 2 mM DTT, 1 mM benzamidine, 2 mM EDTA, 3 mg/l leupeptin, 3 mg/l pepstatin, and 30% glycerol). Mycelium is then disrupted with a French press (sonication disruption can be used but the lysis efficiency is lower for Streptomyces), and insoluble cellular debris is removed by centrifugation (15,000 rpm, 1 h, 4 ). DNA is precipitated by adding 0.2% polyethyleneimine (PEI) and is removed by centrifugation (15000 rpm, 1 h, 4 . Ammonium sulfate precipitation can be used to precipitate KS-CLF, which can normally be found between the 30 and 50% cuts of (NH4)2SO4 (Tang et al., 2003a).

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If no purification tag is introduced into KS-CLF, multiple chromatography steps following ammonium sulfate precipitation provides an effective purification method. For example, the (NH4)2SO4 precipitated tcm KSCLF was re-dissolved in buffer E [100 mM NaH2PO4, 2 mM DTT, and 2 mM EDTA (pH 7.4)] and loaded onto a phenyl-sepharose column preequilibrated with buffer F (buffer E with 1.5 M (NH4)2SO4). The following gradient was then applied to the column: 100% buffer F from 0 to 30 min, 40% buffer F from 30 to 60 min, 10% buffer F from 60 to 200 min, and 0% buffer F from 200 to 240 min. The heterodimeric KS-CLF complex elutes near the end of the gradient (Tang et al., 2003a). Fractions containing tcm KS-CLF were pooled and buffer exchanged into buffer E and loaded onto a HiTrap-Q column pre-equilibrated with buffer E. Elution is performed with increasing concentration of NaCl in buffer E (0 to 400 mM ) and tcm KS-CLF elutes at NaCl concentration 200 mM. The KS-CLF obtained by this method is typically greater than 80% pure. An additional gel filtration step can further purify the enzyme. For hexahistidine-tagged KS-CLF, Ni-NTA and ion-exchange chromatography (if necessary) are applied, starting from re-suspending (NH4)2SO4 precipitated protein in buffer A, and following the protocols described earlier. Alternatively, the FLAG affinity tag has been introduced to the act CLF, which resulted in the recovery of high-purity KS-CLF that led to the crystallization of act KS-CLF (Keatinge-Clay et al., 2004). To purify FLAGtagged act KS-CLF, the phenyl-sepharose chromatography step was used as described previously. This is necessary to avoid fouling of the expensive Anti-FLAG resin. Fractions containing act KS-CLF are pooled and the buffer is exchanged into TBS buffer (50 mM Tris (pH 7.4), 0.15 M NaCl, and 10 mM CaCl2) and loaded onto a column packed with Anti-FLAG M1 agarose affinity gel (5 ml). The column is washed with 30 ml of TBS, and act KS-CLF is eluted with 3 5 ml of TBS containing 100 mg/ml FLAG peptide (Sigma). The eluent is then concentrated and the buffer is exchanged into buffer E containing 20% glycerol (Tang et al., 2003a). Although protein purification procedures are only discussed for KS-CLF using Streptomyces/ pRM5 derived systems here, other type II PKS components can be expressed and purified from Streptomyces using similar methods.

3. In Vitro Activity Assays 3.1. Minimal PKS activity assays 3.1.1. Product assays for minimal PKS The most direct method to assay the minimal PKS is to perform the product formation assay. The outcome of the assay will inform whether the purified enzyme components are active. Most type II minimal PKSs can utilize


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malonyl-CoA as both starter (decarboxylative priming) and extension units. Depending on the chain length specificity of the KS-CLF, a fixed set of products can be expected as result of spontaneous cyclization of the polyb-ketone backbone in the absence of tailoring enzymes. Examples of these products are octaketides SEK4 and SEK4b, decaketides SEK15 and SEK15b, and dodecaketides TW93c and TW93d (Fig. 16.4). Therefore, formation of these products can confirm the chain length specificity of the minimal PKS, or can be used to determine the product size of unknown minimal PKSs isolated from genome mining or metagenomic approaches. In addition, the product-based assays can test the overall catalytic efficiency of the minimal PKS complex. The reactions are typically performed in 100 mM NaH2PO4 with 1 to 20 mM KS-CLF, 20 to 50 mM holo-ACP, 2 mM malonyl-CoA, 0.1 to 1 mM MAT, 2 mM DTT, 10% glycerol. The use of DTT has been found to lead to undesired thioester reduction of acyl-S-ACP, and the less nucleophilic TCEP can be used to prevent oxidation of free thiols (Beltran-Alvarez et al., 2007). The reactions are incubated at 30 , and 100-ml reaction solution is extracted with 300 ml ethyl acetate (EA)/acetic acid (AcOH) (99:1) twice. The reactions can also be terminated by the addition of solid NaH2PO4 to saturation, or HCl to pH 3 and extracted with EA. The combined organic phases are evaporated to dryness, re-dissolved in 20 ml of DMSO and subjected to HPLC analysis. Alternatively, analytical scale solid-phase extraction columns (such as DSC-18, Supelco) can be used to isolate the compounds (Kallio et al., 2008). Reverse phase C18 column chromatography is used for aromatic polyketide analysis with mobile phases containing water and acetonitrile (supplemented with 0.1% trifluoroacetic acid [TFA]). In minimal PKS product assays, the substrate malonyl-CoA can also be generated in situ using the Rhizobium trifolii malonyl-CoA synthetase, MatB (An and Kim, 1998). MatB synthesizes malonyl-CoA from malonic acid and CoA in an ATP-dependent fashion. MatB can be expressed and purified from E. coli at very high levels (>30 mg/l). When MatB is used, the following reagents are added to the assay mixture in place of malonylCoA: 100 mM sodium malonate, 5 mM MgCl2, 20 mM ATP, 5 mM CoA, and 20 mM MatB. The MatB system has several important advantages over malonyl-CoA. (1) For large-scale in vitro production and characterization of polyketides, malonyl-CoA can become prohibitively expensive. Using MatB circumvents this obstacle and the much more economical free CoA is used. (2) Using MatB, different 13C-labeled malonic acid can be used as extender units to produce isotopically-labeled polyketide backbones. The most important application of 13C labeling is to confirm whether a particular product is derived from malonyl-CoA (hence is a polyketide) using LC/MS analysis. For example, when 12C-malonic acid is replaced by 2-13C-malonic acid, and 2-13C-malonyl-CoA is synthesized by MatB and is incorporated into the polyketide backbone, leading to a mass increase of

O

O

HO 8 X malonyl-CoA octaketide minimal PKS O

O O

O O

O

O

O 9

O 13

O S

Enz

CoA

10 X malonyl-CoA decaketide minimal PKS

O

11 O

S

O 9

O 5

7

15

12 X malonyl-CoA dodecaketide minimal PKS O

O

O 17

O

O

O 1 S

3

O

O

O

O

O

O

19 Enz

O

O

O

O 9

S

Enz

OH OH

OH 15 12 HO

10

HO OH 12

OH

O HO

7

O O

O HO SEK4

OH

O

O

HO

O

OH

OH O

14 O

HO

OH O 12

12

9

7

HO O

7

O

O

OH

HO O

O

7

HO

OH HO O

O

HO SEK4b

O

HO

O HO SEK15

O

SEK15b

O

HO

O TW93c

HO

Figure 16.4 Examples of polyketides produced in type II minimal PKS product^based assays.

O TW93d

O


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þ1 for each ketide unit. A mass increase of 8 and 10 should be observed for octaketides and decaketides, respectively. By comparing the mass of the product with a 12C or 13C precursor, the size of the polyketide products can be readily established. (3) Using MatB and triply 13C labeled malonic acid can produce polyketides that are isotopically labeled at each carbon, which will significantly enhance the 13C NMR signals and simplify the identification of products. The product-based assay has been used to investigate the polyketide chain length control mechanism (Tang et al., 2003b). Using homology structure–directed mutagenesis, key residues in the CLF were identified and manipulated to alter polyketide chain length. Mutant act KS/CLFs were purified from S. coelicolor and the distribution of octaketides and decaketides produced in vitro using the minimal PKS assay was analyzed. It was found that the point mutation F116A was sufficient to increase the levels of decaketides from 0% to over 60% of total polyketide products, and the F109A/F116A double mutant completely converted the act octaketide synthase into a decaketide synthase. Another application of product assays is to probe the starter unit selection and incorporation in type II polyketide synthesis (see Section 3.2). The HPLC or LC/MS analysis can be quantitative, but large amounts of proteins and substrates are needed, making this method impractical for kinetic assays. Instead, a more sensitive radioactive assay using 14C-labeled malonyl-CoA can be performed, which can be used to detect as little as 25 pmol of polyketide product (Dreier et al., 1999). For kinetic parameter determination, malonyl-CoA is usually added to 100 to 2000 mM (specific activity between 0.55 and 55 nCi/nmol). The minimal PKS components can be combined at different concentrations depending on the assay goal. Typically, KS-CLF can be added to a final concentration of 0.05 to 5 mM, holo-ACP to 0.1 to 200 mM, and MAT 1 to 1000 nM. The reactions are initiated by the addition of 2-14C-malonyl-CoA and the products are extracted in the same way as described above. Products are re-dissolved in 15 ml EA, separated by thin-layer chromatography (TLC) (EA/AcOH, 99:1) and quantified with a phosphorimager. A typical apparent Km of type II ACP is in the 1- to 7-mM range (Tang et al., 2003a), the Km of malonyl-CoA is in the 70- to 220-mM range (Dreier et al., 1999), and the apparent turnover kcat is between 0.1 to 0.5 min–1 (Dreier et al., 1999; Lee et al., 2005; Tang et al., 2003a). Recently, a continuous spectrophotometric method that measures the release of free CoA has been used to study the kinetics of act minimal PKS (Beltran-Alvarez et al., 2007). The free CoA generated as a result of malonyl-CoA consumption is used as a substrate by a-ketoglutarate dehydrogenase (KDH) to produce succinyl-CoA with concomitant reduction of NADþ to NADH, which can be followed spectrophotometrically at 340 nm. The final concentrations of the components in KDH assays were reported as follows: 100 mM phosphate

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buffer (pH 7.3), 1 mM EDTA, 1 mM tris(2-carboxyethyl)phosphine (TCEP), 2 mM a-ketoglutaric acid, 0.5 mM NADþ, 0.4 mM thiamine pyrophosphate (TPP), 1 mM MgCl2 and 80 mU KDH. The spectrophotometric assay is highly convenient and numerous assays can be performed in parallel. Since the assay measures release of CoA, this technique can be used to monitor the kinetics of any minimal PKS. 3.1.2. MAT-ACP labeling assay The malonyl-CoA:ACP transacylation assay catalyzed by MAT can be used to determine the rate of ACP malonylation in the minimal PKS. The data is important towards establishing the rate-limiting step of the product-based assay described above (Beltran-Alvarez et al., 2007). Typically, the kcat range of S. coelicolor MAT toward type II ACPs is 3000 to 9000 min–1, which indicates that the formation of malonyl-ACP is not the ratelimiting step in type II polyketide synthesis. However, when the ACP is mutated or engineered, the MAT:ACP interaction may be less efficient. It is notable here that at least act ACP has been shown to possess an intrinsic self-malonylation activity, albeit only at elevated concentrations of malonyl-CoA (Arthur et al., 2005). The conversion of holo-ACP to malonyl-ACP can be assayed using 14C-labeled malonyl-CoA (Summers et al., 1995). A typical reaction mixture contains 200 mM 2-14C-malonyl-CoA (55 mCi/mmol), 100 mM holoACP, 1 to 10 nM MAT, 1 mM DTT in 100 mM NaH2PO4 buffer (pH 7.0). Reactions are performed at 25 for short time intervals (usually less than 2 min). Aliquots (10 ml) are quenched by adding SDS-PAGE loading buffer and applied to a 4 to 20% SDS gel to separate ACPs from MAT and malonyl-CoA. The gel is subsequently dried and autoradiographed. Alternatively, a trichloroacetic acid (TCA) precipitation assay can be performed to precipitate malonyl-ACP, followed by scintillation counting. The protein gel autoradiography technique can also be used to detect covalent intermediates and malonyl transfer among minimal PKS components by incubating 14C-labeled malonyl-CoA with one or several of minimal PKS components (Carreras and Khosla, 1998). 3.1.3. KS-ACP protein–protein interaction assays The successful recognition between KS and ACP is essential for product turnover in a minimal PKS. The transient and noncovalent nature of these interactions hinders the direct observation of their compatibility, leaving the product-based assay the only investigation method. Recently, a new method was developed to trap the ACP during its interaction with KS, providing direct insight into the protein–protein interactions between ACP and KS (Worthington et al., 2008). Chemo-enzymatically synthesized CoA analogues and Sfp were used to produce crypto-ACPs, in which a nonhydrolyzable electrophilic probe was tethered in place of the thiol terminus of


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holo-ACP. A covalent crosslink is formed between KS and crypto-ACPs in the event of fruitful protein interactions. The complex can be detected by mobility shift during SDS-PAGE and can be confirmed by MALDI MS/MS analyses. 3.1.4. High-resolution mass spectrometry The application of electrospray ionization Fourier-transform mass spectrometry (ESI-FTMS) in the study of modular PKSs and NRPSs has been reviewed recently (Dorrestein and Kelleher, 2006). ESI-FTMS has the advantage of high resolution and mass accuracy compared to traditional mass spectrometry, and can be used to investigate substrate tolerance, timing of intermediate transfer and tailoring reactions on polypeptides of greater than 70 KDa. ESI-FTMS analysis has been employed in type II PKS study to observe the phosphopantetheinyl functionality of ACPs. Undoubtedly, ESI-FTMS will be a powerful tool in dissecting the individual steps of the minimal PKS, and provide significantly higher resolution snapshots than the aforementioned polyketide product-based assay and autoradiography. It is notable that a nonhydrolyzable malonyl-CoA analogue has been demonstrated to trap polyketide intermediates in iterative condensation, which may facilitate the detection of various protein-bound intermediates in type II PKS assembly by MS analysis (Spiteller et al., 2005).

3.2. Starter unit synthesis and incorporation assays The starter unit represents an attractive position to introduce structural variations into type II polyketides, especially considering that malonylCoA is the only extender unit accepted by all known type II PKSs. Most type II PKSs are primed by an acetate unit with acetyl-ACP or through the decarboxylation of a malonyl-ACP. Numerous medicinally important polyketides are initiated with nonacetate starter units (Fig. 16.1), including daunorubicin (propionate) (Bao et al., 1999), tetracycline (malonamate) (Zhang et al., 2006), frenolicin (butyrate) (Bibb et al., 1994), R1128 (medium-length alkylacyl groups) (Tang et al., 2004b), enterocin (benzoate) (Cheng et al., 2007), and hedamycin (a reduced polyketide starter unit) (Bililign et al., 2004). Genetic characterizations of the corresponding gene clusters has revealed the presence of various enzymes that can synthesize the starter unit, bypass the acetate priming, and transfer the starter unit to the minimal PKS. In vitro analyses of selected nonacetate-primed PKSs have revealed the enzymatic basis of these strategies. While biogenesis of starter units can be assayed independently, priming and incorporation of the starter units are frequently probed using the product-based assays in the presence of the minimal PKS.

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3.2.1. Direct priming of minimal PKS The starter unit specificity of the minimal PKS can be directly probed by adding acyl-CoA primers to the product assay described above. The typical reaction can consist of 1 mM KS-CLF, 200 nM MAT, 2 mM malonyl-CoA, 30 mM holo-ACP, and 2 mM of the desired acyl-CoA. The reaction can then be extracted with organic solvents and analyzed by either radio-TLC (if 2-14C malonyl-CoA is used) or LC-MS. An important finding of this assay is that many normally acetyl-primed minimal PKSs can instead be primed with alternative starter units. For example, the act KS-CLF was shown to be primed by either alkyl starter units such as C4 to C8 fatty acylCoAs (Nicholson et al., 2003), or aryl starter units such as benzyol-ACP (Izumikawa et al., 2006). Similarly, the tcm KS-CLF can be primed by C3 to C10 fatty acyl-CoAs to afford polyketides containing nonacetate starter units (Tang et al., 2004a). These results can also be extended to dodecaketide synthases such as spore-pigment sch and pradimicin ( pms) PKSs (Tang, Y., unpublished data). In each case, the resulting nonacetate primed products contained the same number of carbons in the polyketide backbone. The increases in sizes of the starter units led the corresponding decreases of the number of ketide units in the final polyketides, suggesting that the KS-CLF can maintain chain length through a ‘‘ruler’’ mechanism (Nicholson et al., 2003). The efficiency of starter unit incorporation varies between different minimal PKSs and the starter units. The efficiency of incorporating nonacetate primers can be improved by adding an acetylACP thioesterase such as ZhuC from the R1128 pathway (see Section 3.2.2) (Tang et al., 2004a). Recently fluoroacetate has been incorporated into an octaketide backbone in vitro (Hong et al., 2008). Fluoroacetyl-ACP was synthesized using fluoroacetyl-CoA, Sfp, and apo-ACP, and added to 30 mM act KS-CLF. The reaction was then preincubated at 30 for 10 min to facilitate loading of the fluoroacetyl starter unit onto the KS active site. This was followed by the addition of MAT and malonyl-CoA for chain elongation. A fluorinated variant of SEK4b was produced in addition to SEK4 and SEK4b, as determined by LC-MS and NMR analysis. 3.2.2. Starter unit biosynthesis The frenolicin, R1128 and daunorubicin PKSs each contains a b-ketoacyl: ACP synthase III (KSIII) enzyme that is required for the incorporation of the alkyl starter units. DpsC, the KSIII for daunorubicin, is found to specify the selection of propionyl-CoA as the starter unit during daunorubicin biosynthesis (Bao et al., 1999). Native DpsC was purified from S. lividans by multiple purification steps and was shown to have high substrate specificity toward propionyl-CoA. DpsC exhibited both KSIII and acyltransferase activities toward both DpsG (the ACP encoded in the daunorubicin


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biosynthetic gene cluster) and E. coli ACPp (the ACP of FAS). ES-MS/ HPLC was used to confirm formation of the adduct propionyl-DpsC, and trypsin digestion followed by ES-MS/HPLC analysis revealed that Ser118 is the active site of DpsC. ZhuH and FrenI are the gatekeeping KSIII in the R1128 and fren gene clusters, respectively. Both KSIIIs catalyze the condensation of a short-chain acyl-CoA with malonyl-ACP to yield a b-keto-alkylacyl-ACP species. Reduction of the b-keto-alkylacyl-ACP by the KR/DH/ER cascade associated with the endogenous FAS yields a saturated alkylacyl-ACP that is used to prime the minimal PKS (Tang et al., 2006). The acyl-CoA substrate specificity of ZhuH has been studied thoroughly in vitro using radiolabeled acyl-CoA precursors. Formation of radiolabeled b-keto-alkylacyl-ACP products were detected and quantified by using a TCA protein precipitation assay (Meadows and Khosla, 2001). Propionyl-CoA was shown to be the best substrate of ZhuH, and is the precursor to the compound R1128b. FrenI was found to be highly specific for acetyl-CoA, which led to the formation of the butyryl starter group present in frenolicin. Both the R1128 and frenolicin PKSs contain two ACPs. Upon initial gene cluster identification, the roles of the two ACPs were thought to be interchangeable. After expression and purification of all four ACPs from E. coli, the ACPs were assayed for their roles in the synthesis of the aromatic polyketides in vitro (Tang et al., 2003a). KS-CLFs from several pathways were mixed with each ACP and were assayed for the formation of polyketides using 2-14C-malony-CoA. Similarly, the KSIII enzymes from the two clusters were also assayed in the presence of these ACPs. Interestingly, one ACP from each gene cluster was found to be a dedicated priming ACPp (ZhuG/ FrenJ), while the other ACP was part of the minimal PKS (ZhuN/ FrenN). The functions of the two ACPs from each gene cluster cannot be interchanged as the KS-CLF and KSIII each displays high selectivity towards its cognate ACP. Surprisingly, a single residue was largely responsible for this orthogonality, as mutation of this residue in either ZhuG or ZhuN resulted in recognition of the mutant ACP by both KS-CLF and KSIII. A third member that is critical for the incorporation of the starter unit in R1128 is ZhuC, which was first identified to be a dedicated MAT based on primary sequence similarity to acyltransferases. However, in vivo reconstitution experiments showed that ZhuC was indispensable for the incorporation of the alkylacyl-ZhuG starter units generated by ZhuH. In vitro analysis of ZhuC confirmed its role as an acetyl-ACP thioesterase that attenuates the acetyl priming pathway in favor of nonacetate priming. When ZhuC was added to acetyl-ACP, which can be generated from apo-ACP, acetyl-CoA and Sfp as described in section 2.2, rapid hydrolysis of the acetyl group was observed and holo-ACP with a free thiol was recovered. The hydrolysis assay was performed using either TCA precipitation or SDS-PAGE gel

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autoradiography (Tang et al., 2004a). A typical reaction consists of radiolabeled acetyl-ACP, ZhuC, 2 mM DTT, and 10% glycerol in 100 mM NaH2PO4 buffer. The role of ZhuC was further confirmed by using the product-based assay described previously. When alkylacyl-ZhuG was added to a minimal PKS without ZhuC, acetyl-primed products were the dominant species produced by the KS-CLF. However, when ZhuC was added, the levels of acetyl-primed compounds were significantly attenuated, with a corresponding increase in the amount of alkylacyl-primed polyketides. Benzoyl-priming during enterocin biosynthesis requires the biosynthesis of the starter unit benzoyl-ACP. A benzoate-CoA ligase EncN can synthesize benzoyl-ACP using two different mechanisms (Izumikawa et al., 2006). The first route is to transfer the benzoyl group from benzoyl-CoA to benzoyl-ACP, which can subsequently prime the ent KS-CLF. The benzoyl-transferase activity was measured using 7-14C-benzoyl-CoA and holo-ACP in a TCA precipitation assay. ATP (or AMP/PPi) was found to be an essential cofactor and was proposed to be an allosteric modulator of EncN. The second route is the direct ligation of benzoate to ACP, catalyzed by EncN. The assay was performed in 100 mM Tris-HCl (pH 7.5) buffer with 5.0 mM ATP, 5.0 mM MgCl2, 5 to 50 mM holo-ACP, 20 to 100 mM benzoic acid, and 2 to 5 mM of EncN. Benzoyl-ACP was detected by MALDI-TOF MS analysis of TCA-precipitated ACP. Various holoACPs such as ent, act, and tcm ACPs can be modified with the benzoyl group by EncN.

3.3. Assays of tailoring enzymes responsible for aromatic polyketide scaffold formation: KR, ARO, and CYC Enzymes that act on the nascent polyketide backbone, such as KR, ARO and CYC, are commonly referred to as immediate tailoring enzymes. These enzymes regioselectively modify the reactive poly-b-keto backbone synthesized by the minimal PKS to generate the aromatic scaffold. The fate of the nascent type II PKSs can be divided into two paths, reduced and unreduced, dependent on the presence of C-9 KR (reduction at the ninth carbon from the carboxy terminus of the assembled polyketide). In the reduced path, the first ring is usually cyclized through C-7/C-12 connectivity. There is increasing evidence to suggest that the C-9 KR may be responsible for fixing this regioselectivity (Korman et al., 2008; Ma et al., 2008). The cyclized first ring can then be dehydrated/aromatized by a dedicated ARO. Examples of C-9 reduced aromatic polyketide include tetracycline and anthracyclines. In the unreduced path, the first ring can be spontaneously cyclized at C-7/C-12 or catalyzed by dedicated CYCs. Mithramycin is synthesized from this path. The unreduced polyketide backbone can also be cyclized via C-9/C-14 connectivity by dedicated CYCs to afford the D ring found in tetracenomycin and related compounds.


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In vitro assays with the immediate tailoring enzymes are typically performed in the presence of the minimal PKS, which serves to provide the otherwise inaccessible poly-b-ketone substrates in situ (Zawada and Khosla, 1999; Zhang et al., 2007). The KR or CYCs are added to final concentrations of two- to five-fold excess to that of KS-CLF, which are generally sufficient to completely reduce or cyclize the substrates, respectively. Through both in vivo genetic analysis and in vitro assays, it is evident that the C-9 KRs, and the first- and second-ring CYCs with the same regioselectivity in either path can be interchanged. The functional equivalence of these enzymes is highly useful during in vitro assays. An insoluble cyclase from the pathway of interest can be substituted with equivalent enzymes that can be solubly expressed in E. coli. We demonstrated this versatility in the reconstitution of oxytetracycline cyclization steps. To produce a tricyclic intermediate, we used the oxy minimal PKS, act C-9 KR, gris ARO, and oxy second ring CYC (Zhang et al., 2007). The native oxy C-9 KR and ARO were not solubly expressed in E. coli. Although stable protein–protein interactions are not detected between minimal PKS and immediate tailoring enzymes, kinetic assays reveal that the addition of some of these tailoring enzymes can dramatically increase the turnover rate of a minimal PKS (Zawada and Khosla, 1999). One possible explanation for the increase turnover rate is that enzymatic cyclization of the polyketide backbone increases the rate of product release from the ACP. Synthetic substrate mimics have been used to probe the activity of KRs. Although this can only provide limited insight into the natural functions of the KR, this method has been used to probe the substrate specificity and cofactor requirements of KRs. A typical kinetic assay for KR is performed in 100 mM NaH2PO4 (pH 7.4) containing 100 mM NADPH, 40 mM substrate analogue, and 35 nM enzyme. Two-percent DMSO is used in the reaction solution to increase the solubility of the synthetic substrates when necessary. The decrease of A340 from the conversion of NADPH to NADPþ is monitored by UV-vis spectrophotometer. Reaction velocities are calculated on the basis of the A340 molar absorption coefficient of NADPH (6220 M–1 cm–1). The act C-9 KR has been assayed using a series of synthetic substrate candidates, among which bicyclic compounds such as trans-1-decalone, 2-decalone and a-tetralone can be stereoselectively reduced (Korman et al., 2008). Synthetic analogues were also used as substrates in the enzymatic assay for RED1, a second stereospecific KR encoded in the act pathway (Itoh et al., 2007). RED1 is assigned to the reduction of a b-ketone group in an unstable bicyclic intermediate. Interestingly, RED1 prefers 3-oxo-4-naphthylburyric acid (ONBA) over the N-acetylcysteamine thioester or methyl ester versions of ONBA. The in vitro result strongly suggests that the in vivo substrate of RED1 is a free acid instead of an acyl thioester attached to act ACP. Four terminal-ring cyclases have been characterized individually in vitro due to the availability of stable substrates. These assays can directly test the

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substrate specificity and optimal reaction conditions of cyclases in the absence of any influence from a minimal PKS complex. DnrD and AknH have been shown to convert aklanonic acid methyl ester (AAME) to aklaviketone (AK) in the daunorubicin and aclacinomycin biosynthetic pathways, respectively (Kallio et al., 2006; Kendrew et al., 1999). Analogously, SnoaL was confirmed to catalyze nogalaviketone formation from nogalonic acid methyl ester (NAME) during nogalamycin biosynthesis (Sultana et al., 2004). In addition, the Tcm F2 cyclase TcmI, which cyclizes the last ring during tetracenomycin biosynthesis, was also characterized in vitro (Thompson et al., 2004). Cyclase assays were typically performed in 100 mM Tris-HCl buffer (pH 7.5) at 30 and the polyketide products are extracted by standard methods and analyzed by TLC or HPLC. Strong pH dependence was observed for these cyclases. Although the four enzymes are all fourth ring cyclases catalyzing intramolecular aldol condensation during decaketide formation, they have different substrate requirements and product outcomes. In contrast to TcmI, DnrD, AknH, and SnoaL recognize methyl esters of anthraquinone carboxylic acids, and no dehydration of the last ring occurs after cyclization. SonaL catalyzes the cyclization and affords nogalaviketone with an S-configuration, in contrast to the products of DnrD and AknH which have R-configurations. Since the stereoisomers of aklaviketone cannot be easily separated using conventional RP HPLC, the stereochemistry of products was determined using an aklavinone-10hydroxylase (RdmE)-coupled assay system, in which RdmE can only hydroxylate the species in the R-configuration (Kallio et al., 2006; Niemi et al., 1999).

3.4. Assays of downstream tailoring enzymes Polyketide downstream tailoring enzymes include methyltransferase, oxygenase, glycosyltransferase, acyltransferase, halogenase, aminotransferase, and so on, most of which are important for product structural variations and bioactivities. These enzymes are not limited to type II PKSs, and have also been found and studied in other polyketide and nonribosomal peptide biosynthetic pathways. Due to space limitations, only assays of methyltransferase, oxygenase and glycosyltransferase will be discussed in this section. 3.4.1. Methyltransferase Most methyltransferases use the cofactor S-adenosyl-L-methionine (SAM) and transfer its activated methyl moiety to substrates. A typical reaction mixture contains 100 mM KH2PO4 (pH 7.5), 80 mM SAM, 40 mM substrate, and 2 mM enzymes. Less than 10% DMSO, MeOH or other organic solvents are added to aqueous solutions to increase substrate solubility when needed. The presence of organic solvents in reaction solutions are common for assays involving type II PKS tailoring enzymes, since most of the


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substrates are polycyclic compounds that have only limited solubility in water. The reaction is incubated for 1 h at 37 , and the product is extracted and analyzed by HPLC. Radioactive SAM can be used to label the product and quantify the reaction by scintillation counting. At least five methyltransferases involved in aromatic polyketide biosynthesis have been characterized in vitro, including those that transfer a methyl group from SAM to oxygen, nitrogen, or carbon atom. TcmN and DnrK are O-methyltransferases that install the methyl groups on the D-ring phenols in tetracenomycin and daunorubicin, respectively ( Jansson et al., 2004; Shen and Hutchinson, 1996) (Fig. 16.1). During mithramycin biosynthesis, an O-methyltransferase and a C-methyltransferase have been identified and characterized (Lozano et al., 2000). Recently, the transformation of 4-amino-ATC to ATC has been reconstituted in vitro with OxyT, which is an N,N-dimethyltransfrease catalyzing the reaction in a stepwise manner (Zhang et al., 2008). 3.4.2. Oxygenases There are several types of oxygenases involved in modifying the aromatic polyketide scaffold, including anthrone oxygenases, flavin-dependent oxygenases, and cytochrome P-450 monooxygenases. They differ extensively in their substrate specificity, cofactor requirement and mode of action, and their activities are accordingly assayed by different methods. Example of anthrone oxygenases are TcmH, ActVA-ORF6 and AknX (Chung et al., 2002; Kendrew et al., 1997; Shen and Hutchinson, 1993). No apparent prosthetic group is found for oxygen activation, and no specific cofactor is needed for the activity of these oxygenases. In addition, the enzymes do not seem to be sensitive to pH (6.5 to 9.5) or temperature. Anthrone oxygenases usually have relaxed substrate specificity; for example, tetracenomycin F1 is recognized by ActVA-ORF6 and the substrate analog emodinanthrone has been used for AknX in vitro assay. The anthrone compounds have absorbance in the range of 400 to 500 nm which will be shifted after quinone formation; therefore, the reaction can be monitored spectrophotometrically according to the molecular extinction coefficient differences between the substrate and product at certain wavelength. Recently an interesting oxygenase, JadH from the jadomycin biosynthetic pathway, has been characterized (Chen et al., 2005). By testing different intermediates of the jad pathway as substrates, JadH was revealed to have both anthrone oxygenase and dehydrase/aromatase activities and relaxed substrate specificity. Flavin-dependent oxygenases catalyze versatile reactions including hydroxylations, epoxidations, Baeyer-Villiger and Favorskii rearrangements. The presence and identity of the prosthetic group can be determined as follows (Shen and Hutchinson, 1994): pure protein is boiled for 5 min, cooled on ice immediately, and centrifuged to pellet the denatured protein.

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The supernatant is then subjected to HPLC analysis to identify and quantify the flavin prosthetic group released by this process through comparison to FAD and FMN standards. The concentrations of FAD and FMN are determined according to their A450 molar absorption coefficient 11.3 mM–1 cm–1 and 12.2 mM–1 cm–1, respectively. Yellow holo-enzyme can be partially reconstituted from colorless apo-enzyme in vitro by mixing 1 mole of apo-enzyme, 10 mole of either FAD or FMN (dependent on native prosthetic group), 25 mM Tris-HCl (pH 8.0), 1 mM DTT, and incubating on ice for 1.5 h. NADPH is needed as a cofactor for almost all flavin-dependent oxygenases, while the optimal pH is different for each enzyme. Following the assay, the oxidized products can be analyzed by TLC or HPLC, and the reaction kinetics can be usually determined by monitoring NADPH or substrate consumption. 18O isotope incorporation reactions followed by MS analysis can be used to establish the origins of oxygen atom(s) in final products. 18O2 gas reactions have been carried out in a Thunberg cuvette, in which oxygen is completely removed with a combined vacuum and repeated nitrogen rinses before mixing the reaction components. 18O-H2O reactions are performed in the reaction buffer where H2O is replaced by 18O-H2O (Kallio et al., 2008). Examples of flavin-dependent oxygenases characterized thus far include the tri-hydroxylase TcmG/ElmG in tetracenomycin biosynthesis (Rafanan et al., 2000; Shen and Hutchinson, 1994), di-hydroxylase OxyL in tetracycline biosynthesis (Zhang et al., 2008), and PgaE and PgaM involved in gaudimycin C biosynthesis (Kallio et al., 2008). Reconstitution of EncM activity in vitro confirmed that EncM is responsible for catalyzing the Favorskii-like oxidative rearrangement during enterocin biosynthesis (Cheng et al., 2007). In addition, MtmOIV from the mithramycin biosynthetic pathway has been confirmed to adopt the Baeyer-Villiger mechanism by the isolation of the lactone intermediate premithramycin B-lactone in the in vitro reaction (Gibson et al., 2005). Very few cytochrome P-450 monooxygenases from type II PKSs have been characterized in vitro. Examples include EncR and DoxA, which have been purified from E. coli TB1 and S. lividans TK24, respectively (Piel et al., 2000; Walczak et al., 1999). An extinction coefficient of 91 M–1 cm–1 is employed to quantify the cytochrome P-450 heme using the absorbance difference between 450 and 490 nm. P-450 enzymes require electron transport from NADPH via ferredoxin and ferredoxin:NADPþ oxidoreductase. Instead of the natural P450:reductase pairs, commercially available spinach ferredoxin and spinach ferredoxin:NADPþ reductase (Sigma) are commonly used in assays. A relatively high concentration of NADPH is maintained to ensure electron shuttling efficiency, which can be realized by using NADPH together with the NADPH regenerating system (NADPþ, glucose-6-phosphate, and glucose-6-phosphate dehydrogenase) (Walczak et al., 1999). In addition, inhibition assays on P-450 are usually carried out using known inhibitors such as 4-methylpyrazole, quinidine,


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troleandomycin, sulfaphenazole, and ancymidol (Piel et al., 2000; Walczak et al., 1999). 3.4.3. Glycosyltransferases Although genes encoding glycosyltransferases are widely spread among type II polyketide gene clusters, very few of them have been characterized in vitro due to the difficulties associated with obtaining active enzymes, the aromatic aglycones or the NDP-deoxysugars. Recent examples are AknK and AknS, which are involved in the biosynthesis of aclacinomycin (Leimkuhler et al., 2007; Lu et al., 2004, 2005). The aklavinone aglycone was prepared through the degradation of commercially available aclacinomycin A, and the TDP sugars were synthesized chemically. A typical reaction buffer includes 75 mM Tris-HCl (pH 7.5) and 10 mM MgCl2. It is worth noting that AknS needs an activating protein partner, AknT, to be fully functional; AknT accelerates AknS turnover rate by 200-fold without affecting its Km. Both in vitro and in vivo analysis have revealed that many glycosyltransferases have relaxed substrate specificities toward sugar donors and aglycons (Hertweck et al., 2007), indicating the biocatalysis potential of this group of enzymes in generating novel glycosylated polyketides with improved pharmaceutical properties. 3.4.4. Enzymatic total synthesis Recently, in vitro enzymatic total synthesis has been demonstrated to be a powerful method of producing complex aromatic polyketides (Cheng et al., 2007). The aromatic polyketide enterocin was synthesized in a single reaction vessel from simple benzoate and malonate substrates (Cheng et al., 2007). The recombinant proteins EncA-B (KS-CLF), holo-EncC (ACP), FadD (MAT), EncN (benzoyl-ACP ligase), EncD (C-9 KR), EncM (flavin-dependent oxygenase), EncK (SAM-dependent methyltransferase), EncR (cytochrome P-450 monooxygenase), and MatB (malonyl-CoA synthetase) were mixed with substrates and necessary cofactors (ATP, Mg2þ, NADPH, SAM, ferredoxin, ferredoxin-NADPþ reductase) to afford enterocin in 25% overall yield. This method offers opportunities to produce and engineer aromatic polyketides without fermentation and genetic engineering of Streptomyces, and may lead to the in vitro enzymatic combinatorial synthesis of natural products.

ACKNOWLEDGMENTS Research in our laboratory on this topic has been supported by grants from the National Science Foundation (CBET #0545860) and Universitywide AIDS Research Program of the University of California (D06-LA-193). Yi Tang thanks Chaitan Khosla for introducing him to the genetics and biochemistry of type II PKSs.

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Shen, B., and Hutchinson, C. R. (1996). Deciphering the mechanism for the assembly of aromatic polyketides by a bacterial polyketide synthase. Proc. Natl. Acad. Sci. USA 93, 6600–6604. Spiteller, D., Waterman, C. L., and Spencer, J. B. (2005). A method for trapping intermediates of polyketide biosynthesis with a nonhydrolyzable malonyl-coenzyme A analogue. Angew. Chem. Int. Ed. 44, 7079–7082. Sultana, A., Kallio, P., Jansson, A., Wang, J. S., Niemi, J., Mantsala, P., and Schneider, G. (2004). Structure of the polyketide cyclase SnoaL reveals a novel mechanism for enzymatic aldol condensation. EMBO J. 23, 1911–1921. Summers, R. G., Ali, A., Shen, B., Wessel, W. A., and Hutchinson, C. R. (1995). Malonylcoenzyme A:acyl carrier protein acyltransferase of Streptomyces glaucescens: A possible link between fatty acid and polyketide biosynthesis. Biochemistry 34, 9389–9402. Takano, E., White, J., Thompson, C. J., and Bibb, M. J. (1995). Construction of thiostrepton-inducible, high–copy-number expression vectors for use in Streptomyces spp. Gene 166, 133–137. Tang, Y., Koppisch, A. T., and Khosla, C. (2004a). The acyltransferase homologue from the initiation module of the R1128 polyketide synthase is an acyl-ACP thioesterase that edits acetyl primer units. Biochemistry 43, 9546–9555. Tang, Y., Lee, T. S., and Khosla, C. (2004b). Engineered biosynthesis of regioselectively modified aromatic polyketides using bimodular polyketide synthases. PLoS Biol. 2, 227–238. Tang, Y., Lee, H. Y., Tang, Y., Kim, C. Y., Mathews, I., and Khosla, C. (2006). Structural and functional studies on SCO1815: A beta-ketoacyl-acyl carrier protein reductase from Streptomyces coelicolor A3(2). Biochemistry 45, 14085–14093. Tang, Y., Lee, T. S., Kobayashi, S., and Khosla, C. (2003a). Ketosynthases in the initiation and elongation modules of aromatic polyketide synthases have orthogonal acyl carrier protein specificity. Biochemistry 42, 6588–6595. Tang, Y., Tsai, S. C., and Khosla, C. (2003b). Polyketide chain length control by chain length factor. J. Am. Chem. Soc. 125, 12708–12709. Thompson, T. B., Katayama, K., Watanabe, K., Hutchinson, C. R., and Rayment, I. (2004). Structural and functional analysis of tetracenomycin F2 cyclase from Streptomyces glaucescens. A type II polyketide cyclase. J. Biol. Chem. 279, 37956–37963. Walczak, R. J., Dickens, M. L., Priestley, N. D., and Strohl, W. R. (1999). Purification, properties, and characterization of recombinant Streptomyces sp. strain C5 DoxA, a cytochrome P-450 catalyzing multiple steps in doxorubicin biosynthesis. J. Bacteriol. 181, 298–304. Worthington, A. S., Hur, G. H., Meier, J. L., Cheng, Q., Moore, B. S., and Burkart, M. D. (2008). Probing the compatibility of type II ketosynthase–carrier protein partners. Chem. Bio. Chem. 9, 2096–2103. Zawada, R. J., and Khosla, C. (1999). Heterologous expression, purification, reconstitution and kinetic analysis of an extended type II polyketide synthase. Chem. Biol. 6, 607–615. Zhang, W., Ames, B. D., Tsai, S. C., and Tang, Y. (2006). Engineered biosynthesis of a novel amidated polyketide, using the malonamyl-specific initiation module from the oxytetracycline polyketide synthase. Appl. Environ. Microbiol. 72, 2573–2580. Zhang, W., Watanabe, K., Cai, X., Jung, M. E., Tang, Y., and Zhan, J. (2008). Identifying the minimal enzymes required for anhydrotetracycline biosynthesis. J. Am. Chem. Soc. 130, 6068–6069. Zhang, W., Watanabe, K., Wang, C. C., and Tang, Y. (2007). Investigation of early tailoring reactions in the oxytetracycline biosynthetic pathway. J. Biol. Chem. 282, 25717–25725. Ziermann, R., and Betlach, M. C. (1999). Recombinant polyketide synthesis in Streptomyces: Engineering of improved host strains. Biotechniques 26, 106–110.

C H A P T E R

S E V E N T E E N

Bacterial Fatty Acid Synthesis and its Relationships with Polyketide Synthetic Pathways John E. Cronan*,† and Jacob Thomas* Contents 1. Introduction 2. Bacterial Fatty Acids 3. Acyl Carrier Protein, the Key Component of Bacterial Fatty Acid Synthesis 4. Overview of the Reactions of Fatty Acid Biosynthesis 5. The Initiation Steps of Fatty Acid Synthesis 6. The Enzymes of the Fatty Acid Elongation Cycle 6.1. The 3-ketoacyl-ACP synthase reaction (FabB, FabF, FabH) 6.2. 3-ketoacyl-ACP reductase (FabG) 6.3. The 3-hydroxyacyl-ACP dehydratase (FabZ) 6.4. The enoyl-ACP reductase (FabI) 7. Unsaturated Fatty Acid Synthesis in E. coli 7.1. The 3-hydroxydecanoyl-ACP dehydratase (FabA) 8. The Abundant Exceptions to the E. coli Fatty Acid Synthesis Paradigm 8.1. Branched-chain fatty acids 8.2. Anaerobic synthesis of unsaturated fatty acids 8.3. Diversity of enoyl-ACP reductases 8.4. Type I megasynthase fatty acid synthesis 9. Relationships between Fatty Acid Synthesis and Polyketide Synthesis 10. Methods for Study of Bacterial Fatty Acid Synthesis 10.1. Preparation of the holo and apo forms of E. coli ACP 10.2. Strains and plasmids used 10.3. Holo-ACP 10.4. Apo-ACP 10.5. Synthesis of Acyl-ACP substrates

* {

396 396 397 400 402 404 404 406 406 407 407 407 410 410 410 410 411 412 415 415 416 416 418 418

Department of Microbiology, University of Illinois, Urbana, Illinois, USA Department of Biochemistry, University of Illinois, Urbana, Illinois, USA


#


395

396

John E. Cronan and Jacob Thomas

10.6. 10.7. 10.8. 10.9.

Purification of fatty acid synthetic enzymes Direct assays of fatty acid synthetic enzyme activities Reconstituted fatty acid synthesis systems Resolution of ACP species by conformationally sensitive gel electrophoresis 10.10. ACPs behave abnormally in SDS-polyacrylamide gel electrophoresis and gel filtration References

420 420 422 423 425 426

Abstract This review presents the most thoroughly studied bacterial fatty acid synthetic pathway, that of Escherichia coli and then discusses the exceptions to the E. coli pathway present in other bacteria. The known interrelationships between the fatty acid and polyketide synthetic pathways are also assessed, mainly in the Streptomyces group of bacteria. Finally, we present a compendium of methods for analysis of bacterial fatty acid synthetic pathways.

1. Introduction The primary role of bacterial fatty acids is to act as the hydrophobic component of the membrane lipids (generally phospholipids). In a number of bacteria, fatty acids also are found as components of storage lipids, the most prevalent being polyhydroxyalkanoic acids (although the lengths of the alkanoic chains are often too short to be considered fatty acids). Other bacteria accumulate storage forms reminiscent of eukaryotic lipids. Actinomycetes such as Streptomyces, Mycobacterium and Rhodococcus accumulate triglycerides (often having unusual acyl chains) whereas, upon growth on hydrocarbons, Acinetobacter and a variety of other hydrocarbon-utilizing bacteria accumulate wax esters in which a long chain fatty acid is esterified to a long chain fatty alcohol. Because accumulation of storage lipids usually depends on exogenous sources of acyl chains rather than de novo fatty acid synthesis, storage lipid fatty acids will not be further discussed. For the same reason postsynthetic modifications of fatty acyl chains such as cyclopropane ring formation (Grogan and Cronan, 1997) and cis-trans isomerization (Cronan, 2002) will also not be discussed.

2. Bacterial Fatty Acids The fatty acids synthesized by bacteria (Fig. 17.1) are similar to the most abundant species present in eukaryotic cells except that the bacterial acids tend to be slightly shorter, generally lack polyunsaturation and the monoenoic C18 acids have different double-bond positions. Moreover,

397

Bacterial Fatty Acid and Polyketide Synthesis

A

COOH

B

COOH

C

COOH

COOH

D

COOH

E OH F

COOH

Figure 17.1 Typical bacterial fatty acids. Structure A is hexadecanoic (palmitic) acid, a very abundant saturated fatty acid whereas structure B is cis-9-hexadecenoic (palmitoleic) acid which together with the product formed by one additional spin of the elongation cycle, cis-11-octadecenoic (cis-vaccenic) acid, are the dominant unsaturated species found in bacteria. Structure C isthe cyclopropanefattyacid cis-9,10-methylenehexadecanoic acid formed by methylene incorporation from S-adenosyl-L-methionine into phospholipid-bound palmitoleic acid.Thistogether with its C19 homologue are distributed very widely in bacteria. Structures D and E are C15 branched chain fatty acids where D is the anteiso species and E is the iso species. Structure F is 3-hydroxytetadecanoic acid, a major component of the lipid Aof most Gram-negative bacteria.

some bacteria make branched chain fatty acids, whereas others make 3-hydroxyacyl acids.

3. Acyl Carrier Protein, the Key Component of Bacterial Fatty Acid Synthesis The mechanism of the synthesis of saturated fatty acids is strongly conserved between bacteria and eukaryotes (the archaea synthesize isoprenoid-derived lipids) although the catalytic entities reside in markedly different protein arrangements. The pathway proceeds in two stages, initiation and cyclic elongation. Intermediates of the pathway are diverted to introduce the double bond of the unsaturated fatty acid species, to provide the 3-hydroxy and short chain fatty acids of the lipid A component of the outer membrane of typical Gram-negative bacteria (Raetz and Whitfield, 2002), the octanoyl-ACP used in the lipoic acid synthetic pathway, the

398


unknown early intermediates of the biotin synthetic pathway and the acyl homoserine lactones of quorum sensing. A key feature of the fatty acid synthetic pathway is that all of the intermediates are covalently bound to a protein called acyl carrier protein (ACP), a small, very acidic and extremely soluble protein (Prescott and Vagelos, 1972). The carboxyl groups of the fatty acyl intermediates are in thioester linkage to the thiol of the 40 -phosphopanthetheine (40 -PP) prosthetic group which in turn is linked to Ser-36 of ACP through a phosphodiester bond. Such ACP thioesters are the substrates for the enzymes of the pathway. ACP is one of the most abundant proteins in E. coli and constitutes about 0.25% of the total soluble protein (6 to 8 104 molecules/cell) (Lu et al., 2007; Rock and Cronan, 1981). Indeed, ACP is reported to be the third most abundant protein present in E. coli (being slightly exceeded by RplL and TufB, two translation apparatus proteins) (Lu et al., 2007). The ACP secondary structure predicted from the amino acid sequence (Rock and Cronan, 1979) has been largely confirmed by high-resolution nuclear (NMR) magnetic resonance spectroscopy and x-ray crystallography. ACP (MW 8860) is composed of a preponderance of acidic residues largely grouped into three a-helices (helices I, II, and IV) oriented in an up-down-down topological arrangement to form a helical bundle plus a short fourth helix (helix III) that seems of lower stability and is found both almost parallel and almost perpendicular to the three helix bundle in the various structures now available (Kim and Prestegard, 1990; Roujeinikova et al., 2002b, 2007). The structural plasticity of ACP seen in the absence of acylation of the prosthetic group thiol is also seen, albeit to a lesser extent, in the acylated forms. The current picture is that ACP and its acylated derivatives can adopt many different structures by sliding and twisting the helices relative to one another and by rearranging the prosthetic group and loops. This plasticity may allow acyl groups to slide in and out of the hydrophobic cavity such that a compromise is achieved between shielding the acyl chain from solvent and allowing access to the thioester-proximal acyl carbon atoms such that fatty acid synthesis can proceed. If so, the dynamics of this process will depend on the polarity and length of the acyl chain and on interactions with the other fatty acid synthetic proteins. The prosthetic group of ACP undergoes metabolic turnover (Fig. 17.2) ( Jackowski and Rock, 1983; Thomas and Cronan, 2005) and the apoprotein is not only inactive in fatty acid synthesis but at high levels can be growth inhibitory (Keating et al., 1995). The primary enzyme catalyzing attachment of the prosthetic group is AcpS (De Lay and Cronan, 2006; Flugel et al., 2000) although a backup enzyme, AcpT, is also present (De Lay and Cronan, 2006; Flugel et al., 2000). AcpS and AcpT are 40 -phosphopantetheine transferases (40 -PP transferases) that transfer the 40 -phosphopantetheine portion from CoA to apo ACP to give holo-ACP plus 30 , 50 -ADP. Although the known 40 -PP transferases comprise a single

399


O

3´ 5´ -ADP

O P

O

O

OH

OH

OH SH

N H

HN

O

+

ACP

Coenzyme A

Apo-ACP

AcpS

ADP

3´ 5´ -ADP O

ATP O

Dephospho-coenzyme A

ACP

OH P

HN O

N H

SH

O

O

OH

Holo-ACP

PPi ATP

AcpH

O OH O- P

HN O

O

N H O

OH

SH

+

OH

ACP Apo-ACP

4´ - phosphopantetheine

Figure 17.2 ACP prosthetic group metabolism in E. coli. Apo-ACP, the product of the acpP gene, is nonfunctional in fatty acid synthesis. It is post-translationally activated by the attachment of a 40 -phosphopantetheine arm derived from coenzyme A, a reaction catalyzed by the holo-ACP synthase AcpS. Turnover of ACP involves removal of the prosthetic groupbyACP hydrolyase, AcpHwhichcan then be used in the synthesis of CoA.

protein superfamily, the quaternary structures of these proteins vary from monomers to trimers with E. coli AcpS reported to be dimeric (Lambalot and Walsh, 1997; McAllister et al., 2006). Unlike the enzymes that attach biotin and lipoic acid to their cognate proteins, the 40 -PP transferases are generally quite specific for their protein substrates (Lambalot et al., 1996) (see below). Although E. coli AcpS attaches the prosthetic group to many fatty acid ACPs, some other fatty acid ACPs are not substrates (De Lay and Cronan, 2007) and it fails to modify the structurally related carrier proteins of polyketide and nonribosomal polypeptide synthesis (Lambalot et al., 1996). However, Bacillus subtilis Sfp, which is responsible for modification of the carrier protein used in synthesis of surfactin, a nonribosomal peptide, readily modifies most carrier proteins regardless of origin or metabolic role (Lambalot et al., 1996). It is thought that ACP helix II is a major determinant of this substrate specificity (Mofid et al., 2002) Surprisingly, all known 40 -PP transferases accept acylated-CoA substrates and transfer acyl-40 -PP moieties to apo-ACP. E. coli AcpS utilizes acetyl-ACP in place of ACP at about half the catalytic efficiency and is

400


also active with butyryl-, acetoacetyl-, and malonyl-CoAs (McAllister et al., 2006). Again, Sfp is much more promiscuous and transfers acyl-40 PP moieties modified with molecules such as biotin and fluorescein (Mercer and Burkart, 2007). It is unclear whether or not transfer of acyl-40 -PP moieties is physiologically relevant. However, the low levels of AcpS in E. coli argue that acyl-40 -PP transfer should be unable to replace a fatty acid synthetic enzyme such as FabD. Soon after the identification of the prosthetic group of ACP as 40 -PP and of the AcpS 40 -PP transferase, Vagelos and Larrabee (1967) reported an E. coli activity that removed the 40 -PP moiety of ACP (Fig. 17.2). The enzyme (called ACP hydrolyase and ACP phosphodiesterase) was purified (Fischl and Kennedy, 1990). ACP hydrolyase was shown to be an unusually stable enzyme of molecular weight 25,000 and an N-terminal sequence was reported. Unfortunately, the N-terminal sequence was found to be that of AzoR, a flavincontaining protein later shown to be an azoreductase (Nakanishi et al., 2001). Hence, it is clear that the N-terminal sequence determined was the sequence of a major contaminating protein rather than that of the phosphodiesterase. Regrettably this erroneous sequence attribution has led to many genes (often called acpD) being annotated in bacterial genomes as encoding ACP phosphodiesterase rather than azoreductase (Nakanishi et al., 2001). Thus, in order to prevent confusion with the mistaken AcpD annotations the recently identified gene was named acpH (Thomas and Cronan, 2005) based on the original enzyme name (ACP hydrolyase) given by Vagelos and Larrabee (Vagelos and Larrabee, 1967). The acpH gene was identified by expression cloning and was found to encode a protein of 23 kDa that readily aggregates upon overexpression (Thomas and Cronan, 2005; Thomas et al., 2007). Active enzyme was recovered by folding solubilized inclusion bodies and AcpH was found to be active on acyl-ACPs of fatty acyl chain lengths from C6 to C16. AcpH was active on Bacillus subtilis ACP, but inactive on Lactococcus lactis ACP. Strains carrying deletions of acpH are fully viable, but the ACP prosthetic group is metabolically stable unlike that of wildtype strains (Thomas and Cronan, 2005). Upon AcpH overproduction all of the cellular ACP is converted to the apo form (Thomas and Cronan, 2005). AcpH is not essential either in the laboratory or in the natural habitat (Thomas and Cronan, 2005). AcpH has been shown to be a noncanonical member of the HD phosphatase/phosphodiesterase family (Thomas et al., 2007).

4. Overview of the Reactions of Fatty Acid Biosynthesis The fatty acid synthesis system of E. coli is the archetype of the type II or dissociated fatty acid synthesis systems and the E. coli gene names are very often used for their homologues in other organisms. The precursors for fatty acid biosynthesis are derived from the acetyl-CoA pool. Malonyl-CoA is

401


required for all the elongation steps and is formed by the first step in fatty acid biosynthesis catalyzed by acetyl-CoA carboxylase. Malonyl-CoA is utilized for fatty acid biosynthesis only following its conversion to malonyl-ACP by malonyl-CoA:ACP transacylase (Fig. 17.3). Acetyl-CoA carboxylase will not be further discussed in this review since strictly speaking the enzyme is not a dedicated fatty acid synthesis enzyme because malonyl-CoA is used in synthesis of other molecules (e.g., polyketides). Moreover, a recent review of the E. coli and other multisubunit acetyl-CoA carboxylase is available (Cronan and Waldrop, 2002). There are three known mechanisms for the initiation of fatty acid biosynthesis in E. coli. First, FabH (3-ketoacyl-ACP synthase III) catalyzes the condensation of acetyl-CoA with malonyl-ACP to yield acetoacetylACP (Fig. 17.3). In the second pathway, the acetate moiety is first transferred from acetyl-CoA to acetyl-ACP by the transacylase activity of FabH. The acetyl-ACP is then condensed with malonyl-ACP by FabB (synthase I) or alternatively by FabF (synthase II). The third pathway involves the decarboxylation of malonyl-ACP by FabH, FabB or FabF to form acetylACP followed by subsequent condensation with malonyl-ACP. The evidence for the existence of these pathways and their relative contributions to the initiation of fatty acid biosynthesis is an area of current interest and is discussed in more detail below. The reactions of the fatty acid elongation cycle are outlined in Fig. 17.4. The first step is the Claisen condensation of malonyl-ACP with a growing acyl chain catalyzed by a 3-ketoacyl-ACP synthase (either FabB or FabF). This is the only irreversible step in the elongation cycle and thus it is not surprising that the 3-ketoacyl-ACP synthases play key roles in regulating the product distribution of the pathway. The 3-keto-thioester produced is reduced by FabG, an NADPH-dependent 3-ketoacyl-ACP reductase, followed by removal of a water molecule by FabZ, a 3-hydroxyacyl-ACP dehydratase. The final reduction is catalyzed by FabI, an enoyl-ACP reductase, to give an acyl-ACP, which can serve as a substrate for another round of elongation or, if of sufficient chain length, be transferred into complex lipids. The equilibrium of the FabI enoyl-ACP reductase reaction acts to pull the reversible 3-ketoacyl-ACP reductase and 3-hydroxyacyl-ACP dehydratase reactions to the right (Heath and Rock, 1995). Malonyl-CoA ACP AcpS

AcpH

P-PanSH CoA Apo-ACP

FabD

Acetyl-CoA FabH

Malonyl-ACP Acetoacetyl-ACP

Figure 17.3 The initiation steps in the type II fatty acid synthesis pathway of E. coli. Malonyl-CoA is converted to malonyl-ACP by malonyl transacylase (FabD). Fatty acid synthesis is initiated by FabH, which condenses malonyl-ACP with acetyl-CoA.

402


(NADP+) NAD+ NADH (NADPH)

Acyl-ACP

FabI

Enoyl-ACP

E.coli fatty acid synthesis

Malonyl-ACP FabB FabF

ACP

3-Ketoacyl-ACP NADPH

H2O

FabG

FabZ 3-OH-acyl-ACP

NADP+

Figure 17.4 Production of unsaturated fatty acids. Unsaturated fatty acids arise from a branch in the biosynthetic pathway at the 3-hydroxydecanoyl-ACP intermediate. FabA is a unique 3-hydroxyacyl-ACP dehydratase that is capable for forming trans2-decenoyl-ACP and isomerizing this intermediate to cis-3-decenoyl-ACP. FabA is very selective for the 10-carbon substrates in vivo. FabB is absolutely required for the subsequent elongation of cis-3-decenoyl-ACP to 16:1-ACP. Interestingly, 16:1-ACP is a poor substrate for FabB, and its elongation to 18:1-ACP is controlled by the activity of FabF. FabF is a naturally temperature-sensitive enzyme, and this property account for the greater proportion of 18:1 in bacteria grown at low temperatures compared to those grown at higher temperatures. The trans-3-decenoyl-ACP is used by FabI followed by elongation by either FabB or FabF to form palmitic acid, the major saturated fatty acid in E. coli.

Two of the reactions of the cycle can be carried out by multiple discrete enzymes. As noted above in g-proteobacteria such as E. coli there are three 3-ketoacyl-ACP synthases and two 3-hydroxyacyl-ACP dehydratases are also present with one of each playing a key role in unsaturated fatty acid synthesis. Due to their differing substrate specificities, each isozyme makes a unique contribution to the regulation of the distribution of products from the pathway (see below). In most other bacteria that use the type II pathway there are two 3-ketoacyl-ACP synthases and two 3-hydroxyacyl-ACP dehydratases. However, in bacteria that synthesize branched chain fatty acids there can be three 3-ketoacyl-ACP synthases. The fatty acid biosynthetic pathway ends in the transfer of the acyl chains of the acyl-ACP end products by various acyltransferase systems. In general, transfer of the acyl chain is to a glycerol-based backbone molecule to form membrane lipids.

5. The Initiation Steps of Fatty Acid Synthesis The first step in fatty acid synthesis is condensation of a primer acylCoA with malonyl-ACP (Fig. 17.3). The conversion of malonyl-CoA to malonyl-ACP is catalyzed by malonyl-CoA:ACP transacylase, the product


403

of the fabD gene (Magnuson et al., 1992; Verwoert et al., 1992). FabD is a monomeric protein that accepts the malonyl moiety from malonyl-CoA to form a stable malonyl enzyme intermediate in which the malonyl moiety is in ester linkage to a serine hydroxyl (Oefner et al., 2006; Serre et al., 1995). Nucleophilic attack of this intermediate by the sulfhydryl of ACP gives malonyl-ACP, the major building block of fatty acids. Crystal structures of FabD, FabD complexed with malonate and the FabD-malonyl-CoA complex have been reported (Oefner et al., 2006; Serre et al., 1995). It should be noted that several polyketide ACPs have been reported to self-malonate (Arthur et al., 2005, 2006; Hitchman et al., 1998). That is, the malonyl moiety is transferred from CoA to ACP in the absence of malonyl-CoA: ACP transacylase. This reaction will be discussed below. In contrast to the reactions that produce malonyl-ACP, the reactions whereby the methyl carbon atom and the immediately adjacent carbon atom (the last two carbons of the fatty acid chain in chemical nomenclature) are incorporated into fatty acid remain somewhat unclear. In straight chain acids the methyl and penultimate carbons are derived from acetyl-CoA. Acetyl-CoA is a substrate for FabH (3-ketoacyl-ACP synthase III, which has also been called acetoacetyl-ACP synthase) and is incorporated directly to form the first four-carbon fatty acyl-ACP species ( Jackowski and Rock, 1987; Tsay et al., 1992a). In E. coli malonyl-ACP is utilized only in the elongation steps in fatty acid biosynthesis. However, FabH, FabB, and FabF are capable of initiating fatty acid synthesis in vitro in the absence of an added acetyl-ACP or acetyl-CoA as the primer. This synthesis occurs through a side reaction; decarboxylation of malonyl- ACP to give acetyl-ACP. This reaction is readily demonstrated in vitro (Alberts et al., 1972; McGuire et al., 2001), but its role in initiation in vivo awaits experimental verification. FabH has been shown to be essential for fatty acid synthesis in Lactococcus lactis by construction of a deletion mutant (Lai and Cronan, 2004). However, the deleted strain retains about 10% of the normal fatty acid synthetic ability indicating a partial bypass of FabH activity (Lai and Cronan, 2004). This seems likely to be due to malonyl-ACP decarboxylation followed by FabFcatalyzed condensation of the resulting acetyl-ACP with malonyl-ACP. However, this scenario requires experimental testing. Another possible source of acetyl-ACP would be transfer of acetyl-40 -PP from acetyl-CoA to apo ACP catalyzed by AcpS (see above). However, the intracellular level of AcpS activity seems far too low to provide the needed carbon flow. In the synthesis of branched chain acids acetyl-CoA is replaced by isovaleryl-CoA, isobutyryl-CoA, or 2-methylbutyryl-CoA, which are derived from the metabolic pathways for the amino acids, valine, leucine, and isoleucine (Wallace et al., 1995). The enzyme responsible for the synthesis of these acyl-CoAs is a specialized branched-chain–keto acid dehydrogenase complex, and mutations in the activity of this enzyme system result in strains that are auxotrophic for branched-chain acids

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(Cropp et al., 2000). Bacillus subtilis has two FabH homologues that accept the branched acyl-CoAs (Choi et al., 2000a) whereas Streptomyces glaucescens has only a single such enzyme (Han et al., 1998). It should be noted that the branched chain organisms also contain some straight chain acids and this is reflected in the fact that their FabH enzymes accept acetyl-CoA, although branched chain primers are the preferred substrates. In contrast, E. coli FabH ignores branched chain primers (Choi et al., 2000a).

6. The Enzymes of the Fatty Acid Elongation Cycle 6.1. The 3-ketoacyl-ACP synthase reaction (FabB, FabF, FabH) The first reaction of each cycle is the Claisen condensation of an acyl thioester (acyl-ACP or for FabH, acetyl-CoA) with malonyl-ACP to form a 3-ketoacyl-ACP plus ACP (or CoA). Three E. coli enzymes are known to catalyze 3-ketoacyl-ACP synthase reactions. These enzymes were referred to as synthases I, II and III, but more recently have come to be called FabB, FabF and FabH, respectively, after their gene names. The fabH and fabF genes are located within the fatty acid synthetic gene cluster (Rawlings and Cronan, 1992) whereas fabB maps alone at a distant site. Siggaard-Andersen and coworkers (1994) reported a putative fourth KAS activity in E. coli and assigned an open reading frame to this activity. However, this report was based on a series of indirect inferences and was in error; the gene sequenced was the fabF gene (Magnuson et al., 1995; Rawlings and Cronan, 1992) and the enzyme activity described seems likely to be a mixture of FabH with FabB and/or FabF. The three E. coli 3-ketoacyl-ACP synthases represent two classes of decarboxylating Claisen condensing enzymes (Heath and Rock, 2002; White et al., 2005). The FabB and FabF enzymes have Cys-His-His active sites whereas FabH has a CysHis-Asn active site triad. These differences are reflected in the rest of the primary sequences of the proteins. FabB and FabF are about 37% identical whereas alignment of either FabB or FabF with FabH gives only scattered alignments of very low quality. High-resolution crystal structures of all three enzymes are available (White et al., 2005). The functional FabB is composed of two identical subunits (Garwin et al., 1980b) and contains both malonyl-ACP and fatty acyl-ACP binding sites (D’Agnolo et al., 1975b). In the condensation reaction, the acyl group becomes covalently linked to a specific FabB cysteine sulfhydryl (D’Agnolo et al., 1975b). The acyl-enzyme undergoes condensation with malonylACP to form 3-ketoacyl-ACP, CO2, holo-ACP and free enzyme. Inhibition studies using cerulenin (see below) show the active site cysteine to be Cys-163 (Kauppinen et al., 1988).


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Further investigation revealed the presence of a second long chain synthase activity in E. coli, FabF (D’Agnolo et al., 1975a). Like FabB, FabF has a dimeric structure and is inhibited by cerulenin (Ulrich et al., 1983), although FabF is less sensitive to the antibiotic than is FabB and is not essential for growth of E. coli. Both synthases are both capable of participating in saturated and unsaturated fatty acid synthesis. The enzymes have been shown, in vitro, to function similarly with all substrates except palmitoleoylACP; palmitoleoyl-ACP is an excellent substrate for FabF, but not for FabB (Edwards et al., 1997; Garwin et al., 1980b). This observation is consistent with the role of FabF in the regulation of fatty acid composition of the membrane phospholipid in response to temperature (see below). Mutant strains having fabB missense or null mutations (Garwin et al., 1980a), however, require unsaturated fatty acids for growth. Therefore, in vivo FabB must catalyze a key reaction in unsaturated fatty acid synthesis that FabF cannot. This reaction is probably the elongation of cis-3-decenoylACP, although this has not been demonstrated experimentally. This step is the rate-limiting step in unsaturated fatty acid synthesis (Clark et al., 1983). FabB overproduction has two known effects. First, overproduction of the enzyme overcomes its poor ability to elongate palmitoleoyl-ACP and an increased amount of cis-vaccenic acid is incorporated into phospholipid (de Mendoza et al., 1983). The increase, however, has no effect on the temperature regulation of fatty acid composition (de Mendoza et al., 1983). Second, excess cellular FabB renders E. coli resistant to the antibiotic thiolactomycin which is an inhibitor of all three 3-ketoacyl-ACP synthases. It was thought that FabB-catalyzed decarboxylation of malonyl-ACP offered the cell an alternative initiation pathway for fatty acid biosynthesis; malonyl-ACP decarboxylation to give acetyl-ACP that was used as a primer for chain elongation. The isolation of thiolactomycin-resistant strains that have a resistant FabB has borne out this hypothesis in vivo ( Jackowski et al., 2002). Moreover, in the presence of the antibiotic, excess FabB appears to allow the cell to bypass the standard FabH initiation pathway probably by decarboxylation of malonyl-ACP to form acetyl-ACP ( Jackowski et al., 2002; Tsay et al., 1992b). Given the above observations, FabB appears to be the only E. coli 3-ketoacyl-ACP synthase absolutely required for growth ( Jackowski et al., 2002; Tsay et al., 1992b). FabH is a dimeric protein of 33.5 kDa first detected as a condensation activity resistant to cerulenin both in vivo and in vitro ( Jackowski and Rock, 1987). Although cerulenin blocks the synthesis of long chain fatty acids, short chain (C4 to C8) acids linked to ACP are accumulated both in vivo and in cell extracts. The fabH gene has been shown to encode KAS III. As noted above the FabH sequence has no similarity to those of FabB or FabF, although there is good alignment with other enzymes known to catalyze condensation reactions. From the chain length of the acyl-ACPs produced and the behavior of fabB fabF double mutants mentioned above, it is clear

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that FabH does not participate in the progressive condensation steps of fatty acid synthesis that produce the long chain acids found in the phospholipids and lipid A. However, the enzyme could produce the fatty acid synthetic intermediates used in lipoic acid (and perhaps biotin) synthesis. However, the high activity of the enzyme suggests that it does not exclusively function in this pathway (E. coli requires only a few hundred lipoic acid molecules per cell).

6.2. 3-ketoacyl-ACP reductase (FabG) An open reading frame encoding a protein with strong similarities to several acetoacetyl-CoA reductases and particularly plant 3-ketoacyl-ACP reductases (>50 % identical residues) was found between the fabD and acpP genes in the fab gene cluster (Rawlings and Cronan, 1992). This gene was designated as fabG and is obligatorily co-transcribed with the upstream genes (Zhang and Cronan, 1996, 1998). Blocking fabG transcription blocked cell growth indicating that it is an essential gene (Zhang and Cronan, 1998). However, no mutants in this gene were available until the recent isolation of temperature-sensitive mutants of both E. coli and Salmonella enterica (Lai and Cronan, 2004). Strikingly, all of the mutations were located in or near the subunit interfaces of the FabG homotetramer suggesting that monomers and dimers of the enzyme are inactive. At the nonpermissive temperature, fatty acids synthesis was blocked in the fabG(Ts) mutants following the initial condensation such that only four carbon acyl-ACP species accumulated. Thus, only a single NADPH-specific 3-ketoacyl-ACP reductase exists in E. coli and it functions with all acyl chain lengths (Lai and Cronan, 2004). Crystal structures of FabG are available (White et al., 2005).

6.3. The 3-hydroxyacyl-ACP dehydratase (FabZ) This enzyme is not to be confused with the 3-hydroxydecanoyl-ACP dehydratase specifically required for introduction of the double bond of the unsaturated acids in E. coli and related bacteria. Prior biochemical data gave a puzzling picture of this reaction. One group reported that this step is catalyzed by a single enzyme active with substrates of all chain lengths (Birge et al., 1967) whereas another laboratory reported the presence of three enzymes specific for short, medium, and long chain length substrates (Mizugaki et al., 1968). More recent work with recombinant FabZ showed that the dehydratase efficiently catalyzed the dehydration of short chain 3-hydroxyacyl-ACPs and long chain saturated and unsaturated 3-hydroxyacyl-ACPs (Heath and Rock, 1996). The fabZ gene was isolated as a suppressor of a mutation in lipid A biosynthesis. The suppression is thought to be due to increased intracellular levels of 3-hydroxymyristoyl-ACP (Mohan et al., 1994). Unfortunately, the


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phenotype of the original mutant strain is too feeble for physiological studies. Although no structure of E. coli FabZ is presently available, several FabZ homologue structures have been reported the most closely related of which is that of the Pseudomonas aeruginosa protein (Kimber et al., 2004). This FabZ shows sequence homology to FabA (which was a key to its discovery) and thus it was no surprise that the monomers have a hot-dog fold (see Section 7.1) and the active sites lie at the dimer interface. FabZ has a Glu residue in place of the active site Asp residue of FabA. This substitution was thought to play an important role in the differing specificities of the two enzymes. However, the discovery of a FabZ homologue (FabN) that has FabA activity and has a Glu active site residue (Wang and Cronan, 2004) indicates that this simple hypothesis is incorrect.

6.4. The enoyl-ACP reductase (FabI) Two forms of enoyl-ACP reductase, the last enzyme of the fatty acid cycle, were originally reported, one dependent on NADH and the other on NADPH (Weeks and Wakil, 1968). However, both activities were subsequently shown to be due to the same enzyme, FabI (Bergler et al., 1996), the sole enoyl-ACP reductase of E. coli (Heath and Rock, 1995). The identification of the protein encoded by this gene, now called fabI, was the result of studying mutants of E. coli and S. enterica resistant to diazaborines, a class of potent antimicrobial agents that inhibit lipid synthesis (Turnowsky et al., 1989). FabI is also the target of the widely used antimicrobial compound, triclosan (Heath et al., 1999; McMurry et al., 1998). FabI plays a role in the fatty acid synthesis cycle in that its action pulls the other reversible steps of the cycle (FabG, and FabZ) to the right such that each cycle of fatty acid biosynthesis is completed (Heath and Rock, 1995). Several FabI crystal structures (White et al., 2005) and a useful fabI temperature sensitive mutant (Bergler et al., 1994) are available. It should be noted that E. coli FabI has been reported to be inhibited by palmitoyl-CoA (Bergler et al., 1996). Although this has been ascribed a physiological role, it remains to be demonstrated that the observed inhibition is not due to the well-known detergent properties of long chain acyl-CoAs.

7. Unsaturated Fatty Acid Synthesis in E. coli 7.1. The 3-hydroxydecanoyl-ACP dehydratase (FabA) This enzyme essentially extracts a major fraction of the 3-hydroxydecanoylACP from the elongation cycle and introduces a cis double bond (Fig. 17.4). Following elongation of the cis-3-decenoyl-ACP by FabB (Fig. 17.5), these modified acyl chains are returned to the cycle for elongation to the long

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Octanoyl-ACP + Malonyl-ACP

3-OH-10-ACP FabA FabZ 10:1Δ2t-ACP FabB FabF 16:0-ACP

FabA (FabM) (FabN)

10:1Δ3c-ACP FabB (FabO) 16:1-ACP FabF 18:1-ACP

Figure 17.5 The elongation cycle. There are four steps in fatty acid elongation. Each new cycle of two-carbon elongation is initiated by the condensation of acyl-ACP and malonyl-ACP by one of the elongation condensing enzymes, FabB or FabF. The next step is the reduction of the 3-ketoacyl-ACP by the NADPH-dependent FabG. This 3-hydroxylacyl-ACP is dehydrated to trans-2-acyl-ACP by FabZ. The final step is the NADH-dependent reduction of enoyl-ACP to acyl-ACP by FabI.The parentheses show isozymes from other bacteria that catalyze the reaction shown.

chain unsaturated acids needed for phospholipid function (Bloch, 1971). 3-Hydroxydecanoyl-ACP dehydratase specifically catalyzes the dehydration of 3-hydroxydecanoyl-ACP to a mixture of trans-2-decenoyl-ACP and cis-3-decenoyl-ACP (Bloch, 1971). The reaction proceeds via the formation of trans-2-decenoyl-ACP as an enzyme-bound intermediate which can disassociate from the enzyme (Bloch, 1971). When disassociation occurs, the trans-2 intermediate is reduced by an enoyl-ACP reductase and subsequently converted to saturated fatty acids as in the standard elongation cycle (Clark et al., 1983; Guerra and Browse, 1990; Heath and Rock, 1996). Enzyme-bound trans-2-decenoyl-ACP, however, is isomerized to cis-3-decenoyl-ACP. The double bond is preserved and the cis-3 intermediate is elongated to the unsaturated fatty acids of E. coli, palmitoleic acid, and cis-vaccenic acid. The enzyme, a homodimer of 18 kDa subunits (Leesong et al., 1996), is distinct from the elongation cycle FabZ dehydratase discussed above, although the two enzymes contain sequences in common. The first mutants isolated that were blocked in fatty acid biosynthesis, called fabA, lacked 3-hydroxydecanoyl-ACP dehydratase (Silbert and Vagelos, 1967). These mutants are unable to synthesize unsaturated fatty acids, but synthesize saturated fatty acids normally. In vitro mutant fabA enzymes could form neither the cis-3 nor trans-2-decenoyl products (Cronan and Gelmann, 1973). This finding, along with the observation that saturated fatty acid synthesis continues in vivo indicated that another


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dehydratase was available for saturated fatty acid synthesis, the enzyme, now called FabZ, which catalyzes the formation of trans-2-decenoyl-ACP, but cannot catalyze the isomerase reaction (Heath and Rock, 1996). In the absence of thermal regulation, the ratio of unsaturated to saturated fatty acids in E. coli is dependent on the levels of FabA and FabB. It was shown that overproduction of FabA in vivo failed to increase the level of unsaturated fatty acids, but significantly increased the amount of saturated fatty acids incorporated into membrane phospholipids (Clark et al., 1983). This indicated that, although FabA is required for the synthesis of unsaturated fatty acids, the level of enzyme activity does not limit the rate of unsaturated fatty acid synthesis. Introduction of multiple copies of the fabB gene (encoding synthase I) reversed the effect of dehydratase overproduction resulting in wildtype fatty acid compositions (Clark et al., 1983). Thus, the step more likely to limit the rate of unsaturated fatty acid synthesis is the elongation of cis-3-decenoyl-ACP catalyzed by FabB. The levels of expression of the fabA and fabB genes, therefore, appear to establish a basal ratio of unsaturated to saturated fatty acid synthesis in the absence of thermal regulation. Modulation of the fatty acid composition of membrane phospholipid in response to temperature shift is discussed below. FabA functions with acyl chain lengths from C4 to C12 in an in vitro fatty acid synthesis system reconstituted from purified enzymes (Heath and Rock, 1996). This is a considerably wider range of FabA substrates than originally reported in studies using synthetic model substrates (Bloch, 1971). The reasons for this disagreement are unclear, but it should be noted that the ratios of products produced by the enzyme using acyl-ACP substrates (Guerra and Browse, 1990) differs from that obtained using model substrates (Bloch, 1971). The mechanism of FabA has been thoroughly studied (Bloch, 1971; Schwab and Henderson, 1990) since it was the first enzyme for which a mechanism-activated ("suicide") inhibitor was described. This inhibitor, 3-decynoyl-N-acetylcysteamine, forms a covalent adduct with the active site histidine residue resulting in loss of all of the partial reactions of the enzyme (Bloch, 1971; Cronan et al., 1988; Leesong et al., 1996) The x-ray crystal structure of E. coli FabA was the first structure determined for an E. coli fatty acid synthetic protein. FabA was the first example of the ‘‘hot-dog’’ fold in which a long central a-helix is wrapped by a six-stranded antiparallel b-sheet (Leesong et al., 1996). FabA forms an unusually stable dimer and it seems that the dimerization is essential for activity since the two active sites are formed along the dimer interface with the critical His and Asp active site residues being contributed by different monomers (Leesong et al., 1996).

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8. The Abundant Exceptions to the E. coli Fatty Acid Synthesis Paradigm 8.1. Branched-chain fatty acids The synthesis of branched-chain fatty acids by use of FabH homologues has been discussed above. Such fatty acids occur widely in bacteria (e.g., Bacilli, Streptomyces, Staphylococci).

8.2. Anaerobic synthesis of unsaturated fatty acids The classical pathway of anaerobic unsaturated fatty acid synthesis in E. coli is not widely distributed in bacteria. Genomic analyses indicate that only the a- and g-proteobacteria encode the proteins of this pathway. Most other bacteria including many pathogens make unsaturated fatty acids under anaerobic conditions, but lack recognizable homologues of the key E. coli unsaturated fatty acid synthetic enzymes, FabA and FabB. Thus far, Streptococcus pneumoniae has been shown to introduce the cis double bond using FabM (Marrakchi et al., 2002), an isomerase of a sequence unrelated to FabA, that performs only one of the FabA reactions (FabA is both an isomerase and a dehydratase). In contrast, Enterococcus faecalis was shown to have proteins that are homologues of FabZ and FabF that perform the roles of FabA and FabB, respectively (Wang and Cronan, 2004). This work has been extended (Altabe et al., 2007; Lu et al., 2005). However, riddles remain in a number of bacteria (e.g., the clostridia and the lactic acid bacteria). Pseudomonas aeruginosa has not only the classical FabA-FabB pathway, but also two oxygen-requiring desaturase pathways, one of which uses endogenous phospholipid acyl chains whereas the other uses exogenous saturated fatty acids (Cronan, 2006a; Zhu et al., 2006). The bacilli also have desaturases that are induced by low-growth temperatures that utilize endogenous phospholipid acyl chains (Mansilla and de Mendoza, 2005). Other bacteria lack FabM, FabA, FabB, and desaturases, and lack extra copies of genes encoding FabZ and FabF homologues. Therefore, it seems that other pathways for unsaturated synthesis exist.

8.3. Diversity of enoyl-ACP reductases A curious and important divergence is seen in the conservation of bacterial fatty acid synthetic (FAS) proteins. Most FAS II enzymes are strongly conserved among bacteria both in sequence and (where data are available) in structure and most domains of the FAS I proteins are clearly derived from FAS II proteins (Cronan, 2004, 2006b). An exception is the last step of the elongation cycle, formation of a saturated acyl-ACP by reduction (generally NAD(P)H-dependent) of the enoyl-ACP double bond. As mentioned above


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in E. coli this reaction is catalyzed by the product of the fabI gene that was discovered as the target for the antibacterial action of a set of diazaborine compounds (Bergler et al., 1994). FabI was later shown to also be the site of the antibacterial action of triclosan (McMurry et al., 1998), a compound used in antibacterial hand soaps and a large variety of other everyday products. Although FabI homologues are widely distributed in bacteria and other FAS II-containing organisms, the existence of a number of bacterial species naturally resistant to triclosan was soon recognized (Heath et al., 1999; Marrakchi et al., 2003). In these bacteria triclosan resistance was due to the presence of enoyl-ACP reductase isozymes of varying resistance to triclosan. Bacillus subtilis contains two isozymes, a FabI homologue and an isozyme called FabL that is moderately resistant to triclosan (Heath et al., 2000) and, like FabI, is a member of the short chain dehydrogenase reductase superfamily. Streptococcus pneumoniae contains a single enoyl-ACP reductase, FabK, that is refractory to triclosan and is a TIM barrel flavoprotein unrelated to the short chain dehydrogenase reductase isozymes (Marrakchi et al., 2003). A fourth class of enoyl-ACP reductase is present in vibrios and related bacteria (Massengo-Tiasse and Cronan, 2008). The diversity of the bacterial enoylACP reductases when compared to the lack of structural and mechanistic diversity seen in the other enzymes of the FAS II elongation cycle argues that naturally occurring compounds exist that selectively inhibit one or another of these enzymes. This hypothesis has recently been confirmed by the recent discoveries of natural enoyl-ACP reductase inhibitors of fungal origin that specifically target FabI (Cephalochromin) (Zheng et al., 2007) and FabK (Atromentin and Leucomelone) (Zheng et al., 2006).

8.4. Type I megasynthase fatty acid synthesis The most striking divergence from the E. coli type II fatty acid synthetic pathway is the type I megasynthase pathway found in the mycolic acidproducing branch of the Actimomycetales (mycobacteria, corynebacteria, rhodococci, and nocardiae) (Schweizer and Hofmann, 2004). Mycolic acids are high-molecular-weight a-alkyl b-hydroxy fatty acids (70 to 90 carbon atoms in length) composed of a species-specific ‘‘short’’ arm of 22 to 26 carbon atoms and a ‘‘long’’ meromycolic acid arm of 50 to 60 carbon atoms (although this arm is shorter in corynebacteria) (Gokhale et al., 2007; Schweizer and Hofmann, 2004). These essential molecules are found in the matrix of the cell wall and are formed by head-to-head Claisen-type condensations of two long-chain fatty acyl thioesters followed by reduction of the keto group. De novo synthesis of the long-chain fatty acid esters is accomplished by several strategies. In mycobacteria, the type II system is unable to perform de novo synthesis, which is the function of the type I system. The type I system synthesizes C16 and C26 saturated acyl-CoAs. The C16-CoA is elongated by the type II FabH protein and

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successive condensation (by two very long-chain FabF-like enzymes) and reduction cycles to give the C56-CoA. This is condensed with the C26CoA by the Pks13 polyketide synthase that catalyzes the Claisen condensation. Reduction of the resulting ketone by specific reductase gives the mycolic acids (Gokhale et al., 2007). It should be noted that the type I system may play only a minor role in the growth of pathogenic mycobacteria since they readily acquire host fatty acids. Excepting its specificity for long-chain lengths, the mycobacterial type II system is fairly typical. The multifunctional fatty acid synthesis (FAS) polypeptides of mycobacterial and corynebacteria contain all the functional domains required for de novo fatty acid synthesis (Schweizer and Hofmann, 2004). These domains are organized in the following order: acyltransferase, enoyl reductase, dehydratase, malonyl/palmitoyl transferase, acyl carrier protein, 3-ketoacyl reductase and 3-ketoacyl synthase. All intermediates generated remain enzyme-bound during the process of elongation and undergo transacylation to other catalytic sites within the enzyme. FAS I generates short-chain, fatty acyl-CoA primers that are further elongated by the type II system. It is remarkable that the domain order of the bacterial type I protein is precisely that given by in silico fusion of the yeast b and a fatty acid synthase subunits with b being the N-terminal partner (Schweizer and Hofmann, 2004). Moreover, both the bacterial type I protein and yeast enzymes are functional hexamers. It will be interesting to see if the bacterial type I protein has the fantastic porous barrel architecture of the fungal enzyme (Cronan, 2006c). Corynebacteria synthesize unsaturated fatty acids by use of a second type I synthase that contains a FabA-like domain (Schweizer and Hofmann, 2004). Mutational inactivation of the gene encoding this second synthase results in an unsaturated fatty acid requirement. However, some corynebacteria lack this second enzyme and make only saturated species (Schweizer and Hofmann, 2004), whereas others lack a type I protein and require fatty acids for growth (Tauch et al., 2005). Finally, it should be noted that proteins annotated as fatty acid synthetic proteins based on sequence homologies with the E. coli proteins can have no involvement in fatty acid synthesis. For example a Pseudomonas aeruginosa FabH homologue catalyzes a key step in the synthesis of an extracellular quinoline derivative (Zhang et al., 2008) that acts as a quorum-sensing molecule and a Mesorhizobium FabH homologue plays a role in biotin synthesis (Sullivan et al., 2001).

9. Relationships between Fatty Acid Synthesis and Polyketide Synthesis There seems no doubt that polyketide synthesis is an evolutionary descendant of fatty acid synthesis. The fatty acid synthase (FAS-I) responsible for de novo fatty acid synthesis in the cytosol of animal cells is


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homologous in sequence and analogous in architecture with the very large family of type I modular polyketide synthases (PKSs). Both ‘‘megasynthase’’ assembly lines use homologous domains (ACP or PCP) to carry the growing fatty acid or polyketide via a pantetheine-linked thioester (Smith and Sherman, 2008). The common thioester chemistry, similar structures and the adaptable architecture have resulted in the proliferation of hybrid PKSFAS pathways found in phylogenetically diverse bacteria (Smith and Sherman, 2008). Indeed, the very complex lipids of mycobacteria are made via a partnership between an FAS and a modular PKS (Gokhale et al., 2007) and a class of fatty acids are produced by modular PKS systems (Metz et al., 2001; Wallis et al., 2002). This close relationship raises the question of whether or not FAS and PKS systems share enzymatic components. The two systems clearly share the precursors, acetyl-CoA and malonyl-CoA, but this is probably not a competitive situation because fatty acid synthesis proceeds primarily in growing cells, whereas polyketides are secondary metabolites whose synthesis generally occurs after a culture ceases net growth. Indeed, ectopic expression of sets of whiE-PKS genes presumed sufficient to assemble a carbon chain caused inhibition of early growth of the strains. This growth inhibition was not a nonspecific effect of the protein expression because it required all three genes to be expressed (subsets of two of the proteins gave no inhibition of growth). It has been reasonably proposed that this growth inhibition is due to competition between fatty acid synthesis and the inappropriately timed polyketide synthesis for precursors or proteins common to the two pathways (Yu and Hopwood, 1995). This hypothesis remains to be tested. The more interesting question is whether or not the fatty acid and polyketide synthetic pathways share protein components. This question has been addressed almost exclusively in bacteria of the genus Streptomyces due to the remarkable diversity of polyketides made by these bacteria and has focused on type II polyketide synthases which are composed of discrete monofunctional proteins (analogous to type II fatty acid synthesis) rather than the large, highly modular type I polyketide synthases in which interactions are ‘‘hard-wired.’’ The early finding that S. coelicolor contained three different ACP-like proteins, rather than the single protein expected if the fatty acid ACP (AcpP) could also function in polyketide synthesis, suggested that ACP was not a shared component (Revill et al., 1996). AcpP was identified by its constitutive expression, its genomic clustering with genes encoding other fatty acid synthetic proteins and its ability to function in E. coli fatty acid synthesis. In contrast, the remaining two proteins were developmentally regulated and one of these was shown to be required for synthesis of the polyketide, actinorhodin (Revill et al., 1996). Moreover, Streptomyces AcpPs were shown to only very weakly complement mutants lacking the actinorhodin ACP-like protein; only traces of the polyketide were produced (Khosla et al., 1992; Revill et al., 1996). Later work showed

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that AcpP and some PKS ACPs are modified by distinct phosphopantetheinyl transferases (the AcpS reaction of Fig. 17.2), although other PKS ACPs are modified by AcpS (Cox et al., 2002; Lu et al., 2008; Shen et al., 1992a; Walsh et al., 1997). Hence, some AcpS proteins are less specific for their acceptor protein than E. coli AcpS. Moreover, a few phosphopantetheinyl transferases such as B. subtilis Sfp are known to modify a wide diversity of ACP-like proteins (Finking et al., 2002; Mootz et al., 2001). Recently a new approach to test the compatibility of the ACPs of type II FAS and PKS systems has been developed (Worthington et al., 2008). This approach uses modified ACP species that carry a reactive electrophile in place of the thiol of the 40 -PP moiety. Interaction between one of these modified ACPs and a 3-ketoacyl-ACP synthase results in formation of a covalent bound between the ACP prosthetic group and the active site cysteine of a synthase. The systems compared were the E. coli FAS AcpP together with the E. coli FabB and FabF synthases and EncC (the ACP) and EncAB synthase of the Streptomyces maritimus polyketide, enterocin. Two different 40 -PP analogues were tested for each ACP. One analogue mimicked a short chain acyl-ACP whereas the other mimicked a longer chain species. As expected both the E. coli and S. maritimus ACP species reacted with their cognate synthase. Both versions of E. coli AcpP also reacted with S. maritimus EncAB. However, only a partially reciprocal result was seen in that FabF failed to react with either version of EncC, whereas FabB reacted only with EncC carrying the shorter analogue (Worthington et al., 2008). These findings raise the question of whether or not EncC functions in both the FAS and entericin pathways of S. maritimus even though the ACP is encoded in the gene cluster of the latter pathway. However, no S. maritimus genome sequence is available and thus the possibility of an FAS ACP cannot be evaluated at present. The other fatty acid synthetic protein suspected to play a role in polyketide synthesis is FabD, the malonyl-CoA:AcpP transacylase (Fig. 17.3). FabD proteins purified from Streptomyces were shown to be active on both AcpPs and type II PKS ACPs (Florova et al., 2002; Revill et al., 1995; Summers et al., 1995). These enzymes were demonstrated to be fatty acid synthetic proteins by their genomic clustering with other fatty acid genes and in one case the ability to functionally replace the E. coli FabD protein when expressed in E. coli (Summers et al., 1995). There seems only a single discrete malonyl-CoA:AcpP transacylase encoded in the S. coelicolor genome sequence (although domains of similar sequence are found in the modular type I PKS genes) and this is also the case in S. avermitilis. However, the genome of S. griseus subsp. griseus encodes three FabD homologues. Hence, there is a possibility that there may be malonyl-CoA:ACP transacylases targeted to specific PKS ACPs in some Streptomyces but not others. The malonyl-CoA:AcpP transacylase of S. glaucescens has been reported to play an indirect role in providing a building block for synthesis of the polyketide


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tetracenomycin C (Florova et al., 2002). In vivo studies with deuterium labeled acetate indicates that, in direct contrast to fatty acid synthesis, the starter acetate unit is not derived directly from acetyl-CoA as shown by loss of deuterium upon incorporation into the polyketide. The starter unit is thought to be formed indirectly by synthesis of malonyl-CoA from acetyl-CoA followed by transfer of the malonyl moiety to the tetracenomycin ACP (TcmM) (Florova et al., 2002. The resulting malonyl-TcmM would then be decarboxylated to give acetyl-TcmM, the acetate moiety of which is the proposed starter unit (Florova et al., 2002). Although malonyl-ACP decarboxylation is a known side reaction of 3-ketoacylACP synthases, the identity of the enzyme catalyzing decarboxylation is unclear. However, FabH seems a strong possibility (Smirnova and Reynolds, 2001; Wallace et al., 1995). In mycobacteria the FAS II 3-ketoacyl-ACP synthase (KasA) has been found to interact with PpsB and PpsD, two polyketide modules involved in the biosynthesis of the virulence lipid, phthiocerol in E. coli (Kruh et al., 2008). The interaction with PpsB, a four-module PKS protein containing an ACP domain, was examined in detail. When PspB labeled with a longchain fatty acid was incubated with KasA and AcpM, the Mycobacterium AcpP, the acyl chain was transferred to both proteins and could be elongated when other FAS components were present. Hence, transfer of a PKS product to an FAS for further elongation seems possible. The reverse transfer did not occur. The PKS to FAS transfer was proposed to serve to increase the diversity of mycobacterial lipids (Kruh et al., 2008).

10. Methods for Study of Bacterial Fatty Acid Synthesis 10.1. Preparation of the holo and apo forms of E. coli ACP ACP is one of the most abundant proteins of E. coli and can readily be purified from wildtype cells. However, modern day preparations utilize high-level overexpression of the protein. In some cases the acpP gene has been modified by addition of N-terminal or C-terminal tags to aid purification of the encoded fusion protein. However, bacterial ACPs have atypical and highly conserved properties that make purification of the native proteins very facile. First, ACP is very acidic (isoelectric point of 4.1), small and extremely soluble (solutions of >40 mg/ml can be made). This solubility allows bulk purification steps such as precipitation of most other proteins with 50% 2-propanol or saturated ammonium sulfate while ACP remains in solution. We favor the 50% 2-propanol treatment since the 2-propanol does not interfere with ion exchange chromatography, the final step of the purification. The acidic nature of ACP results in very tight binding of

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ACP to strongly basic anion exchanger resins such as Q (quaternary ammonium) or the weakly basic anion exchanger, DEAE (often abbreviated D). We favor the DEAE matrix because it binds fewer contaminating proteins than Q. ACP is bound to the DEAE column at pH 6.1. At this pH the strongly acidic ACP binds tightly whereas most other proteins bind weakly (if at all) because the pH is below their isoelectric points. Second, ACP readily and quantitatively refolds following denaturation with heat, urea, guanidine or trichloroacetic acid (TCA). In the procedures below ACP species are routinely recovered and concentrated by TCA precipitation. The TCA can be removed by dialysis as below by washing the ACP precipitate with a pH 4.1 buffer followed by dissolution in a buffer of higher pH (Cronan and Klages, 1981). Both procedures remove any deoxycholate (used as a coprecipitant) remaining from the precipitation step. The procedure outlined below is modified from those described previously (Keating et al., 1995; Rock and Cronan, 1980, 1981) and applies to Escherichia coli ACP, but may be adaptable to the ACPs of other organisms provided that they have similar sizes and isoelectric points.

10.2. Strains and plasmids used The E. coli K-12 strain used for ACP expression is DK574 (Keating et al., 1995) which is strain SJ16 ( Jackowski and Rock, 1981) containing plasmid pMR19 (Rawlings, 1993) which encodes the structural gene for ACP from E. coli under control of a tac promoter and the Lacl repressor overexpression plasmid pMS421 (Grana et al., 1988). The strain also has a chromosomal lesion in the gene encoding the aspartate decarboxylase, panD. This mutation does not affect growth in rich medium but imposes a requirement for exogenous supplementation with the CoA precursor b-alanine in minimal medium (Cronan, 1982). This permits the isolation of radiolabeled ACP by supplementation with radioactive b-alanine. Plasmid pJT93 carrying the holo-ACP synthase gene acpS under control of a lac promoter was constructed as follows: acpS was amplified from E. coli K-12 genomic DNA. The PCR product was digested with restriction enzymes KpnI and PstI and ligated into the vector pDHC30 (Phillips et al., 2000) digested with the same enzymes. Plasmid pJT94 encoding acpH under lac promoter control was constructed in a similar fashion. The acpH gene was amplified from E. coli K-12 genomic DNA, the PCR product digested with enzymes KpnI and PstI and ligated into pDHC30 digested with the same enzymes.

10.3. Holo-ACP At high levels of expression acyl carrier protein is isolated primarily in the apo-form (Keating et al., 1995). To ensure complete phosphopantetheinylation of the protein, strain DK574 is additionally transformed with the


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plasmid pJT93 expressing the Escherichia coli AcpS 40 -PP transferase under tac promoter control, a system similar to the BAP1 system for the purification of PKS ACPs discussed in Chapter 16 of this volume. Strain DK574 carrying pJT93 is cultured overnight in LB medium supplemented with 15 mg/ml kanamycin, 50 mg/ml streptomycin and 10 mg/ml chloramphenicol. This starter culture is diluted 100-fold into 500 ml of the same medium and incubated with shaking at 37 until it reaches an optical density of 0.8. When ACP labeled in the 40 -PP arm is desired, the following changes are made: the expression strain is cultured overnight in 5 ml M9 medium supplemented with 15 mg/ml kanamycin, 50 mg/ml streptomycin, 10 mg/ ml chloramphenicol, 0.4% glucose, and limiting (0.5 mM) b-alanine in order to reduce the CoA pools. These b-alanine starved cultures are then diluted 100-fold into 100 to 500 ml of M9 medium containing 0.4% glucose and 8 mM [3H] b-alanine (American Radiolabeled Chemicals) and grown to an optical density of 0.8. The expression of ACP and AcpS are simultaneously induced by the addition of 100 mM IPTG followed by incubation for a further 3 to 4 h. The cells are harvested by centrifugation at 17,700g for 10 min and washed in an equal volume of 50 mM Tris-HCl, pH 8.8. The cells are then resuspended in 5 ml AcpS reaction buffer (50 mM Tris-HCl pH 8.8, 10 mM MgCl2, 5 mM dithiothreitol) and lysed by sonication (six 20-s pulses) or by two passages through a French pressure cell. The cell lysate is cleared by centrifugation at 18,000g for 20 min at 4 , 1 mM CoA is added to the supernatant and incubated at 37 for 4 h. The alkaline pH and high concentration of dithiothreitol (DTT) also serve to hydrolyze acyl-ACP thioesters that may be present. At this stage, complete conversion to holoACP is verified by conformationally sensitive gel electrophoresis on a nondenaturing 20% polyacrylamide gel containing 0.5 M urea (Rock et al., 1981b) followed by staining with Coomassie Brilliant Blue 250. The treated cell extract is dialyzed against 50 mM potassium-2(N-morpholino)ethanesulfonic acid (K-MES) (pH 6.1) overnight or subjected to ultrafiltration using a centrifugal filter of MWCO 5000 with three buffer replacements. An equal volume of ice-cold isopropanol is added to the dialyzed extract and incubated with stirring at 4 for 2 to 14 h to allow precipitation of cellular protein. The suspension is cleared by centrifugation at 18,000g for 20 min at 4 . A Vivaspin D Maxi H DEAE centrifuge column (Sartorius) is equilibrated by two washes with 10 ml of 0.5 M K-MES, pH 6.1, and one wash with 10 ml of 25 mM K-MES, pH 6.1 at 2,000g. The supernatant resulting from centrifugation of the isopropanol treatment is applied to the Vivaspin column followed by two washes of the column with 10 ml 25 mM potassium-MES containing 0.3 M LiCl. ACP is eluted from the Vivaspin column with 10 ml of 25-mM K-MES containing 0.5 M LiCl. LiCl is preferred over NaCl as the latter results in the formation of sodium adducts of ACP which may interfere with downstream analyses

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such as mass spectrometry. ACP is precipitated from the eluate by the addition of TCA to a concentration of 5% together with sodium deoxycholate to 0.02% as a coprecipitant (Bensadoun and Weinstein, 1976) followed by incubation at 4 for 30 min. This suspension is centrifuged at 18,000g for 30 min and ACP is resuspended in 1 ml 0.5 M Tris-HCl, pH 8.0. The buffer containing the concentrated protein is replaced by dialysis against 25 mM K-MES, pH 6.1 containing 1 mM DTT and visualized by conformationally sensitive gel electrophoresis (see below) followed by staining with Coomassie Brilliant Blue 250. A 500-ml culture gives about 2 mg of pure ACP (the poorer medium used in preparation of tritium-labeled ACP results in about half this yield).

10.4. Apo-ACP Apo-ACP is often required in the study of 40 -PP transferases or as a substrate for the preparation of acyl-ACP thioesters using a 40 -PP transferase. In this case, the strain DK574 is transformed with the plasmid pJT94 expressing the AcpH ACP hydrolyase AcpH under tac control. Culture of the expression strain and coexpression of ACP and AcpH are carried out as described above. When expression is complete, cells are lysed by sonication or passage through a French pressure cell in 5 ml of AcpH buffer: 50 mM Tris-HCl pH 8.8, 25 mM MgCl2, 1 mM DTT, 0.2 mM MnCl2, and incubated for 4 h at 37 . The purification follows that of holo-ACP.

10.5. Synthesis of Acyl-ACP substrates It is often necessary to synthesize acyl-ACP thioesters as substrates for fatty acid synthesis enzymes. Acyl ACP thioesters may be enzymatically synthesized by three complementary methods. The first utilizes E. coli acyl ACP synthetase (Aas) to form an acyl ACP thioester using holo-ACP and free fatty acid as substrates. The acyl ACP synthetase of E. coli (Ray and Cronan, 1976) is a membrane protein later discovered to have two active sites, an Aas active site and 2-acylglycerophosphoethanolamine acyltransferase active site ( Jackowski et al., 1994). The overall reaction salvages lysophospholipids formed during lipoprotein synthesis by acylation ( Jackowski et al., 1994). The Aas activity of the protein has been extensively used for making acylACPs (Rock et al., 1981b; Shanklin, 2000). More recently, the gene encoding acyl ACP synthetase from Vibrio harveyi was identified and the protein (AasS) expressed and purified ( Jiang et al., 2006). AasS is a soluble cytoplasmic enzyme readily purified in hexahistidine-tagged form and so presents another attractive choice for synthesis of acyl-ACP thioesters. A typical reaction mixture consists of 20 mM ACP, 200 mM fatty acid and 170 nM AasS in a buffer containing 100 mM Tris-HCl (pH 7.8), 10 mM MgCl2, 1 mM TCEP, and 10 mM ATP in a reaction volume of 1 ml.


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The reaction is allowed to proceed for 4 h at 37 and stopped by addition of two volumes of acetone and the proteins allowed to precipitate at –20 overnight. The precipitate is centrifuged at 18,000g for 20 min followed by two washes with three volumes of acetone. The pellet is air dried and resuspended in 20 mM Tris-HCl (pH 7.4) containing 1 mM Tris [2-carboxyethyl] phosphine (TCEP). For most purposes the acetone precipitation product will suffice, although it maybe further purified by chromatography on octyl-sepharose, if the acyl chain is C8 or longer (Rock et al., 1981b). It should be noted that the preferred substrates for AasS are saturated fatty acids of 5 to 16 carbon-atom–chain length (Shen et al., 1992b). For preparation of unsaturated acyl-ACPs E. coli Aas may be required. It should also be noted that AasS is active on only a subset of bacterial ACPs ( Jiang et al., 2006) whereas E. coli Aas has a broader specificity (Shanklin, 2000; Sharma et al., 2005). A second method for acyl-ACP synthesis is the use of a phosphopantetheinyl transferase such as the Sfp of Bacillus subtilis to transfer an acyl-phosphopantetheine group from acyl coenzyme A to apo ACP as discussed in Chapter 10 of volume 458, although the use of CoA thioesters makes this an expensive synthetic route and many acyl-CoAs are not commercially available. The third method is specific chemical acylation of the thiol of the ACP prosthetic group (Cronan and Klages, 1981). In the presence of high concentrations of imidazole at pH 6.5, N-acylimidazoles specifically acylate only the prosthetic group thiol; there is no acylation of the other nucleophilic groups of the protein and the reactions are often quantitative (Cronan and Klages, 1981; Roujeinikova et al., 2002a; Sharma et al., 2005). N-acylimidazoles are readily synthesized and can be used without purification. There, also, is a fourth synthetic method that is very specialized, but that fulfills a shortcoming of the other methods. Synthesis of 3-ketothioesters of ACP is problematical due to the instability of these species. As noted above M. tuberculosis FabH uses saturated acyl-CoAs of chain length C8–C12 as primers in the condensation reaction with malonyl-ACP to give 3-ketoacyl-ACPs two carbon atoms longer than the primer (Choi et al., 2000b). Therefore, M. tuberculosis FabH together with the appropriate acyl-CoA can be added to the reconstructed fatty acid synthesis system in place of the generic FabH and will generate the needed 3-ketoacyl-ACP in situ (Marrakchi et al., 2002). It should be noted that due to the lack of tryptophan and the paucity of other aromatic residues in E. coli ACP (and most other ACPs) standard protein colorimetric assays often seriously underestimate the concentrations of ACP and acyl-ACP solutions. For E. coli ACP a molar extinction coefficient of 1.8 103 at 278 nm has been determined based on amino acid analysis (Rock and Cronan, 1980) and a similar value is found in the older literature (Sauer et al., 1964). This value is accurate unless the ACP sample is grossly contaminated with nucleic acid (this is mainly tRNA that

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elutes from the DEAE matrix with ACP). Given overexpression of ACP this is not a problem, but when ACP was purified from wildtype cells (where the ratio of tRNA to ACP is much higher) this contamination was readily detected by inspection of the UV spectrum of the sample and the tRNA was readily removed by isoelectric point precipitation (Rock and Cronan, 1980). It should also be noted that Coomassie Blue staining of ACP also underestimates (by 5- to10-fold depending on the staining protocol) the amount of ACP present because of a lack of basic groups to bind the dye.

10.6. Purification of fatty acid synthetic enzymes Although each of the E. coli fatty acid synthetic enzymes has been purified from wildtype cells using classical chromatographic procedures, the enzymes are now almost invariably purified as fusion proteins carrying various purification tags, most often N-terminal hexahistidine tags (Heath and Rock, 1995; Hoang et al., 2002). Each of the E. coli and Pseudomonas aeruginosa fatty acid synthetic enzymes remains active with an N-terminal hexahistidine tag (although in most cases the activities of the tagged and native forms have not been compared directly). Similarly the fatty acid synthetic enzymes of a wide variety of organisms retain activity when hexahistidine tagged. Purification of soluble hexahistidine tagged proteins by metal chelate chromatography is routine in most laboratories and will not be described here. Although the tagging approach has also been successful with a wide variety of type II fatty acid synthetic enzymes, occasional problems have arisen. For example, a C-terminally hexahistidine tagged version of the Vibrio cholerae enoyl-ACP reductase, FabV, was several orders of magnitude less active that the native protein (Massengo-Tiasse and Cronan, 2008). In contrast the N-terminal hexahistidine tagged version was much more active than the C-terminally tagged version (although it was somewhat less active than the native protein in vivo). It should be noted that the nickel ions that bleed from the standard metal chelate chromatographic columns and the imidazole used for elution must be efficiently removed from the enzyme preparations. Niþþ can catalyze air oxidation of the thiol (usually DTT) added to insure reduction of ACP and enzyme cysteine residues. Oxidation results in inactive enzymes and inactivates ACP by formation of ACP disulfide-linked dimers. Imidazole can attack acyl thioesters to form acyl-imidazoles ( Jencks and Carriuolo, 1959).

10.7. Direct assays of fatty acid synthetic enzyme activities Although the acyl substrates of fatty acid synthesis are bound to ACP, some of the fatty acid synthetic enzymes show activity with acyl-CoA and acylN-acetylcysteamine substrates (Bloch, 1971; Heath et al., 2000; MassengoTiasse and Cronan, 2008). These latter substrates are generally less active


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than the cognate ACP substrates and have lower VMAX and higher KM values consistent with their serving as model compounds. The simplest of the assays are determination of the 3-ketoacyl-ACP reductase (FabG) and enoyl-ACP reductase (FabI, FabL, FabV) activities because the progress of these reactions is readily monitored by oxidation of the reduced pyridine nucleotide substrate by spectrophotometry. Moreover, these enzymes are often active with commercially available acyl-CoA substrates (Choi et al., 2000b; Heath et al., 2000; Massengo-Tiasse and Cronan, 2008). Two of the enzymes, 3-ketoacyl-ACP synthase III (FabH) and malonylCoA:ACP transacylase (FabD) may be assayed by TCA precipitation assays (Alberts et al., 1974; Han et al., 1998). Short chain acyl-CoAs are soluble in TCA whereas acyl-ACPs are TCA insoluble. Hence, the conversion of a 14C-labeled acyl-CoA substrate to a 14C-labeled, ACP-bound acyl chain results in conversion of a TCA soluble compound to an insoluble compound. Therefore, a simple precipitation step followed by scintillation counting of the washed precipitates gives quantitative data. The 3-hydroxyacyl-ACP dehydratases (FabZ, FabA) may be assayed by hydration of the enoyl-thioester trans-2 double bond, the reversal of the physiological reaction. Loss of the characteristic adsorption of enoyl-thioesters at 263 nm can be followed spectrophotometrically (Bloch, 1971). This assay can be problematical using ACP thioesters and purified enzymes due to protein adsorption at this wavelength and CoA thioesters cannot be used since they absorb strongly at this wavelength. Moreover, assay of crude extracts is precluded by protein and nucleic acid absorption. For these reasons N-acetylcysteamine thioesters are the preferred substrates providing the enzyme will accept these model compounds. The reaction can also be run in the forward reaction by following the increase in absorption at 263 nm. The double-bond isomerization reactions catalyzed by FabA and FabM can likewise be followed by gain or loss of absorption at 263 nm (depending on the substrate used) since cis-3-acylthioesters do not absorb at this wavelength (Bloch, 1971; Marrakchi et al., 2002; Schwab et al., 1985). Determination of the long-chain 3-ketoacyl-ACP synthase (FabF, FabB) activity requires the most complicated of the assays since the enzymes generally require that both substrates (malonyl-ACP and the acyl-ACP) are ACP thioesters, which precludes TCA precipitation assays. The reaction uses malonyl-ACP labeled with 14C either at malonate carbon 2 of or at carbons 1 and 3 plus an unlabeled acyl-ACP. Following incubation with the long-chain 3-ketoacyl-ACP synthase the assay is stopped by addition of buffered sodium borohydride which cleaves the thioester bonds and reduces both the acyl thioester moiety and the newly introduced keto moiety to alcohol moieties (Edwards et al., 1997; Garwin et al., 1980b). The products are radioactive 1,3-acyl diols, a stable product that is readily extracted into toluene, a solvent compatible with direct scintillation counting (the product

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of malonyl-ACP reduction remains in the aqueous phase). It should be noted that both the 3-ketoacyl-ACP synthase and 3-hydroxyacyl-ACP dehydratase reactions can be followed spectrophotometrically using assays coupled to pyridine nucleotide conversion (oxidation or reduction, respectively) by 3-ketoacyl-ACP reductase (FabG) (Birge et al., 1967). However, this enzyme is not commercially available. We have chosen to give literature references on a range of bacteria rather than specific assay conditions since the assays used for the E. coli enzyme may not suffice for enzymes from other bacteria. For example, the FabH of Mycobacterium tuberculosis uses long-chain acyl-CoA substrates rather than acetyl-CoA (Choi et al., 2000b). Long chain acyl-CoAs are insoluble in TCA and thus the E. coli FabH TCA precipitation assay will not work. Instead, the appropriate assay would be the borohydride reduction assay used for FabF and FabB. A second example is the FabH proteins of Bacilli and Streptomyces where short branched-chain acyl-CoAs are used in place of acetyl-CoA (Heath et al., 2000; Li et al., 2005; Smirnova and Reynolds, 2001).

10.8. Reconstituted fatty acid synthesis systems Crude extracts of E. coli synthesize fatty acids when the extracts are supplemented with a thiol reducing agent, NADPH, malonyl-CoA, acetyl-CoA and ACP (Lennarz et al., 1962). An NADPH regenerating system is often included due to the potent NADH oxidase activity of membrane fragments in the crude extract (the pyridine nucleotide transhydrogenase present in the extracts interconverts NADPH and NADH). ACP is added to augment that of the extract and this is probably needed to compensate for the massive dilution relative to in vivo conditions resulting from extract preparation. If the malonyl-CoA is 14C labeled, the reaction can be followed by extraction of the synthesized fatty acids followed by scintillation counting (alternatively the acetyl-CoA can be labeled but incorporation is much less efficient). The reaction can also be followed by use of the conformationally sensitive gel electrophoresis system described below. If an extract deficient in the enzymatic step of interest is available, such extracts can be used to assay activity of an enzyme that catalyzes that step of fatty acid synthesis. Extracts deficient in specific enzymes can be obtained from a mutant strain lacking the enzyme to be assayed (Cronan et al., 1969; Gelmann and Cronan, 1972; Silbert and Vagelos, 1967). Another method is by treatment of the extract with an inhibitor that specifically and irreversibly inactivates the enzyme of interest (followed by removal of excess inhibitor before assay) (McGuire et al., 2001). In both cases the purified enzyme is then added to the deficient extract and its ability to restore fatty acid synthetic ability is measured. Similarly, if one has two mutant strains defective in fatty acid synthesis that have the same phenotype, the two extracts can be mixed. If the mutations are in the same gene (protein)


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normal fatty acid synthesis will not be restored whereas if the strains are mutant in different genes, normal synthesis results. Such ‘‘extract complementation’’ assays were used to show that the E coli unsaturated fatty acid auxotrophs, fabA and fabB, had lesions in two different genes (Cronan et al., 1969) and demonstrated the enoyl-ACP reductase activities of FabV and FabK (Marrakchi et al., 2003; Massengo-Tiasse and Cronan, 2008). Another more flexible approach takes advantage of the ease with which the bacterial fatty acid synthesis proteins can be purified. In this approach (called ‘‘reconstructed fatty acid synthesis’’) each of the proteins is purified as the hexahistidine-tagged species and is then used to assemble a system that synthesizes fatty acids from 14C-labeled malonyl-CoA and acetyl-CoA using NADH and NADPH as cofactors (Heath and Rock, 1995, 1996). The products of the reaction have generally been assayed by the conformationally sensitive gel electrophoresis system described below although the solvent extraction used with extract complementation could also be used. Since the purity of each of component is readily tested by its omission, this system can be used to test functionality of any FAS candidate protein. A criticism of this approach is that in general equal masses of each enzyme were added which may not reflect the in vivo situation. However, this approach has subsequently been validated by mass spectrophotometric analysis of the levels of the fatty acid synthetic proteins in E. coli that indicate that the proteins (except ACP) are each present in a few thousands of molecules per cell (Lu et al., 2007). Since the subunit molecular weights of the E. coli fatty acid synthetic enzymes vary by only about twofold, use of equal masses of the enzymes in the reconstructed fatty acid synthesis system is not a great departure from the in vivo situation. Moreover, the chosen ratio of ACP to the synthetic enzymes also approximates the in vivo ratio. A similar reconstructed system was later assembled from the P. aeruginosa proteins (Hoang et al., 2002).

10.9. Resolution of ACP species by conformationally sensitive gel electrophoresis With the advent of facile synthesis of acyl-ACP species comparisons of the properties of E. coli ACP and long-chain acyl-ACPs showed that acyl-ACPs were more resistant to partial denaturation at elevated pH values (pH 9 to 9.5) than the nonacylated species (Rock and Cronan, 1979). The two protein species coeluted from size exclusion chromatography columns at neutral pH, whereas at pH 9.4 the nonacylated species eluted first. Since the two protein species had essentially the same mass and charge, the earlier elution indicated that the nonacylated species had a larger hydrodynamic radius and that the acyl group acted to stabilize ACP structure toward this pH-induced expansion. A greater degree of separation was obtained by polyacrylamide gel electrophoresis using the standard Tris-based buffer

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system generally used for SDS gels but with the SDS omitted (Cronan, 1982; Rock et al., 1981b). The pH of the separating gel in this system increases from pH 8.8 to about pH 9.5 during electrophoresis (Cronan, 1982), resulting in good resolution of acylated and nonacylated ACP species with the acylated species migrating more rapidly. Moreover, it was found that the acetyl-, butyryl-, hexanoyl-, and octanoyl-ACPs could be resolved with a direct relationship between electrophoretic mobility and acyl chain length (Cronan, 1982). During these investigations it was found that heating of the gel by electrical resistance increased resolution and when commercial temperature-controlled electrophoresis units became available, the system was standardized to 20% polyacrylamide gels run at 37 ( Jackowski and Rock, 1987). However, workers in the plant lipid field found that addition of various concentrations of urea to the gels markedly improved resolution of ACP species (Post-Beittenmiller et al., 1991), such that the separations could be done in a minigel format. Short chain acyl-ACP species were separated at low urea concentrations (0.5 to 1 M) whereas resolution of longer chain (C16–C18) species required higher urea concentrations (up to 5 M). Apo-, holo-, and acyl-ACP may be distinguished from each other by conformationally sensitive gel electrophoresis. This method takes advantage of the differential partial denaturation of ACP species under alkaline conditions in the presence of urea. ACP species carrying a hydrophobic acyl chain are more stable than holo-ACP which in turn is more stable than apoACP and ACP species acylated with hydrophilic acyl groups (e.g., malonylACP) (Rock et al., 1981a). In the gel systems commonly used to resolve ACP species, partially denaturing conditions are maintained by urea and alkaline pH and the mobility of a protein is inversely related to its hydrodynamic radius (all species of a given ACP have essentially the same net charge). Apo- and holo-ACP are best distinguished from each other by using a resolving gel composed of 20% acrylamide, 0.67% N, N0 -methylenebisacrylamide, 375 mM Tris-HCl (pH 8.8), 0.5 M urea, and a stacking gel of 5% acrylamide, 0.17% N, N0 -methylenebisacrylamide, 125 mM Tris-HCl, (pH 6.8). Both gels contain N, N, N0 , N0 -tetramethylethylenediamine, and polymerization is initiated by addition of ammonium persulfate to 0.04%. The minigels are loaded and the samples are subjected to electrophoresis at 120 V for 100 to 110 min in a running buffer containing 25 mM Tris-HCl and 0.19 M glycine. To resolve acyl-ACP thioesters, the concentration of urea is increased to 2.5 M and the gel is run at 120V for 120—150 min. The proteins is then visualized by staining the gel in a solution of 50% methanol, 10% acetic acid and 0.1% Coomassie Brilliant Blue R250 followed by destaining in a solution of 10% methanol and 10% acetic acid. If the protein samples are radioactive, the gel is dried and subjected to autoradiography or phosphorimaging. These gel systems have been very useful in analysis of fatty acid synthesis in vivo and in vitro and allow detection of each of the intermediates of the


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early cycles of fatty acid synthesis. However, they must be augmented with other analytical techniques such at mass spectroscopy. This is because some intermediates are labile (such as 3-ketoacyl-ACPs) and others are difficult to resolve (trans-2-enoic and saturated species of the same chain length). Moreover, the presence of hydrophilic groups such as a 3-hydroxy moiety can be offset by greater hydrophobicity (increased chain length), which can complicate interpretation. The x-ray (Roujeinikova et al., 2002b, 2007) and NMR (Kim and Prestegard, 1989, 1990) structures of ACP provide some rationale for the observed properties of ACP and its acyl thioesters. These data confirmed the original prediction that ACP is structured into a four helix bundle although certain of the helixes are plastic and the protein probably exists in at least two conformations. The acyl chain stabilizes the structure mainly through hydrophobic interactions between the hydrocarbon portion of the acyl chain and residues of the hydrophobic cavity primarily formed by the last three helices of the protein. The prosthetic group also interacts with residues other than that to which it is attached accounting for the separation of the holo and apo forms of ACP. It must be noted that the system described above is for E. coli ACP and its acyl species. Despite the strong sequence conservation, bacterial ACPs differ markedly in their electrophoretic mobilities on these gels (de la Roche et al., 1997; De Lay and Cronan, 2007) and the urea concentrations must be optimized for each ACP. For example separation of the acyl species of the ACP-I and ACP-II of spinach (62% identity) required markedly different urea concentrations (Post-Beittenmiller et al., 1991). It should be noted that in the absence of urea, this gel system resolves ACP from the other proteins present in crude extracts of E. coli (Flugel et al., 2000). This is because other proteins are either too large to enter the gel or lack sufficient negative charge to migrate as rapidly as ACP (or both).

10.10. ACPs behave abnormally in SDS-polyacrylamide gel electrophoresis and gel filtration ACPs generally migrate abnormally in standard SDS-polyacrylamide gel electrophoresis systems because the highly charged nature of the protein results in aberrantly low SDS binding. Indeed, both E. coli and V. harveyi ACPs migrate as 20- to 22-kDa proteins despite having molecular weights of 8.6 kDa (de la Roche et al., 1997; Rock and Cronan, 1979). Long chain acyl-ACPs migrate more rapidly than ACP due to increased binding of SDS to the acyl group which imparts greater negative charge (Rock and Cronan, 1979). However, the thioester linkage of acyl-ACPs can be hydrolyzed during SDS gel electrophoresis. E. coli ACP migrates on gel filtration columns as though a protein of 19 to 20 kDa (Rock and Cronan, 1979). This is not due to dimer formation, but rather to the atypically large hydrodynamic radius of this very flexible and dynamic protein.

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C H A P T E R

E I G H T E E N

Aminocoumarins: Mutasynthesis, Chemoenzymatic Synthesis, and Metabolic Engineering Lutz Heide Contents 1. Introduction 2. Generation of Integrative Cosmids and Heterologous Expression of Novobiocin, Clorobiocin, and Coumermycin A1 Gene Clusters 3. Generation of Single and or Multiple Deletions in the Integrative Cosmids 4. Mutasynthetic Generation of New Aminocoumarin Antibiotics 5. In Vitro Amide Synthetase Assays for the Identification of Suitable Ring A Analogues for Mutasynthesis 6. Use of Various Amide Synthetase Genes for Expanding Mutasynthesis Product Range 7. Generation of Substrates for Chemoenzymatic Synthesis 8. Chemoenzymatic Synthesis of New Clorobiocin Analogues 9. Generation of New Aminocoumarin Antibiotics by Metabolic Engineering 10. Conclusion Acknowledgments References

438 441 442 444 446 447 447 449 451 453 453 454

Abstract The aminocoumarin antibiotics novobiocin, clorobiocin and coumermycin A1 are formed by different Streptomyces strains and are potent inhibitors of bacterial gyrase. Their biosynthetic gene clusters have been analyzed in detail by genetic and biochemical investigations. Heterologous expression of these gene clusters by site-specific integration into the genome of the fully sequenced host Streptomyces coelicolor A3(2) readily results in an accumulation of the antibiotics in yields similar to the wildtype strains.

Pharmazeutische Biologie, Pharmazeutisches Institut, Universita¨t Tu¨bingen, Tu¨bingen, Germany Methods in Enzymology, Volume 459 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)04618-7

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In recent years, the aminocoumarins have developed into a model system for the generation of new antibiotics by genetic methods. Prior to heterologous expression in S. coelicolor, cosmids containing the complete biosynthetic clusters can be manipulated in Escherichia coli by l RED–mediated recombination, creating single or multiple gene replacements or gene deletions. Thereby, mutant strains are generated which are blocked in the synthesis of certain intermediates or in specific tailoring reactions. For instance, mutasynthetic experiments can subsequently be carried out to generate aminocoumarin antibiotics that contain modified acyl moieties attached to the aminocoumarin core, and chemoenzymatic synthesis can be employed for the acylation of the deoxysugar moiety of structural analogues of the aminocoumarin antibiotics. Metabolic engineering—the combination of gene deletions and foreign gene expression via replicative expression vectors—can be used to generate further structural variants of these antibiotics. These methods can be combined, allowing the generation of a wide variety of new compounds. This chapter may provide general pointers for the use of genetic methods in the generation of new antibiotics.

1. Introduction The aminocoumarin antibiotics novobiocin, clorobiocin and coumermycin A1 (Fig. 18.1), produced by different Streptomyces strains, are potent inhibitors of bacterial gyrase (Maxwell and Lawson, 2003). They show excellent antibacterial activity against Gram-positive pathogens, including methicillin-resistant strains. The first aminocoumarin to be discovered was novobiocin, which was licensed as an antibiotic for human therapy under the trade name AlbamycinÒ . Clorobiocin and coumermycin A1 are structurally similar to novobiocin and interact with gyrase in a similar fashion, but with significantly higher affinity than novobiocin. Notably, the latter two antibiotics are also potent inhibitors of topoisomerase IV, a second vital target in the bacterial cell. Simocyclinone D8 (Fig. 18.2) also contains the characteristic aminocoumarin moiety, but otherwise its structure is completely different from the aforementioned compounds. It is also a very potent gyrase inhibitor, but interestingly it acts on a completely different subunit of this heterotetrameric enzyme (Flatman et al., 2005). The structurally complex aminocoumarin antibiotic rubradirin (Kim et al., 2008) does not act on gyrase. No further aminocoumarin antibiotics have been discovered; therefore the chemical diversity of aminocoumarins found in nature appears to be limited. The total chemical synthesis of aminocoumarin antibiotics is possible, but only in a complicated multistep process (Ferroud et al., 1999). In recent years, however, the aminocoumarins have developed into a successful model system for expanding the chemical diversity of natural products by

439

Aminocoumarins

A

Ring C 6´ 6´´ CH3 7´´5´´ H3C O O 4´´ O 3´´

H3C

4´´´ 6´´´ HC 3

5´´´ N H

3´´´ C

OH 4´

5´

O 1´´ H

7´

O C

10 2 1

O 2´ O

8´ Cl

2´´ OH

H N 3´

3

6

Ring B

5

9

8

11 HC 3

4 OH H 3C

Ring A

O

O

O

CH

O

O

N

C

O

OH

O

Ring B

3

C

H

O

CH3

Clorobiocin

O

OH

Ring C

7

Ring A

OH

Novobiocin

NH2

2´´´

MePyC OH

Ring C

CH

HC 3

H3C

H

H 3C

O

O

O

CH

O

3

O

OH O

N

C

O

C

O C

O

3

H3C

MePyC

H N

O

O O

N H

Ring B

OH

C

O

Ring B

CH

O

O

3

Coumermycin A1

N

gyrB

nov E F

3

3

H

B

CH

CH

Ring C

MePyC

H

O

MePyDC

CH3

N

CH3

E F G

GY H I

H

J KL M

I JK L M

N

N 1

N 2

N 3

N 4

N 5

N 6

N 2

N 3

N 4

N 5

N 6

N 7

N 7

R

O P Q R S T U VW hal P

1kb

R

S T U V W Z gyrB

QR

R

parY

clo E

G Y H

I J K L M N1

OP

R R R R R 1 2a 2b 3 4

R 5

R 6

S T U V W gyrB

R

parY

R

cou

Ring A biosynthesis

Terminal pyrrole biosynthesis

3‘‘-O-acylation

Ring B biosynthesis

Central pyrrole biosynthesis

Resistance

Ring C biosynthesis

Amide and glycoside bond formation

Regulation

8‘-methylation/halogenation

4‘‘-O-methylation

Unknown

Figure 18.1 (A) Structures of aminocoumarin antibiotics.‘‘Ring A’’ ¼ 3-dimethylallyl4-hydroxybenzoyl moiety.‘‘Ring B’’ ¼ 3-amino-4,7-dihydroxy-coumarin moiety.‘‘Ring C’’ ¼ deoxysugar moiety. ‘‘MePyC’’ ¼ terminal 5-methyl-pyrrole-2-carboxyl moiety. ‘‘MePyDC’’ ¼central 3-methyl-pyrrole-2,4-dicarboxyl moiety. (B) Organization of the biosynthetic gene clusters for novobiocin (nov), clorobiocin (clo), and coumermycin A1 (cou). The sequences of these gene clusters are accessible in GenBank under accession numbers AF170880, AF329398, and AF235050, respectively.

O O HO

O OH

HO

O Cl

H O N C O

CH3

O O O

CH3

O

CH3 O

OH OH OH

Simocyclinone D8

Figure 18.2 Structure of simocyclinone D8.

OH

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Lutz Heide

genetic methods. The basis for this development was the cloning of the biosynthetic gene clusters of these antibiotics (Fig. 18.1) and the detailed analysis of the function of the genes contained therein for catalysis of biosynthetic reactions, regulation and self-resistance. Figure 18.3 shows as

glucose-1HO phosphate HO

CH2OH

OPO32– dTTP

CloV

L-proline

HO

CloN4

PPi

CloI dTDP

OH

CloQ

C S CloH O

CloR OH

O

3,5-epimerization

H3C

CloR

CloN5

OH

O

O

HO

NH2

OH

FAD CloN3

CloU

FADH2

O2 CO2

HOOC

Ring B

HO

SAM

CO2

OH

??

dTDP

O

OH O2

HOOC

C S CloH O

HO

DMAPP

HOOC

CloJ CloK dTDP

NADH 4-hydroxyphenylpyruvate OH

O

NH2 CloW

C S

HOOC

OH

HO

O

HO

AMP

N H

O

C S CloH O

CloN5

CloN4

OH NAD++H+ CloF

AMP + PPi NH2

HO

H2O

CH3

O

CloH

NH2

CloT

C AMP O

N H

OH

HOOC

ATP

O

HO ATP

COOH

PPi

CH2OH

COOH

N H

prephenate

L-tyrosine HO

OH

COOH

O

NH2

O

O

OH

O ATP

C S

N H

H3C

CloN5

O

O

CloN5

H N

O

O

CH3 dTDP

O

OH

H N

O

O

HO HO

O C OH

Clo-hal

OH

HO

O C

Cl

CloN1

CloN2

OH

OH

CloS 4-ketoreduction H3C

C S O

HO

AMP + PPi

dTDP

O

HO

spontaneous?

N H

CloL

CH3

CloM

CloN5

H O N C

OH

C S O

N H

H3C

CloN1

O

CH3

O

O

HO

O

OH

Cl HO

OH CloP

4´´-O-methyl transfer

CloN6

5´´´-C-methyltransfer

CloN7

3´´-O-acyl transfer 10

5´

6´´ CH3 7´´5´´ H3C

H3C O 4´´

4´´´ 6´´´ H3C 5´´´ N

6´

H N 3´

O

O 2

C

7

1

3

8

9

11

1´´ O

O 3´´

3´´´ C

OH 4´

2´´ OH

H

7´

8´

O 2´ O

6

4 5

OH

Cl

Clorobiocin

O

2´´´

H

Figure 18.3 Biosynthetic pathway of clorobiocin.

OH

Ring A

Aminocoumarins

441

example the biosynthetic pathway of clorobiocin and the catalytic functions of the individual gene products in this process. Details of the functional analysis of the aminocoumarin clusters have been summarized in several reviews (Heide et al., 2008a; Li and Heide, 2004, 2006).

2. Generation of Integrative Cosmids and Heterologous Expression of Novobiocin, Clorobiocin, and Coumermycin A1 Gene Clusters For the generation of new aminocoumarins by genetic methods, heterologous expression of their gene clusters offers two principal advantages: one is the possibility to rapidly manipulate the clusters by l RED– mediated recombination in Escherichia coli, prior to integration into the genome of a Streptomyces strain that is more difficult to manipulate genetically; the second advantage is the possibility to work in a completely sequenced host with the potential to influence the supply of primary metabolic precursors and the expression of regulatory genes outside of the cluster in a systems biology approach (Rokem et al., 2007). The integration functions of phage FC31 (Thorpe et al., 2000) are used for site-specific integration of cosmids containing the entire aminocoumarin biosynthetic gene clusters into the genome of Streptomyces coelicolor M512 (Fig. 18.4). M512 (DredD DactII-ORF4 SCP1– SCP2–) is a derivative of the A3(2) wildtype and lacks the capacity to form three of the genuine antibiotics of S. coelicolor A3(2), that is, undecylprodigiosin, actinorhodin, and methylenomycin (Floriano and Bibb, 1996). The procedure for heterologous expression of the aminocoumarin clusters involves the following steps. 1. Cosmids containing the entire novobiocin or clorobiocin gene cluster in a SuperCosI vector (Stratagene) are generated from the genomic DNA of the respective producer strains using standard techniques (Sambrook and Russell, 2001). The coumermycin A1 gene cluster is relatively large (38.2 kb) and can hardly be obtained on a single cosmid by standard techniques. Therefore, it is obtained by ‘‘stitching’’ of the inserts of two overlapping cosmids, each containing parts of the cluster, using l RED– mediated recombination. This method is described in Vol. 458 Chapter 7 of this series and in Wolpert et al. (2008). 2. The DraI-BsaI fragment of pIJ787 (Eusta´quio et al., 2005), which contains the integrase gene and the attP site of phage FC31 as well as a tetracycline resistance gene and which is flanked by about 100 bp of b-lactamase (bla) sequence on one side and about 300 bp of bla sequence on the other side, is used to replace, via l RED–mediated recombination, the bla gene in the SuperCos1 backbone of the cosmids containing the aminocoumarin gene clusters, thereby generating the desired integrative cosmids.

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Lutz Heide

A

912 bp B

13.7 kb

B

7274 bp

B B B B 3815 bp 5583 bp 4443 bp

ORF A B C D E F G H I J K L M N OP Q R S T U VW gyrBR ORF 13 7 6 21 20

Novobiocin cluster in integrative cosmid

T7 B

∼ 8.5 kb

B ∼ 10 kb

Unknown sequence

T3

neo int

B S. coelicolor M512 chromosome

attP

tet

attB

Site-specific integration

B

attL

Integration mutant

int

neo

B

B

tet

attR B

Nov-cluster 13 kb

∼ 5.2 kb

Figure 18.4 Schematic representation of site-specific integration of the novobiocin gene cluster into the genome of Streptomyces coelicolor M512. B ¼ BglII restriction sites.

3. Because of the potent methylation-dependent restriction system of S. coelicolor, cosmid DNA has to be passed through E. coli ET12567 as a nonmethylating host (MacNeil et al., 1992) prior to introduction into the Streptomyces host. 4. The integrative cosmids, still carrying the kanamycin resistance gene neo, are then introduced into S. coelicolor M512 via polyethylene glycol– mediated protoplast transformation (Kieser et al., 2000). Kanamycinresistant clones are checked for site-specific integration into the genome by Southern blot analysis. 5. Cultivation of the integration mutants in the production media for novobiocin (Kominek and Sebek, 1974), clorobiocin (see Section 4 below) and coumermycin A1 (Wolpert et al., 2008) results in the accumulation of the antibiotics in yields of 20 to 42 mg/l, 18 to 34 mg/l, and 7 mg/l, respectively.

3. Generation of Single and or Multiple Deletions in the Integrative Cosmids Single gene replacements or gene deletions within the integrative cosmids can be carried out by l RED–mediated recombination, for example using the apramycin resistance cassette of plasmid pIJ773, which is

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flanked by FLP recombinase recognition targets (FRT) (Gust et al., 2004). After gene replacement, this cassette can be conveniently removed by action of FLP recombinase, leading to excision of the DNA region between the FRT sites and leaving an 81-bp ‘‘scar’’ sequence. However, the presence of this scar sequence makes further knockouts in the same cosmid difficult, because it represents a functional FRT site as well as a target for l RED– mediated recombination. Therefore, another procedure has been developed which opens the possibility for multiple deletions within the cosmid insert. An apramycin-resistance cassette is generated by PCR using primers with either an XbaI or a SpeI recognition site between the apramycin resistance marker and the 39-bp flanking sequence for l RED–mediated recombination. XbaI and SpeI sites are rare in the GC-rich Streptomyces genome. After gene replacement, this cassette can be removed by digestion with XbaI and SpeI and re-ligation of the resulting compatible ends. This procedure leaves a minimal in-frame ‘‘scar’’ of 18 bp which is not recognized by XbaI or SpeI and which does not interfere with further gene deletions or replacements. This procedure is therefore the method of choice for modification of the aminocoumarin gene clusters in E. coli prior to their heterologous expression in S. coelicolor (see also Vol. 458 Chapter 7 of this series). It involves the following steps: 1. A fragment of 97 bp is amplified from pIJ773 (Gust et al., 2003) using primers FRT_P01f (5-CTG CAG GAA TTC GAT ATT CCG GGG ATC TCT AGA TCT-3) (the EcoRI, XbaI, and BglII restriction sites are underlined) and FRT_P01r (5-TGG CGG GGA TAT CGA AGT TCC-3) (the EcoRV restriction site is underlined). After digestion with EcoRI and EcoRV, this fragment is ligated into the same sites of pBluescript SK(-) (Stratagene) to give pUG017. 2. A second fragment of about 1 kb containing the apramycin resistance gene aac(3)IV is amplified from pIJ773 using primers apra_P03f (5-GGG GAT GAT ATC TTT ATC ACC ACC GAC TAT TTG-3) (the EcoRV restriction site is underlined) and apra_P02r (5-TCG ATA AGC TTG ATG ACT AGT CTG GAG CTG GAG CTG CTT CGA-3) (the HindIII and SpeI restriction sites are underlined). After digestion with EcoRV and HindIII, this fragment is ligated into the same sites of pUG017 to give pUG019. 3. The apramycin resistance cassette (approximately 1 kb) is excised from pUG019 by digestion with EcoRI and HindIII and amplified by PCR using the forward primer 5-(N)39 ATT CCG GGG ATC TCT AGA TCT-3 and the reverse primer 5-(N)39 ACT AGT CTG GAG CTG CTT C-3 ((N)39 represents 39-nucleotide extensions for l RED– mediated recombination, homologous to the regions upstream and downstream of the DNA fragment to be deleted; XbaI and SpeI restriction sites are underlined). Amplification is performed using the Expand High Fidelity PCR system (Roche Molecular Biochemicals) according

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to the manufacturer’s instructions, with annealing temperatures of 45 and 48 , respectively. 4. The PCR product is used for gene replacement in cosmids via l RED– mediated recombination (Gust et al., 2003, 2004). 5. For excision of the resistance cassette after gene replacement, cosmid DNA is isolated from E. coli ET12567 and digested with XbaI and SpeI, and 100 ng of DNA is re-ligated overnight at 4 . E. coli XL1-Blue MRF0 cells are transformed with the ligation reaction. Apramycin-sensitive kanamycin-resistant clones are analyzed by restriction enzyme digestion and gel electrophoresis. 6. Subsequent gene deletions within the same cosmid follow the identical procedure.

4. Mutasynthetic Generation of New Aminocoumarin Antibiotics Mutasynthesis—the feeding of synthetic precursor analogues to mutants of microbial producer strains of natural products—is an important and powerful tool for drug discovery and lead optimization. In the aminocoumarin group of antibiotics, it is especially useful to replace the natural 3-dimethylallyl-4-hydroxybenzoyl moiety (termed ‘‘Ring A’’) of novobiocin and clorobiocin by synthetic analogues thereof. In clorobiocin biosynthesis, Ring A is formed via catalysis by the gene products of cloF, cloQ, and cloR (Fig. 18.3). When the prenyltransferase gene cloQ is inactivated, biosynthesis of this moiety is abolished. Since Ring A is the starter unit of the biosynthesis, no aminocoumarin antibiotics are accumulated by such a mutant. If, however, synthetic analogues of Ring A are added to the culture medium, they may be incorporated instead of Ring A and lead to the formation of new aminocoumarin antibiotics. Inactivation of cloQ is carried out by an in-frame deletion on an integrative cosmid containing the entire clorobiocin cluster, using the procedure described above. The entire mutasynthesis procedure involves the following steps: 1. The apramycin-resistance cassette for replacement of cloQ is generated by PCR amplification using pUG019 (see Section 3, Steps 1 and 2) as template with the primers cloQ_for (50-GGCGCGCCCATTGCT CACCGTCTTACCGACACCGTCCTTATTCCGGGGATCTC TAGATC-30) and cloQ_rev (50-CCCATGGTCGATTCCGTGT GTTGGTGAAGTGCGCGCAGACTAGTCTGGAGCTGCTTC30). Bold letters represent the homologous extensions to the DNA regions immediately upstream and downstream of cloQ. Underlined letters indicate the XbaI and SpeI restriction sites. PCR amplification is performed in a

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2.

3.

4.

5.

6.

7. 8. 9.

10.

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50-ml volume with 50 ng template, 0.2 mM dNTPs, 50 pmol of each primer, and 5% (v/v) DMSO with the Expand High Fidelity PCR system (Roche Molecular Biochemicals): denaturation is at 94 C for 2 min, and then 10 cycles with denaturation at 94 for 45 s, annealing at 50 for 45 s, and elongation at 72 for 90 s, followed by 15 cycles with annealing at 55 for 45 s, and the last elongation step at 72 for 5 min. The PCR product is introduced by electroporation into E. coli BW25113/pIJ790 harboring a cosmid containing the entire clorobiocin cluster and the integration functions of phage FC31 (see Section 2). The resulting modified cosmid is isolated, introduced by transformation into the nonmethylating strain E. coli ET12567, reisolated, and digested with XbaI and SpeI to remove the apramycin-resistance cassette. Religation overnight at 4 gives the desired cosmid with the cloQ-defective clorobiocin cluster. The cloQ-defective cosmid isolated from E. coli ET12567 is introduced into S. coelicolor M512 by PEG-mediated protoplast transformation (Kieser et al., 2000). Clones resistant to kanamycin are selected, and integration of the cloQ-defective clorobiocin cluster into the genome is confirmed by Southern blot analysis. For mutasynthetic production of new aminocoumarins, the cloQ-defective heterologous expression strain is first precultured in YMG liquid medium (pH 7.3) containing 1% malt extract, 0.4% yeast extract, 0.4% glucose, and 25 mg/ml kanamycin at 30 and 180 rpm for 2 to 3 days. One milliliter of this YMG preculture is then used to inoculate 50 ml corn starch medium (pH 7.0) (1% corn starch [Becton-Dickinson, Heidelberg, Germany], 1% peptone [Roth, Karlsruhe, Germany], 0.5% meat extract [Merck, Darmstadt, Germany] and 25 mg/ml kanamycin), followed by cultivation for 2 days at 33 and 210 rpm. Five milliliters of this corn starch preculture are inoculated into 80 ml of production medium, prepared from 4.8% distillers’ solubles (SigmaAldrich, Deisenhofen, Germany), 3.7% glucose, 0.0024%CoCl2 6 H2O (at this point, the pH of the mixture is adjusted to 7.8), 0.6% CaCO3, 0.2% (NH4)2SO4, and 25 mg/ml kanamycin. At the time of inoculation of the production medium, 1 mg of the respective ring A analogue, dissolved in 200 ml ethanol, is added to 80 ml of culture medium. Cultivation is carried out in 500-ml baffled flasks for 7 to 10 days at 33 and 210 rpm. For preparative isolation of the products, a total culture volume of 800 ml is pooled and acidified with HCl to pH 4. Lipophilic components are removed by extraction with petrol ether, and subsequently the aqueous phase is extracted twice with an equal volume of ethyl acetate. The residue of the ethyl acetate extract after evaporation of the solvent is dissolved in 3 ml methanol and applied to a column (100 2.6 cm) of

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Sephadex LH 20 (Amersham Biosciences) which is eluted with methanol. 11. The fractions after separation on Sephadex LH 20 are analyzed by HPLC using an analytical RP18 column and the solvent gradient mentioned below. Fractions containing novclobiocins are pooled and further purified on a preparative HPLC column (Multosphere 120 RP18-5, 250 20 mm, C&S Chromatographie Service, Du¨ren, Germany) using a linear gradient from 68 % to 100% MeOH in 0.2% aqueous acetic acid for 37 min and a flow rate of 2.5 ml/min. Detection is carried out by UV absorption at 340 nm. Examples of new aminocoumarins accessible by this procedure are described in Galm et al. (2004).

5. In Vitro Amide Synthetase Assays for the Identification of Suitable Ring A Analogues for Mutasynthesis The most important limiting step for the generation of new aminocoumarin antibiotics by the mutasynthetic procedure described above is the substrate tolerance of the amide synthetase CloL, which attaches the externally added ring A analogue via amide bond formation to the 3-amino group of the aminocoumarin moiety (see Fig. 18.3). In vitro enzyme assays with CloL can therefore be used to predict, with minimal requirement for time and material and with significant accuracy, whether or not a given ring A analogue will yield a new aminocoumarin antibiotic in mutasynthesis (Galm et al., 2004). Moreover, the range of aminocoumarins that can be produced by mutasynthesis can be significantly expanded if other amide synthetases, such as that of coumermycin A1 biosynthesis (CouL) or that of simocyclinone D8 biosynthesis (SimL) (Luft et al., 2005), are also investigated for their substrate tolerance for the respective ring A analogue. If found suitable, these amide synthetases are utilized in the mutasynthesis procedure (see Section 7). 1. The three amide synthetase genes cloL, couL, and siml are cloned into the expression vector pQE70 (Qiagen) according to the manufacturer’s protocol. 2. The C-terminally His6-tagged fusion proteins NovL, CouL, and SimL are expressed in E. coli XL1 Blue MRF0 (Stratagene) and purified by affinity chromatography on Ni-NTA agarose (Qiagen) according to the manufacturer’s protocol. 3. The assay for amide synthetase activity contains 1mM of the ring A analogue, 1 mM 3-amino-4,7-dihydroxy-8-methyl-coumarin, 5 mM ATP, 5 mM MnCl2, 50 mM Tris-HCl (pH 8.0), and 0.5 to 2 mg of the

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respective protein in a final volume of 100 ml. The reaction is carried out for 30 min at 30 and stopped by addition of 5 ml 1.5 M trichloroacetic acid. The assay mixture is extracted with 1 ml of ethyl acetate, the organic phase is evaporated, and the residue is dissolved in H2O/ methanol (50:50, v/v). HPLC analysis is carried out using a Multosphere RP18-5 column (250 4 mm; 5-mm particle size) with a linear gradient from 60 to 100% methanol in 1% aqueous formic acid and detection at 305 nm.

6. Use of Various Amide Synthetase Genes for Expanding Mutasynthesis Product Range Compounds which are found, in the enzyme assay described in Section 5, to be poorly accepted by CloL but readily by CouL (e.g., 3,5dimethyl-4-hydroxybenzoic acid), or by SimL (e.g., benzoic acid), can be successfully used in mutasynthesis if the suitable amide synthetase is cloned into an expression vector and introduced by transformation into the heterologous expression strain containing the cloQ-defective clorobiocin cluster. 1. The amide synthase genes couL and siml are cloned into the E. coliStreptomyces shuttle vector pUWL201, containing the ermE* promoter for strong constitutive expression (Doumith et al., 2000). 2. The couL or siml expression plasmids are introduced by transformation into S. coelicolor M512, harboring the cloQ-defective clorobiocin cluster, by PEG-mediated protoplast transformation (Kieser et al., 2000). 3. Subsequently, the mutasynthetic experiment is carried out as described above (Section 4, Steps 4 to 11), including 25 mg/ml thiostrepton in the cultivation media for selection of cells containing the couL or siml expression plasmids. Examples of new aminocoumarins accessible by expression of couL and siml in the cloQ-defective heterologous expression strain are described in Anderle et al. (2007b).

7. Generation of Substrates for Chemoenzymatic Synthesis The structural skeleton of novobiocin and clorobiocin consists of the 3-dimethylallyl-4-hydroxybenzoyl moiety (ring A), the 3-amino-4,7-dihydroxycoumarin moiety (ring B), substituted at position 8 with a methyl group or a chlorine atom, and the 5-methyl-L-rhamnose moiety (ring C).

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In two final tailoring reactions of the biosynthesis, the deoxysugar moiety (ring C) is decorated with two further substituents (see Fig. 18.3): The 4-OH group of the deoxysugar is methylated by the SAM-dependent methyltransferases NovP or CloP, respectively; and the 3-OH group of the deoxysugar is acylated. In novobiocin biosynthesis, a carbamoyl group is attached via catalysis by the carbamoyl transferase NovN. In clorobiocin biosynthesis, the respective acyl group is a 5-methyl-pryrrol-2-carboxylmoiety, derived from proline via catalysis by CloN3, CloN4 and CloN5 and transferred from the acyl carrier protein CloN5 to the deoxysugar in a twostep process involving two acyl transferases, CloN2 and CloN7, and a further acyl carrier protein, CloN1 (see Fig. 18.3) (Garneau-Tsodikova et al., 2006). Methylation at position 5 of the pyrrole-2-carboxyl moiety takes place prior to attachment of this moiety to the deoxysugar (Anderle et al., 2007a). For the generation of new aminocoumarins by chemoenzymatic synthesis (and by organic-synthetic procedures), it is of interest to generate novobiocin or clorobiocin analogues which lack either the acyl moiety at position 3 of the deoxysugar, or the methyl group at position 4 of the deoxysugar, or both. This can conveniently be achieved by inactivation of the responsible genes on cosmids containing the respective gene cluster and by heterologous expression of the resulting cosmids, using the methods described above. As example, the generation of a clorobiocin analogue (termed novclobiocin 104) lacking the acyl moiety at position 3 of the deoxysugar is described here. 1. The apramycin resistance gene aac(3)IV is amplified from plasmid pUG019 (see Section 3, Steps 1 and 2) by PCR, with use of primers with 39-bp extensions homologous to the regions upstream and downstream of cloN7, that is, primer cloN7-f (50 -GGCAGACTCCCCAACAGCA GAGAGGACCAACTGAGCATGATTCCGGGGATCTCTAGA TC-30 ) and primer cloN7-r (50 -AGTGTGCGTGGTGCGCCAGCA CTCCGACAAGCACCGTTAACTAGTCTGGAGCTGCTTC-30 ). Bold letters represent the homologous extensions to the DNA regions immediately upstream and downstream of cloN7, including the putative start and stop codons of cloN7. PCR amplification is performed in a 50-ml volume with template (50 ng), dNTPs (0.2 mM), each primer (50 pmol) and DMSO (5% v/v), with use of the Expand High Fidelity PCR system (Roche Molecular Biochemicals): denaturation is at 94 for 2 min, then 10 cycles with denaturation at 94 for 45 s, annealing at 45 for 45 s and extension at 72 for 90 s, followed by 15 cycles with annealing at 48 , and the last elongation step at 72 for 5 min. 2. The PCR product is introduced by electroporation into E. coli BW25113/pIJ790 harboring an integrative cosmid with the entire clorobiocin cluster. l RED–mediated recombination results in the

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replacement of the entire open reading frame of cloN7 in the clorobiocin cluster with the apramycin resistance cassette, leaving only the start and the stop codons of cloN7 intact. The modified cosmid is isolated and analysed by restriction enzyme digestion. 3. Since cloN7 is situated at the end of a putative transcription unit, removal of the apramycin resistance cassette to avoid polar effects is unnecessary in this case. Note that in a similar experiment with cloN1, which is situated at the start of a putative transcription unit, it is advisable to remove the resistance cassette by digestion with XbaI and SpeI as described in Section 3, Step 5 (Freitag et al., 2005b). 4. The cloN7-deficient cosmid is then passed through the nonmethylating host E. coli ET12567 and introduced into S. coelicolor M512 as described in Section 2. Site-specific integration into the genome is confirmed by Southern blot analysis. 5. Cultivation of the resulting strain and preparative isolation of the desired product, novclobiocin 104, is carried out as described above for the mutasynthesis experiment (Section 4, Steps 4 to 11), with omission of Step 7, that is, without the external addition of a ring A analogue. A similar procedure can be used, for instance, in generating a strain defective in the methyltransferase gene cloP, which produces clorobiocin analogues lacking the methyl group at 4-OH of the deoxysugar (Freitag et al., 2005a).

8. Chemoenzymatic Synthesis of New Clorobiocin Analogues Most of the enzymes involved in the biosynthesis of the aminocoumarin antibiotics have meanwhile been overexpressed and purified, and are available for the chemoenzymatic generation of new aminocoumarins in vitro. The production of a hybrid antibiotic, containing a carbamoyl group typical of novobiocin in the deoxysugar, and a chlorine atom typical of clorobiocin in the aminocoumarin moiety, is given as an example. 1. The substrate novclobiocin 104, carrying a chlorine atom at position 8 of the aminocoumarin moiety and lacking the acyl group a 3-OH of the deoxysugar, is generated as described above (Section 8). 2. The carbamoyl transferase NovN is expressed and purified in the following way: novN is amplified from a cosmid containing the novobiocin biosynthetic gene cluster using the GC-RICH PCR system (Roche, Mannheim, Germany) and the primer pair novN-BclI (50 - GTG CTC GCT GAT CAG AAC GAC ATG -30 ) and novN-HindIII (50 - AAG GGA AGC TTT ACG GCC GCG AC -30 ). Introduced restriction sites

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are underlined. The DNA fragment is directly ligated into the linearized vector pGEM-T (Promega), resulting in pXHNET. novN is released from pXHNET by digestion with BclI and HindIII and ligated into pRSET-B (Invitrogen), which has been digested with BamHI and HindIII, to give pXHNER. To obtain pXHNEG, pXHNER is fused with the vector pGM9 (Muth et al., 1989) via the HindIII restriction site. pXHNEG contains a kanamycin resistance gene. After transforming S. lividans T7 protoplasts with plasmid pXHNEG, cells are regenerated on R2YE plates and kanamycin- and thiostrepton-resistant colonies are selected. They are grown in YEME medium with kanamycin (10 mg/ml) at 28 for 48 h, and then 1 ml culture is transferred to 100-ml YEMEmedium with kanamycin (10 mg/ml) and thiostrepton (25 mg/ml), and cultured for further 24 h. The cells are harvested by centrifugation (10 min at 5000g) and incubated in ice-cold lysis buffer (50 mM NaH2PO4, pH 8.0, 300 mM NaCl, 15 mM imidazole, 0.2% triton 100 and 6 mg/ml lysozyme) for 30 min. The cell suspension is sonicated for 8 min (Branson Sonifier 250) and the cell debris is removed by centrifugation (30 min at 15,000g). The protein is purified from the soluble cell extract by metal affinity chromatography using Ni-nitrilotriacetic acid (Ni-NTA) resin (Qiagen) according to the manufacturer’s instructions. Subsequently, the Ni-NTA resin eluate is applied to a NAP 10 column (Amersham Biosciences) and eluted with 0.2 M Tris-HCl (pH 8.0). 3. For chemoenzymatic generation of the desired new aminocoumarin antibiotic, the carbamoyl transferase assay mixture contains 0.2 M TrisHCl (pH 8.0), 1 mM novclobiocin 104, 5 mM carbamoyl phosphate (as Liþ salt), 2 mM ATP, 2 mM MgCl2 and purified NovN (0.4 mM ) in a final volume of 5 ml. The reaction is carried out overnight at 30 and terminated by the addition of 250 ml 1.5 M trichloroacetic acid. The assay products are extracted twice with an equal volume of ethyl acetate. After evaporation of the organic solvent, the residue is dissolved in methanol, and the products are isolated by HPLC using the method described for the mutasynthesis experiments (see Section 4, Step 11). (Note that column chromatography on Sephadex LH 20 is unnecessary in this case, as the extract from the enzyme assay contains far less impurities than extracts from microbial cultures.) Using other substrates than novclobiocin 104, hybrid antibiotics with different structures can be obtained by this chemoenzymatic method: the preparative generation of 10 different aminocoumarin antibiotics is described in Xu et al. (2004). Furthermore, it is possible to generate variants of coumermycin A1 by tandem action of four biosynthetic enzymes, CouL, CouM, CouP, and NovN, using 3-methyl-pyrrole-2,4-dicarboxylic acid, the aminocoumarin moiety, dTDP-5-methyl-rhamnose and carbamoyl phosphate as substrates and ATP and SAM as cofactors (Freel Meyers

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et al., 2004). Additional chemoenzymatic approaches for generating new aminocoumarin antibiotics are described in Fridman et al. (2007) and Williams et al. (2008).

9. Generation of New Aminocoumarin Antibiotics by Metabolic Engineering The availability of biosynthetic gene clusters for aminocoumarin antibiotics of different structures and detailed knowledge of the genes contained in these clusters, form an excellent basis for the generation of new aminocoumarin antibiotics by rational metabolic engineering via combining gene inactivations and gene expressions. An example of the systematic generation of a series of new aminocoumarin antibiotics with different substituents at position 80 of the aminocoumarin moiety and position 3" of the deoxysugar is given in Fig. 18.5. These experiments can be carried out with the methods described above for generation of single and/or multiple deletions within integrative cosmids (see Section 3), for heterologous expression of the resulting modified biosynthetic gene clusters (see Section 2), and for the expression of additional biosynthetic genes using replicative expression vectors (see Section 6).

Novobiocin 3² 8¢ Carbamoyl CH3

Clorobiocin 3² 8¢ MePyC Cl

– methyltransferase novO Novclobiocin 117 3² 8¢ Carbamoyl H

Novclobiocin 104 3² 8¢ Cl H

– halogenase clo-hal

Novclobiocin 101 3² 8¢ MePyC H

+ halogenase clo-hal Novclobiocin 114 3² 8¢ Carbamoyl Cl

– acyl transferase cloN2

+ methyltransferase novO

Novclobiocin 102 3² 8¢ MePyC CH3

– acyl transferase cloN2

Novclobiocin 107 3² 8¢ H H

Side product Novclobiocin 103 3² 8¢ H CH3

Figure 18.5 Scheme showing routes for the generation of novobiocin and clorobiocin analogues by metabolic engineering of the producer strains. MePyC ¼ 5-Methylpyrrole-2-carboxyl moiety.

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The following section therefore gives an overview of the steps involved, without describing the methods again in detail. Novobiocin can be produced in the heterologous host S. coelicolor M512, after stable integration of the complete novobiocin biosynthetic gene cluster into its genome (see Section 2). The productivity of the heterologous producer (31 mg/l) is similar to that of the natural producer, Streptomyces spheroides (35 mg/l). Inactivation of the methyltransferase gene novO, responsible for 80 -methylation of the aminocoumarin ring, in the novobiocin cluster, and heterologous expression of the resulting cosmid readily yields a strain which accumulates novclobiocin 117 (30 mg/l), the novobiocin analogue lacking the 80 -methyl group. When the halogenase gene clo-hal, responsible for 80 -halogenation of the aminocoumarin ring in clorobiocin biosynthesis, is cloned into the replicative expression plasmid pUWL201 and expressed in the novO-defective strain, the hybrid antibiotic novclobiocin 114 is produced, which carries the 3"-O-carbamoyl group typical of novobiocin, and the 80 -chlorine typical of clorobiocin (yield 14 mg/l). The clorobiocin producer Streptomyces roseochromogenes can be manipulated genetically with relative ease. The halogenase gene clo-hal can be inactivated in this strain by an in-frame deletion (Eusta´quio et al., 2003), which results in the production of the 80 -unsubstituted compound novclobiocin 101 (40 mg/l). Subsequently, the methyltransferase gene novO can be expressed in this strain using the replicative expression plasmid pUWL201. This leads to the formation of the hybrid antibiotic novclobiocin 102, which carries the 5-methyl-pyrrole-2-carboxyl group typical of clorobiocin at position 3", and the methyl group typical of novobiocin at position 80 (yield 43 mg/l). Besides novclobiocin 102, a side product can be isolated, carrying the methyl group at 80 , but no substituent at 3"-OH (i.e., novclobiocin 103 [11 mg/l]). This makes a separate experiment for the generation of novclobiocin 103 such as via inactivation of the carbamoyl transferase gene novN in the novobiocin cluster, unnecessary. The attachment of the 5-methyl-pyrrole-2-carboxyl (MePyC) moiety to the 3"-OH of noviose requires, besides other proteins, the acyltransferase CloN2. When the cloN2 gene in the clorobiocin producer S. roseochromogenes is inactivated, the resulting mutant accumulates novclobiocin 104, lacking the MePyC unit. This compound is produced in a yield of 85 mg/l, surpassing the productivity of clorobiocin in the wildtype. The gene cloN2 can also be inactivated in the clo-hal defective mutant described above. The resulting clo-hal – cloN2 – double mutant produces novclobiocin 107, which carries no substituent at 80 and at 3"-OH. Therefore, all compounds depicted in Fig. 18.5 can readily be obtained from genetically engineered producer strains. Metabolic engineering can also be carried out with genes from biosynthetic gene clusters outside of the aminocoumarin family. This can be demonstrated by inactivation of the methyltransferase gene cloN6, which

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is responsible for attachment of a methyl group at position 5 of the pyrrole2-carboxyl moiety of clorobiocin (Fig. 18.3), on an integrative cosmid containing the entire clorobiocin cluster. The cloN6-defective cluster is expressed in S. coelicolor M512 (using genomic integration) together with the halogenase gene hrmQ (using the replicative expression plasmid pUWL201). hrmQ is obtained from the biosynthetic gene cluster of hormaomycin (Reinscheid et al., 2005) and is presumed to be responsible for the attachment of a chlorine atom at position 5 of the pyrrole-2-carboxyl moiety of hormaomycin, which is structurally completely different from clorobiocin. This experiment results in the formation of a new clorobiocin derivative, carrying a chlorine atom rather than a methyl group at position 5 of the pyrrole-2-carboxyl moiety (Heide et al., 2008b).

10. Conclusion The family of aminocoumarin antibiotics provides an example for the generation of new antibiotics by genetic methods. The available detailed knowledge of the biosynthetic pathways and of the functions of the genes contained in the biosynthetic clusters facilitates such experiments. Limitations posed by the substrate specificity of biosynthetic enzymes can be overcome by expression of heterologous enzymes (e.g., amide synthetases) with appropriate specificities. Heterologous expression of the entire biosynthetic gene clusters in suitable expression hosts significantly accelerates the engineering procedures and offers excellent prospects to increase production rates by upregulating precursor supply and influencing regulatory cascades. These experiments exemplify that drug structures can be systematically altered by genetic engineering of the producer strains, avoiding the need for multistep organic synthesis. Such genetic methods are likely to become a cornerstone of programmes for drug discovery and development from microorganisms in future.

ACKNOWLEDGMENTS The author gratefully acknowledges the invaluable contributions of many colleagues, coworkers, and collaboration partners in the development of the methods described here, especially of S.-M. Li and B. Gust, as well as of C. Anderle, A. Eusta´quio, A. Freitag, U. Galm, H. Xu, L. Westrich, B. Kammerer, and many others. The work was financially supported from the Deutsche Forschungsgemeinschaft and from the European Community (IP 005244 ActinoGEN).

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REFERENCES Anderle, C., Alt, S., Gulder, T., Bringmann, G., Kammerer, B., Gust, B., and Heide, L. (2007a). Biosynthesis of clorobiocin: Investigation of the transfer and methylation of the pyrrolyl-2-carboxyl moiety. Arch. Microbiol. 187, 227–237. Anderle, C., Hennig, S., Kammerer, B., Li, S. M., Wessjohann, L., Gust, B., and Heide, L. (2007b). Improved mutasynthetic approaches for the production of modified aminocoumarin antibiotics. Chem. Biol. 14, 955–967. Doumith, M., Weingarten, P., Wehmeier, U. F., Salah-Bey, K., Benhamou, B., Capdevila, C., Michel, J. M., Piepersberg, W., and Raynal, M. C. (2000). Analysis of genes involved in 6-deoxyhexose biosynthesis and transfer in Saccharopolyspora erythraea. Mol. Gen. Genet. 264, 477–485. Eusta´quio, A. S., Gust, B., Galm, U., Li, S. M., Chater, K. F., and Heide, L. (2005). Heterologous expression of novobiocin and clorobiocin biosynthetic gene clusters. Appl. Environ. Microbiol. 71, 2452–2459. Eusta´quio, A. S., Gust, B., Luft, T., Li, S. M., Chater, K. F., and Heide, L. (2003). Clorobiocin biosynthesis in Streptomyces. Identification of the halogenase and generation of structural analogs. Chem. Biol. 10, 279–288. Ferroud, D., Collard, J., Klich, M., Dupuis-Hamelin, C., Mauvais, P., Lassaigne, P., Bonnefoy, A., and Musicki, B. (1999). Synthesis and biological evaluation of coumarincarboxylic acids as inhibitors of gyrase B. L-rhamnose as an effective substitute for L-noviose. Bioorg. Med. Chem. Lett. 9, 2881–2886. Flatman, R. H., Howells, A. J., Heide, L., Fiedler, H. P., and Maxwell, A. (2005). Simocyclinone D8, an inhibitor of DNA gyrase with a novel mode of action. Antimicrob. Agents Chemother. 49, 1093–1100. Floriano, B., and Bibb, M. (1996). afsR is a pleiotropic but conditionally required regulatory gene for antibiotic production in Streptomyces coelicolor A3(2). Mol. Microbiol. 21, 385–396. Freel Meyers, C. L., Oberthur, M., Heide, L., Kahne, D., and Walsh, C. T. (2004). Assembly of dimeric variants of coumermycins by tandem action of the four biosynthetic enzymes CouL, CouM, CouP, and NovN. Biochemistry 43, 15022–15036. Freitag, A., Rapp, H., Heide, L., and Li, S. M. (2005a). Metabolic engineering of aminocoumarins: Inactivation of the methyltransferase gene cloP and generation of new clorobiocin derivatives in a heterologous host. ChemBioChem 6, 1411–1418. Freitag, A., Wemakor, E., Li, S. M., and Heide, L. (2005b). Acyl transfer in clorobiocin biosynthesis: Involvement of several proteins in the transfer of the pyrrole-2-carboxyl moiety to the deoxysugar. ChemBioChem 6, 2316–2325. Fridman, M., Balibar, C. J., Lupoli, T., Kahne, D., Walsh, C. T., and GarneauTsodikova, S. (2007). Chemoenzymatic formation of novel aminocoumarin antibiotics by the enzymes CouN1 and CouN7. Biochemistry 46, 8462–8471. Galm, U., Dessoy, M. A., Schmidt, J., Wessjohann, L. A., and Heide, L. (2004). In vitro and in vivo production of new aminocoumarins by a combined biochemical, genetic and synthetic approach. Chem. Biol. 11, 173–183. Garneau-Tsodikova, S., Stapon, A., Kahne, D., and Walsh, C. T. (2006). Installation of the pyrrolyl-2-carboxyl pharmacophore by CouN1 and CouN7 in the late biosynthetic steps of the aminocoumarin antibiotics clorobiocin and coumermycin A1. Biochemistry 45, 8568–8578. Gust, B., Challis, G. L., Fowler, K., Kieser, T., and Chater, K. F. (2003). PCR-targeted Streptomyces gene replacement identifies a protein domain needed for biosynthesis of the sesquiterpene soil odor geosmin. Proc. Natl. Acad. Sci. USA 100, 1541–1546. Gust, B., Chandra, G., Jakimowicz, D., Yuqing, T., Bruton, C. J., and Chater, K. F. (2004). lambda RED–mediated genetic manipulation of antibiotic-producing Streptomyces. Adv. Appl. Microbiol. 54, 107–128.

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Heide, L., Gust, B., Anderle, C., and Li, S. M. (2008a). Combinatorial biosynthesis, metabolic engineering and mutasynthesis for the generation of new aminocoumarin antibiotics. Curr. Top. Med. Chem. 8, 667–79. Heide, L., Westrich, L., Anderle, C., Gust, B., Kammerer, B., and Piel, J. (2008b). Use of a halogenase of hormaomycin biosynthesis for formation of new clorobiocin analogues with 5-chloropyrrole moieties. ChemBioChem 9, 1992–1999. Kieser, T., Bibb, M. J., Buttner, M. J., Chater, K. F., and Hopwood, D. A. (2000). ‘‘Practical Streptomyces genetics.’’ Norwich, UK: John Innes Foundation, Norwich, UK. Kim, C. G., Lamichhane, J., Song, K. I., Nguyen, V. D., Kim, D. H., Jeong, T. S., Kang, S. H., Kim, K. W., Maharjan, J., Hong, Y. S., Kang, J. S., Yoo, J. C., et al. (2008). Biosynthesis of rubradirin as an ansamycin antibiotic from Streptomyces achromogenes var. rubradiris NRRL3061. Arch. Microbiol. 189, 463–473. Kominek, L. A., and Sebek, O. K. (1974). Biosynthesis of novobiocin and related coumarin antibiotics. Dev. Ind. Microbiol. 15, 60–69. Li, S. M., and Heide, L. (2004). Functional analysis of biosynthetic genes of aminocoumarins and production of hybrid antibiotics. Curr. Med. Chem. Anti-Infect. Agents 3, 279–295. Li, S. M., and Heide, L. (2006). The biosynthetic gene clusters of aminocoumarin antibiotics. Planta Med. 72, 1093–1099. Luft, T., Li, S. M., Scheible, H., Kammerer, B., and Heide, L. (2005). Overexpression, purification and characterization of SimL, an amide synthetase involved in simocyclinone biosynthesis. Arch. Microbiol. 183, 277–285. MacNeil, D. J., Gewain, K. M., Ruby, C. L., Dezeny, G., Gibbons, P. H., and MacNeil, T. (1992). Analysis of Streptomyces avermitilis genes required for avermectin biosynthesis utilizing a novel integration vector. Gene 111, 61–68. Maxwell, A., and Lawson, D. M. (2003). The ATP-binding site of type II topoisomerases as a target for antibacterial drugs. Curr. Top. Med. Chem. 3, 283–303. Muth, G., Nussbaumer, B., Wohlleben, W., and Puehler, A. (1989). A vector system with temperature-sensitive replication for gene disruption and mutational cloning in streptomycetes. Mol. Gen. Genet. 219, 341–348. Reinscheid, U. M., Zlatopolskiy, B. D., Griesinger, C., Zeeck, A., and de Meijere, A. (2005). The structure of hormaomycin and one of its all-peptide aza-analogues in solution: Syntheses and biological activities of new hormaomycin analogues. Chemistry 11, 2929–2945. Rokem, J. S., Lantz, A. E., and Nielsen, J. (2007). Systems biology of antibiotic production by microorganisms. Nat. Prod. Rep. 24, 1262–1287. Sambrook, J., and Russell, D. W. (2001). ‘‘Molecular cloning: A laboratory manual.’’ New York: Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Thorpe, H. M., Wilson, S. E., and Smith, M. C. (2000). Control of directionality in the sitespecific recombination system of the Streptomyces phage phiC31. Mol. Microbiol. 38, 232–241. Williams, G. J., Goff, R. D., Zhang, C., and Thorson, J. S. (2008). Optimizing glycosyltransferase specificity via ‘‘hot spot’’ saturation mutagenesis presents a catalyst for novobiocin glycorandomization. Chem. Biol. 15, 393–401. Wolpert, M., Heide, L., Kammerer, B., and Gust, B. (2008). Assembly and heterologous expression of the coumermycin A1 gene cluster and production of new derivatives by genetic engineering. ChemBioChem 9, 603–612. Xu, H., Heide, L., and Li, S. M. (2004). New aminocoumarin antibiotics formed by a combined mutational and chemoenzymatic approach utilizing the carbamoyltransferase NovN. Chem. Biol. 11, 655–662.

C H A P T E R

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Enzymology of Aminoglycoside Biosynthesis—Deduction from Gene Clusters Udo F. Wehmeier* and Wolfgang Piepersberg† Contents 1. Introduction 1.1. Basic concepts of CAG biosynthesis 1.2. Key to the biosynthetic classes of CAGs 2. Some Key Enzymes in the Biosyntheses of Aminoglycoside Antibiotics 2.1. Investigation of a dTDP-6-deoxyhexose pathway in STR-producing Streptomyces griseus subsp. griseus DSM 40236 2.2. dTDP-D-glucose synthase (StrD, RfbA) 2.3. dTDP-D-glucose 4,6-dehydratase (StrE, RfbB) 2.4. dTDP-4-6-glucose 3,5-epimerase (StrM, RfbC) 2.5. dTDP-L-rhamnose synthase (StrL, RfbD) 2.6. L-glutamine:scyllo-inosose aminotransferase StsC (EC 2.6.1.50) from Streptomyces griseus subsp. griseus DSM 40236 2.7. L-arginine:scyllo-inosamine-phosphate amidinotransferase StrB1 from Streptomyces griseus subsp. griseus DSM 40236 (ATCC 10137) 2.8. Streptomycin-phosphate phosphatase StrK (EC 3.1.3.39) from Streptomyces griseus subsp. griseus DSM 40236 (ATCC 10137) 2.9. Myo-inositol-2-dedydrogenase (IDH; scyllo-inosose synthase) ForG (EC 1.1.1.18) from Micromonospora olivasterospora DSM 43868 2.10. Glycosyltransferases involved in CAG pathways 2.11. KanM1, KanM2, and KanN of S. kanamyceticus DSM 40500

* {

460 464 467 469

469 470 471 471 471

472

473

473

475 476 477

Department of Sports Medicine, Bergische University Wuppertal, Wuppertal, Germany Department of Chemical Microbiology, Bergische University Wuppertal, Wuppertal, Germany


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459

460

Udo F. Wehmeier and Wolfgang Piepersberg

3. Acarbose and Related Metabolites 3.1. The synthesis and modification of C7-cyclitols 3.2. The biochemistry of carbophors: A unique system for the acquisition of glucose from starch in actinomycetes References

478 479 481 487

Abstract The classical aminoglycosides are, with very few exceptions, typically actinobacterial secondary metabolites with antimicrobial activities all mediated by inhibiting translation on the 30S subunit of the bacterial ribosome. Some chemically related natural products inhibit glucosidases by mimicking oligo-a1,4-glucosides. The biochemistry of the aminoglycoside biosynthetic pathways is still a developing field since none of the pathways has been analyzed to completeness as yet. In this chapter we treat the enzymology of aminoglycoside biosyntheses as far as it becomes apparent from recent investigations based on the availability of DNA sequence data of biosynthetic gene clusters for all major structural classes of these bacterial metabolites. We give a more general overview of the field, including descriptions of some key enzymes in various aminoglycoside pathways, whereas in Chapter 20 provides a detailed account of the better-studied enzymology thus far known for the neomycin and butirosin pathways.

1. Introduction Cyclitol-containing aminoglycosides (CAGs, see examples in Fig. 19.1) and some structurally related compounds (e.g., mycothiol) (Newton et al., 2006), with very few exceptions, are natural products of actinomycetes. After the discovery of the antibiotic streptomycin in 1943 (Schatz et al., 1944), this group of substances became one of the major groups of natural therapeutics in medicine and are still in focus of much molecular biological, biochemical, and pharmaceutical research (Fig. 19.1 and Table 19.1) (Arya, 2007; Piepersberg, et al., 2007; Piepersberg and Wehmeier, 2009). Producers of CAGs with antibiotic activity are members of the highGþC Gram-positive (Actinobacteria) genera Actinoplanes, Streptomyces, Streptosporangium, Saccharopolyspora, Streptoalloteichus, Micromonospora, Dactylosporangium, and Frankia; an exception is BTR, which is produced by Bacillus circulans, a bacterium belonging to the low-GþC Firmicutes. Two distinct targets are seen with CAGs, the bacterial ribosome (antibiotics) and alpha-glucosidases (enzyme inhibitors). All antibiotically active compounds of this class of secondary metabolites act on the 30S subunit of the bacterial ribosome by binding to one of at least four sites on the 16S rRNA; there they interfere with translation in distinctly different ways: (1) STR binds tightly to the sugar-phosphate backbone in four different

H 6´ CH2NH2 H3C NHCH3 O 4´ 6´ O 2 2´ OH 1´ Gentamicin C1 2´ NH HO 3´ 4 1´ HN NH2 NH2 GEN-C1 NH=C-NH2 4´ O 2 3´ 4 5 NH2 CHO O NH2 1´ CH3 OH 5´ 3 6 2 O NH2 2´ Streptomycin H3C 3´ HO O 3 6 2´ 5 OH O Spectinomycin HO O 1 NH2 STR OH 1´ 5 HO Kanamycin B 1˝ SPC O 1 6˝ H3C-NH 1˝ O CH OH KAN-B 2 5˝ O 3˝ 1˝ OH 5˝ HO 2˝ OH O 3˝ 4˝ OH O 2˝ 5´ 4˝ NH HO 2 CH3 6˝ OH 2˝ 6´ NH 6´ 6´ CH3 OH CH2NH2 CH2NH 6˝ OH 4´ CH 4˝ 2 3 4´ O Butirosin O HN O Neomycin C HO HO 2 2´ 2˝ OH 2´ 7˝ NH NEO-C BTR HO HO 1˝ 1´ HO HOH C 2 1˝ 2 NH2 4 NH 3´ NHCH3 OH Apramycin H OH 3´ NH 2 8´ 2 4 NH2 2 6´ 2 O APR O 6 O 3 5˝ OH 3 6 O NH-CO-CH-(CH2)2 -NH2 5´ O NH2 O O 5˝ HOH2C O 5 OH 1 2´ HOH2C O 5 OH 1 OH OH 1˝ O 1´ 4˝ 6˝´ 1˝ 2˝ 4˝ HO HN NH 4˝´ CH2NH2 3˝ 2˝ 3 2 2 1˝ OH 3˝ 2˝ O 4 OH O HO 7´ 2 6´ OH OH O O OH 2˝´ 1˝´ NH OH 2 HO 2´ HO 2´ 3´ OH 3˝´ 1´ 7 NH2 O HO 6´ 5 4 O HO NH HO 6 1 OH OH O 2 HO Validamycin A HO 1 6 3 2 HC OH 6´ 3 VAL 4´ HO NH2 CH3 H HN NHCH HN 2 3 6´ 1 O OH 3 HO 4´ OH HO O OH 7 4 O 1 HO 2´ 5´ Hygromycin B OH Fortimicin A O OH 1´ HO HO OH OH 3´ H N 1´ O HO HYG-B FOR-A 2 O OH 2 4 4 O HO O 3 6´ NH2 NH 1´ CH OH OH 2 O H OH 5 Acarbose OH HS OH O NH 2 HO O ACB 2´ OH HO 6 O 1 Mycothiol H3CHN OH OCH3 OH H N-H C-C=O OH 6 4

HO 5 O O

2

NH=C-NH2

3

1

NH 2 OH

OH

NH-CH3 1 4 6 3

HO

4´

2

Figure 19.1 Examples of the chemical structures of aminocyclitol-aminoglycosides (CAGs) and of some related actinomycete metabolites. Some of the major families of CAGs are produced as mixtures of several closely related end products, where the various components are distinguished by adding a capital letter to the trivial name. Acarbose, belonging to a family of products also called amylostatins, and validamycins are secreted actinomycete enzyme inhibitors and have no antibiotic activity. Mycothiol is a nonsecreted primary metabolite occuring in most or all actinobacteria and acting as the detoxifying thiol with a role similar to that of glutathione in other groups of organisms.

462 Table 19.1 Examples of CAGs for which sequence data for complete biosynthetic gene clusters are available.a Aminoglycoside (Family, Complexb)

Streptomycin (STR) Hydroxystreptomycin Spectinomycin (SPC) Neomycin (NEO) Butirosin (BTR) Ribostamycin (RIB) Paromomycin (PAR) Lividomycin (LIV) Kanamycin (KAN) Tobramycin (TOB) Apramycin (APR) Gentamicin (GEN)

Producing organism, Strain no.

STR-related family S. griseus ssp. griseus DSM 40236 S. glaucescens GLA.0 (DSM 40716) S. netropsis DSM 40093 NEO-related family S. fradiae DSM 40063 B. circulans ATCC 21557 S. ribosidificus NRRL B-11466 S. rimosus ssp. paromomycinus NRRL 2455 S. sp. (lividus) CBS 844.73 KAN-related familyc S. kanamyceticus DSM 40500 S. sp. (tenebrarius) DSM 40477 S. sp. (tenebrarius) DSM 40477 M. echinospora DSM 43036

Biosynthetic class c

Accession code

Genes in cluster

Ca-6DOH Ca-6DOH Ca-6DOH

AJ862840 AJ006985 U70376

23 (26–31)d 18

2DOS-GA (Cb1) 2DOS-GA (Cb1) 2DOS-GA (Cb1) 2DOS-GA (Cb1) 2DOS-GA (Cb1)

AJ629247 AB097196 AJ744850 AJ628955 AJ748832

22 25 20 22 23

2DOS-GA (Cb1) 2DOS-GA (Cb1) 2DOS-GA (Cb1) 2DOS-GA (Cb1)

AJ628422 AJ810851 AJ629123 AJ628149

21 21 30 32

Fortimicin (FOR) Fortimicin (FOR) Istamycin (IST) Hygromycin B (HYG-B)

Acarbose (ACB) Acarbose (ACB) Validamycin (VAL) a

FOR-rrelated family M. olivasterospora DSM 43868 Frankia sp. CcI3 S. tenjimariensis ATCC 31603 HYG-B-related family S. hygroscopicus ssp. hygroscopicus DSM 40578 ACB-related family Actinoplanes sp. SE 50/110 S. glaucescens GLA.0 (DSM 40716) S. hygrosc. ssp. limoneus IFO 12704

FOR-type (Ca) FOR-type (Ca) FOR-type (Cb1)

AJ628421 (34)d NC_007777.1 (34)d AJ845083 26

HYG-B-type (Cb1)

AJ628642

22

Cb2-Hex Cb2-Hex Cb2-Hex

Y18523 AM409314 DQ223652

25 24 22

Only the most complete stage of sequencing is given where multiple coverage of the same genomic area is reported. The majority of aminoglycosides designated by a common name are composed of a complex mixture; chemically distinct components in a complex are distinguished by adding capital letters, and in some cases additionally by figures to the name (e.g., gentamicin C1); aminoglycoside names are abbreviated as given below. c For definition see text. d The number of genes clearly belonging to these clusters is still unclear; the for gene cluster in Frankia sp. CcI3 was found by chance in the genome sequence of this organism. S., Streptomyces; Streptoallo, Streptoalloteichus; M., Micromonospora; B., Bacillus; S., Streptomyces; c, completely sequenced; p, partially sequenced; r, resistance gene(s) outside biosynthetic clusters; sp., species; ssp., subspecies. b

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regions of the 16 rRNA, at nucleotides 13, 526, 915, and 1490, thereby affecting the initial selection and proofreading steps by fixing the 30S subunit in its ram state, which lowers the fidelity of tRNA recognition in the A-site (Carter et al., 2000); (2) SPC binds to helix 34 and inhibits translocation of peptidyl-tRNA from the A- to the P-site (Carter et al., 2000); (3) KAS binds in the path of the mRNA between the P- and the E-sites, blocking initiator tRNA binding and thereby initiation of translation (Schluenzen et al., 2006); (4) the 2DOS-GA group of CAGs (e.g., NEOs, GENs) interact with the A-site portion of 16S rRNA helix 44, disturbing correct codon-anticodon pairing at the essential nucleotides A1492/A-1493 and also inhibiting 30S subunit assembly (Carter et al., 2000; Mehta and Champney, 2002); (5) HYG-B binds to another region in helix 44 controlling codon-anticodon interaction in the A-, P-, and E-sites, by interacting with nucleotides U-1406, C-1496, and U-1498, and inhibiting ribosomal fidelity, translocation, a ribosomal ATPase, and ribosome biogenesis, also on eukaryotic ribosomes (Brodersen et al., 2000; Borovinskaya et al., 2008; McGaha and Champney, 2007). The glucosidase inhibitors target the alpha-1,4-glucoside hydrolase center of alpha-amylases, amylomaltases and similar enzymes with their nonhydrolyzable N-glycosidically linked pseudodisaccharide unit (Brayer et al., 2000; Leemhuis et al., 2004). The recent analysis of the gene clusters for almost all of the well studied CAGs and related cyclitol-containing metabolites (Table 19.1) revealed that their biosynthetic pathways are all based largely on monofunctional enzymes catalyzing single steps in the pathway, in contrast to the multistep enzymology involved in the production of polyketides or non–ribosomal peptides (see respective chapters in this volume). This knowledge has provided access to a more intense study of the biochemistry involved in the biosynthesis of aminoglycosidic compounds, which has been started in a few cases. However, none of the pathways for CAGs has yet been elucidated completely on the basis of enzymology. Pathways for other CAG-like metabolites, such as mycothiol, do not use any biosynthetic concepts or enzymes with close similarity to those involved in CAG biosyntheses (Newton et al., 2006; Piepersberg et al., 2007).

1.1. Basic concepts of CAG biosynthesis Core components in practically all natural products of this group are cyclitol units which become decorated by side groups introduced (1) via modifying steps directly on the cyclitol and (2) by glycosylation with various sugar moieties, which (3) can become further modified by pathway-specific enzymes either before or after glycosyl transfer. This general principal of design can be used to classify the aminoglycosides into biosynthetic classes mostly comprising structurally related families of end products (Table 19.1).

465

Aminoglycoside Biosynthesis

A proposal for the definition of ‘‘pathway formulas’’ classifying the CAG pathways into a general system has been published previously (Piepersberg, 1997). Therein the cyclitol pathways were given a ‘‘C’’ code according to their biochemistry. Two distinct (amino-)cyclitol pathways are used in the initiation of CAG pathways (Fig. 19.2), one based on myo-inositol (Ca pathway), the other on 1-keto-2-deoxy-cylitols (Cb pathways). The primary precursors in both cases are aldosephosphates, D-glucose-6-phosphate in Ca and Cb1, and sedo-heptulose-7-phosphate in Cb2. Pathway Ca is used in the biosynthetic pathways for STR-related CAGs, FORs, and some others of minor importance (Table 19.1, Figs. 19.1 and 19.2). The Ca pathway starts with the two-step formation of myo-inositol in a pathway and is not encoded in the cognate biosynthetic gene clusters. The reason for this is that the two-step biosynthesis of myo-inositol via myo-inositolphosphate (alternatively designated as D-myo-inositol3-phosphate or L-myo-inositol-1-phosphate) occurs by a eukaryotic type L-myo-inositol-1-phosphate synthase (INO1, EC 5.5.1.4) and

Ca HO

OH 2

4 OH

HO

3 5

D-Glucose-6-P Cb1

6

OPO32–

OH 1

L-myo-Inositol-1-P

4 OH HO 3 HO 5

Sedo-Heptulose-7-P

6

2

OH 1 O

2,5-epi-valiolone

2-Deoxy-scyllo-inosose

MOD/GT

2-DOsIA

sIA

Cb2 HO 2 4 3 OH 6 7 5 HO 1 O HO OH

(6DOH)

(2x)

(UDP-NAGA) (6DOH)

STR, SPC, KAS

2-DOS (UDP-NAGA)

IST, FOR

Paromamine

HYG-B APR

NEO, RIB, BUT, PAR, LIV

ACB

VAL

KAN, TOB, GEN

Figure 19.2 Basic CAG pathways. The biosynthetic routes for (amino-)cyclitol and pseudodisaccharide (in case of the paromamine-based and FOR-related ACAGAs) formation are schematically represented in order to classify the aminoglycosides into biosynthetic families. For details, see text. (2DO)sIS, (2-deoxy-)scyllo-inosamine; NAGA, N-acetyl-glucosamine; 6DOH, 6-deoxyhexose component; MOD/GT ¼ modification and glycosylation phase in all pathways. For aminoglycoside product families seeTable19.1.

466


myo-inositolphosphate phosphatase (IMP, SuhB; EC 3.1.3.25). In contrast to most other bacteria, these are essential metabolic activities in actinomycetes, because all actinobacteria produce mycothiol (the major cellular thiol and redox cocatalyst) as an essential metabolite. Actinobacterial genomes mostly harbor more then one gene, sometimes several genes with either Lmyo-inositol-1-phosphate synthase or inositol monophosphatase signatures. In some CAG gene clusters, however, a putative IMP enzyme for the dephosphorylation of myo-inositolphosphate seems to be encoded. For example, the strO gene in the str/sts-cluster of STR producers encodes a protein with monophosphatase signature; however, since the streptidine pathway involves two dephosphorylation steps, it is not clear at present whether StrO is catalyzing phosphorolysis from L-myo-inositol-1-phosphate (Walker, 1975; Piepersberg et al., 2007). Also, the spcA and forA genes in the respective clusters for SPC and FOR biosynthesis, which also involve a Ca cyclitol pathway, seem to encode IMP enzymes. The reason for this could be that the housekeeping enzyme activity of this enzyme is reduced in the mostly nonvegetative production phases for secondary metabolites to too low levels in these species. To render myo-inositol an aminotransferase substrate for the formation of scyllo-inosamine it has to be oxidized by a myo-inositol 2-dehydrogenase (IDH, EC 1.1.1.18). This enzyme activity, too, is normally not specific for CAG biosynthesis and seems to be encoded by all actinomycete genomes sequenced so far; it normally plays a role in the catabolism of myo-inositol in many organisms. However, in the STR- and SPC-pathways an IDH seems to be encoded by the respective gene clusters (StrI/StsB, SpcB/ SpcH) and a second cyclitol oxidase, a 1-aminocyclitol 3-dehydrogenase, prepares the cyclitol precursor later for a second transamination step. It was proven by us recently that in the FOR-producer the gene forG encodes an IDH and that the gene is essential for FOR production (see Section 2.9). In the Cb pathways the C6- and C7-cyclitols (Cb1 started by 2-deoxyscyllo-inosose synthase [see Chapter 20]; Cb2 started by 2-epi-5-epi-valiolone synthase [Stratmann et al., 1999]), being the precursors of e.g., 2-deoxystreptamine in the 2DOS-CAGs or of valienamine in the acarboserelated metabolites, are synthesized by enzymes (‘‘C-enzymes,’’ such as NeoC, KanC, AcbC) structurally closely related to and following the same mechanism as is used by dehydroquinate synthase (DHQS, EC 4.6.1.3). These enzymes are NADþ-dependent, cyclizing phosphate lyases and catalyze intramolecular aldol condensations on aldosephosphates, yielding nonphosphorylated 1-keto-2-deoxycyclitols. The Cb pathways immediately yield ketocyclitols which can be used directly as aminotransferase substrates. Thus the basic concept of CAG formation can be summarized as follows: (1) single-step enzymes—that is, enzymes with only one reaction centre, and mostly derived from sugar and cyclitol metabolism—successively activate, modify, and glycosidically link sugar precursors; (2) a second phase of decorating modification frequently follows after oligomerization; (3) the


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distribution of modification phases over the pathway varies between biosynthetic classes; and (4) cyclitol amination is a common principle in a majority of CAGs, normally catalyzed by unusual L-glutamine–dependent cyclitol aminotransferases (see description of StsC in Section 2.6). These biosynthetic concepts are similarly found in the quite variable oligosaccharidic building blocks of bacterial heteropolysaccharides, normally used as cell wall components or extracellular surface material. Therefore, anabolic cell wall metabolism might be the evolutionary and ecological (cell protection) source of CAGs (Piepersberg, 2001).

1.2. Key to the biosynthetic classes of CAGs 1. Ca-6DOH: The biosynthesis of these pseudodi- or pseudotrisaccharides is based on the formation of a cyclitol unit via the Ca pathway (streptidine in STR; bluensidine in bluensomycin; actinamine in SPC; unmodified myo-inositol in KAS) followed by a glycosidic attachment of a preformed 6-deoxyhexose (6DOH) moiety. Most modification steps occur on the monomer level. In KAS the cyclitol becomes converted to D-chiro-inositol after having been glycosylated. For the STRs, biosynthetic enzymology has been studied to a considerable extent (see Section 2.1–2.8), whereas for the SPCs and KAS only a few data are available. 2. 2DOS-GA (Cb1): The core biosynthetic design of this largest and pharmaceutically most interesting group of CAGs is based on the formation, via the Cb1-pathway, of the 2-deoxydiaminocyclitol 2DOS from scyllo-inosose and its 4-glycosylation with D-glucosamine (GA), forming the characteristic intermediate paromamine (Fig. 19.2). The modifications on subunits are performed on both the monomer and oligomer levels; for example, various modifications important for the properties of the final products, such as transaminations, methylations, and epimerization, occur during the late pathways on the oligosaccharidic intermediates. The end products are pseudotri- or pseudotetra-saccharides; APR deviates structurally from this pattern, but nevertheless clearly is a member of this group. The biosynthetic route for GENs has been studied in detail by collecting a set of blocked mutants and studying the accumulation of intermediates in them (Kase et al., 1982). Biosynthetic enzymology has been studied to some extent for some of the NEO-related compounds, to a lesser extent, for the KANs/GENs, and not at all for APR. In the NEOfamily paromamine becomes 60 -transaminated and 5-ribosylated forming the pseudotrisaccharidic RIB, an antibiotically active end product itself (Table 19.1). In the BTR-pathway, RIB becomes 1-aminoacylated with 2-hydroxy-4-aminobutyric acid. In NEO-, PAR-, and LIV-producers RIB becomes further glucosaminylated in the 300 -position and modified in this fourth glycosidic moiety. In the KAN/GEN-family paromamine is

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6-glycosylated by neutral sugars, either a C6-pyranose (KAN) or a C5-pyranose (GEN). These pseudotrisaccharidic precursors are 60 - and 300 -transaminated and can be modified further by 300 -N- and 400 -C-methylation steps. In the producers of GEN-related CAGs a quite complicated, branching late pathway, characterized by 60 -C-methylation for only part of the pool of intermediates, 30 ,40 -didehydroxylation, 60 -epimerization, and 60 -N-methylation, creates a complex of related GEN products varying in proportion between individual producer strains (Piepersberg et al., 2007). Another unusual situation exists in the producers of the so-called nebramycin complex, to which TOB, a 30 -dehydroxy-KAN-B, and APR belong, though the latter is a single-substituted 2DOS derivative (Fig. 19.1). We and others have studied the genetics of TOB/APRbiosynthesis in two different organisms, S. sp. ‘‘tenebrarius’’ DSM 40477 (Table 19.1) and Streptoalloteichus hindustanus DSM 44523 (accession codes AJ875019 and AB103327). In both species, there are two independent tob- and apr-clusters, both encoding an almost complete pathway of their own; the TOB-pathway is equivalent to that of KAN-B. However, the two genes necessary for the 30 -dehydroxylation of KAN-B are located in the apr-cluster (aprD3, aprD4 genes), and this modification is shared between the two pathways (Piepersberg et al., 2007). 3. HYG-B-type (Cb1): Biosynthesis of the cyclitol in the third unique pathway is again based on the Cb1 product 2DOS. However, formation of the pseudotrisaccharidic end product principally deviates from the 2DOS-GA type by the glycosidic attachment of a neutral hexose to the 5-position of 2DOS and of an aminoheptose moiety via an unusual orthoester bond to the 20 - and 30 -positions of the hexose (cf. Fig. 19.1). The modification steps have to be assumed to occur on both the monomer and the oligomer levels; however, biosynthetic enzymes have not been studied in vitro. 4. FOR-type (Ca or Cb1): The biosynthesis is based on the formation of a monoamino cyclitol unit, scyllo-inosamine in FORs, and 2-deoxyscyllo-inosamine in ISTs, from either myo-inositolphosphate (Ca route) or scyllo-inosose (Cb1 route) and glycosidic attachment of a D-glucosamine moiety. The modification of subunits is achieved mostly on the condensed level of these pseudodisaccharides, whereby the cyclitol also becomes highly decorated with additional side groups (see the structure of FOR-A in Fig. 19.1). A reasonable picture of the complicated FOR- and IST-pathways comes from structural analyzes of intermediates accumulated in a series of mutants (Odakura et al., 1984). A significant paralleling between the FOR- and GEN-pathways, evident from the use of several common biosynthetic steps for 30 ,40 dehydroxylation and 60 -C-methylation, is reflected by a number of unique genes with very high DNA homology between the for- and gen-clusters (for details, see Piepersberg et al., 2007).


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5. Cb2-Hex: The biosynthesis of these malto-oligosaccharide analogues is based on the formation of a C-7 cyclitol unit (valienol, valiolol, valienamine) via the Cb2 pathway and glycosidic attachment of (6-deoxy)hexose units; they are malto-oligosaccharide analogues, that is, linear chains of up to six (pseudo-)saccharidic units linked by glycosidic bonds structurally mimicking a-1,4-glucosides. Some enzymes involved in their biosynthesis and probable cyclic conversion have been studied (see Section 3).

2. Some Key Enzymes in the Biosyntheses of Aminoglycoside Antibiotics Some enzymology involved in key steps of the STR, NEO, KAN, and ACB-pathways investigated by us are described in subsequent sections; the other enzymes important for the biosyntheses of the 2DOS-CGA compounds BTR and NEO are described in Chapter 20.

2.1. Investigation of a dTDP-6-deoxyhexose pathway in STR-producing Streptomyces griseus subsp. griseus DSM 40236 In our analyses of the STR biosynthetic gene cluster and its encoded enzymatic tool box we found that four genes, strDEL and strM, organized in the two transcription units strDEL and strNB2M, had about equally high similarity to the dTDP-L-rhamnose biosynthetic genes in the rfb-locus of enterobacteria, that is, rfbA (rmlA), rfbB (rmlB), rfbC (rmlC), and rfbD (rmlD) from Gram-negative bacteria producing a lipopolysaccharide containing L-rhamnose (Reeves, 1993). These are equivalent to many genes occurring in all groups of bacteria and are mostly involved in the biosynthesis of extracellular polysaccharides; for example, Escherichia coli harbors a second set of rfbA,B-related genes, rffH and rffG, involved in the synthesis of enterobacterial common antigen. The encoded proteins have the following enzyme activities: RfbA (RmlA) is the dTDP-glucose synthase (glucose-1phosphate thymidylyltransferase; EC 2.7.7.24); RfbB (RmlB), the dTDP-glucose 4,6-dehydratase (EC 4.2.1.46); RfbC (RmlC), the dTDP4-keto-6-deoxyglucose (dTDP-6-deoxy-D-xylo-4-hexulose) 3,5-epimerase (EC 5.1.3.13); and RfbD (RmlD), the dTDP-L-rhamnose synthase (dTDP-6-deoxy-L-lyxo-4-hexulose reductase; EC 1.1.1.133). Grisebach and collaborators had first described a dTDP-hexose pathway involved in STR biosynthesis in S. griseus subsp. griseus N2-3-11 (equivalent to wildtype strain ATCC 10137, DSM 40236) (Ortmann et al., 1979; Grisebach, 1978). This was postulated to comprise the first three of the four steps of the dTDP-L-rhamnose pathway, forming dTDP-4-keto-L-rhamnose, and

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finally its reductive conversion to dTDP-dihydrostreptose. The final step should be similar to dTDP-L-rhamnose synthesis and was postulated to involve, besides reduction at C-5/C-6, a C-chain reorganization, such that C-3 of the hexose is branched out from the linear chain and, forming a furanosidic ring sugar with two C1-branches at the C-3 and C-4 atoms (cf. Fig. 19.1; positions 30 and 40 in the formula of STR). In order to correlate these earlier findings with the genetic record as it became evident from the str/sts-cluster, we investigated the enzyme activities of the heterologously produced proteins StrD,E,M,L in comparison to the well studied enzymes RfbA,B,C,D from Gram-negative bacteria; the dTDP-activated sugars were identified and their accumulation quantitatively measured by HPLC.

2.2. dTDP-D-glucose synthase (StrD, RfbA) This enzyme was tested either (1) in 500-ml assay volume at pH 8.0 and 37 containing 50 mM Tris/HCl buffer, 10 mM MgCl2, dTTP 2 mM, a-Dglucose-1-phosphate 10 mM, 170 ml PPi-reagent (Sigma Deisenhofen, Germany), enzyme preparations at an activity in the range of 1 to 100 nkat; or (2) in 300-ml final volumes with the same composition but with the addition of 35 nkat inorganic pyrophosphatase instead of PPi-reagent. The reaction was monitored in (1) by photometrically measuring the consumption of NADH at 340 nm; and in (2) 20-ml samples were withdrawn from 150-ml reaction mixtures, which were generally taken after 30, 45, or 60 min, and in which reactions were stopped by boiling for 1 min, and insoluble material was separated by centrifugation, and then subjected to HPLC analysis (Beckmann Instruments, Munich, Germany; pump module 125, autosampler 502, UV detector 166, radioisotope detector 171); nucleotide-activated sugars were measured at 260 nm for unlabeled compounds or on a radioisotope detector for 14C-labeled sugars. Chromatography was run on a reversed-phase column (Eurospher 100 C18, Knauer, Berlin, Germany; elution by 0 to 40% acetonitrile gradient). The background activity for dTDP-glucose synthase was surprisingly low in STRproducing S. griseus. StrD could be overexpressed to a considerable extent in Streptomyces lividans and E. coli; the product, dTDP-D-glucose, eluted at 35.9 min from the HPLC. Like the standard enzyme RfbA, StrD had maximal substrate specificities for dTTP and D-glucose-1-phosphate; only with the substrate combinations UTP/D-glucose-1-phosphate, dTTP/Dglucosamine-1-phosphate, and UTP/N-acetyl-D-glucosamine-1-phosphate could minor activities also be detected. RfbA and StrD are both feedback inhibited by dTDP-L-rhamnose; e.g., in reaction assays containing 10 mM and 2.5 mM of the substrates glucose-1-phosphate and dTTP, respectively, 445 mM of dTDP-L-rhamnose inhibited both enzymes to 71 to 72% residual activity.


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2.3. dTDP-D-glucose 4,6-dehydratase (StrE, RfbB) These enzymes were measured in similar assays as in (1.b): 300-ml test volumes contained 50 mM Tris/HCl buffer (pH 7.5), 2 mM dTDP-glucose, and in the range of 1 to 100 nkat of enzyme activity; for the streptomycete enzyme sources the buffer concentration was enhanced to 150 mM and 1.5 mM NADþ was added. For detection of substrates/ products HPLC was used (see Section 2.2); dTDP-4-keto-6-deoxyglucose eluted at 37.8 min. The StrE protein turned out to be a dTDP-glucose 4,6dehydratase with very similar properties to RfbD, with the exception that StrE did not contain firmly bound NADþ as a prosthetic group, as is the case for RfbB. The 4,6-dehydratase background activities were quite low in S. griseus N2-3-11 (vegetative phase) and S. lividans TK23. StrE could be overexpressed in S. lividans, but not in E. coli. StrD did not convert other activated sugars, such as dUDP-D-glucose, UDP-D-glucose, or CDP-D-glucose.

2.4. dTDP-4-6-glucose 3,5-epimerase (StrM, RfbC) The 3,5-epimerase activity was assayed as a coupled reaction together with the subsequent dTDP-L-rhamnose synthase because it turned out that the equilibrium of the epimerization lies fully on the substrate side and no reaction could be detected in the absence of a product-consuming reaction. Test volumes of 300 ml contained 50 mM Tris/HCl buffer pH 7.5, 2 mM dTDP-4-keto-6-deoxyglucose, 2.5 mM NADPH, 35 nkat RfbD (nonlimiting enzyme activity), and variable concentrations of StrM or RfbC preparations. The 3,5-epimerase activity could be measured both photometrically by NADPH consumption and in HPLC separations, where the dTDP-L-rhamnose could be detected and eluted at 35.8 min. In both assay systems, identical values for the specific activity of StrM/RfbC were measured.

2.5. dTDP-L-rhamnose synthase (StrL, RfbD) This enzyme was assayed in the same test system as for the dTDP-4-6glucose 3,5-epimerase (see Section 2.4) with the difference that the content of StrM (RfbC) was fixed to 35 nkat and the activity of the dTDP-Lrhamnose synthase StrL (RfbD) was varied. Both StrL and RfbD could be highly overexpressed over the background activities of their donor strains and in E. coli BL21 and S. lividans TK23, respectively, and they have identical substrate and product specificities. Both enzymes are inhibited by their product dTDP-L-rhamnose; e.g., residual activity is 50 to 55% at a product concentration of 380 mM. Therefore, StrL clearly is a dTDP-L-rhamnose synthase and has no dTDP-dihydrostreptose synthase activity. To further

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exclude production of a different product (dTDP-L-dihydrostreptose) by StrL this enzyme was enriched from a protein preparation of the overproducing strain of S. lividans TK23 and tested in the presence of a RfbDfree RfbC preparation obtained by DEAE cellulose chromatography from the RfbC-overproducing strain of E. coli BL21. The nucleotide-activated sugar product was purified and analyzed by NMR as described elsewhere (Elling et al., 2005) and clearly was dTDP-L-rhamnose. Also, if [14C]dTDP-4-keto-6-deoxyglucose (prepared from 14C-a-D-glucose-1-phosphate by the reactions catalyzed by RfbA and RfbB; see Section 2.2–2.3) was incubated with cell-free extracts of S. griseus N2-3-11 from various culture states and for various incubation times using the same test conditions only [14C]-dTDP-L-rhamnose was yielded in all cases. Thus, no evidence for the presence of a dTDP-L-dihydrostreptose synthase with the same properties as a dTDP-L-rhamnose synthase (Grisebach, 1978; Wahl and Grisebach, 1979) could be detected in STR-producing S. griseus.

2.6. L-glutamine:scyllo-inosose aminotransferase StsC (EC 2.6.1.50) from Streptomyces griseus subsp. griseus DSM 40236 During the pioneering work on streptidine biosynthesis by Walker and collaborators (see overview in Walker, 1975a), the two aminotransferase (AT) activities needed for the introduction of two amino groups into the cyclitol moiety were identified. It turned out that the first step was dependent on L-glutamine as an amino donor and that unusually the alpha amino group of this amino acid was the donor group transferred, whereas in the second step L-alanine was the donor (Walker, 1975b,c). The first step, catalyzed by L-glutamine:scyllo-inosose (2,4,6/3,5-pentahydroxycyclohexanone) AT (EC 2.6.1.50), was correlated with a particular enzyme protein and gene product (StsC) only after most of the DNA sequence of the str/stscluster from the producer S. griseus and methods for overproduction of genetically engineered streptomycete genes became available (Ahlert et al., 1997; Hopwood et al., 1985). Since the str/sts-cluster contains three AT genes, strS, stsA, and stsC, identification of the first-step cyclitol AT had to include all three encoded enzymes. For this purpose, the three genes were subcloned in expression vectors for both E. coli BL21(DE3) (employing vectors pET11a, pET16b) and Streptomyces lividans (pIJ6021) (Takano et al., 1995), and introduced by transformation and overexpressed in both host systems. Except for low amounts of StsA, only insoluble inclusion bodies of all three proteins were obtained in E. coli. However, reasonable amounts of soluble protein of all three enzymes were yielded in the streptomycete host. In our laboratory, the three in vitro test systems described by Walker and coworkers (Walker, 1975b; Lucher et al., 1989) were used in all


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investigations of the StsC enzyme (Ahlert et al., 1997): (1) for the qualitative measurement of this AT the monoamino product scyllo-inosamine (1-amino-1-deoxy-scyllo-inositol) and of the alpha-ketoglutaramate (2-oxoglutaramate) byproduct of the forward reaction were measured, when 2-deoxy-scyllo-inosose and L-glutamine were used as substrates; (2) for higher sensitivity the radioactive exchange reaction between [14C]labeled scyllo-inosamine and nonlabeled scyllo-inosose (backward reaction) was employed; (3) the forward reaction was also measured by use of [14C]scyllo-inosose as acceptor substrate in a test system similar to the first one, where the product, [14C]-labeled scyllo-inosamine was separated from the substrate by cation-exchange chromatography on Dowex 50W-X8. Later it was found that all the aminotransferases involved in the biosynthesis of the diaminocyclitol 2DOS (1-amino-1,2-dideoxy-scyllo-inositol; see Section 1.1 and Fig. 19.2) use the same mechanism as StsC and, based on primary structure similarity, are in the same subfamily of the aminotransferase I–II superfamily (Lucher et al., 1989; Piepersberg et al., 2007).

2.7. L-arginine:scyllo-inosamine-phosphate amidinotransferase StrB1 from Streptomyces griseus subsp. griseus DSM 40236 (ATCC 10137) During the pioneering work on streptidine biosynthesis by Walker and collaborators (Walker, 1975a,d) the two amidinotransferase activities involved in this branch of the STR pathway were first identified and in vitro test systems described. Later work on these enzymes always relied on the nonspecific test system developed by these authors, which uses hydroxylamine as acceptor and L-arginine as donor substrates and yields hydroxyguanidine as the product. Later a well characterized mutant strain (SD141) blocked in the first step amidinotransferase activity was also isolated from the wildtype strain of Streptomyces griseus subsp. griseus ATCC 10137 by others (Ohnuki et al., 1985). This mutant was subsequently used to identify the encoding gene, strB1, on cloned DNA from the same organism. The DNA sequence and regulation of transcriptional activity of this and the surrounding genes became available in genetic studies (Distler et al., 1987, 1992). More recently the crystal structure of the StrB1 protein was also determined (Fritsche et al., 1998).

2.8. Streptomycin-phosphate phosphatase StrK (EC 3.1.3.39) from Streptomyces griseus subsp. griseus DSM 40236 (ATCC 10137) The gene clusters for the production of STRs, PAR, LIV, and APR each encode a gene, strK, parZ, livZ, and aprZ, for an extracellular phosphatase homologous to and perfectly matching the domain structure of alkaline

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phosphatases (EC 3.1.3.39, alkPPc superfamily, pfam00245) (Mansouri et al., 1991). The supernatants of S. griseus DSM 40236 strain N2-3-11, fermented in 10 l of solid-free TSB (Oxoid, Hamburg, Germany) as the production medium, was tested after various times of culturing for STRphosphate phosphatase activity in the following way: assays contained in 100-ml volumes 30 mM Tris/HC1, pH 8.0, 10 mM MgC12, 40 to 60 mM substrate, and variable quantities of supernatants cleared by centrifugation (or enzyme preparations); incubation was at 30 . Hydrolysis was measured by the phosphocellulose paper-binding assay (Haas and Dowding, 1975) in 10-ml aliquots taken at various times from the above incubation mixtures. As specific substrates [32P]-labeled STR-6-phosphate and STR-300 -phosphate were prepared by use of the STR-resistance phosphotransferases AphD (APH(6)) and AphE (APH(300 )), both encoded in the same producer strain S. griseus N2-3-11, expressed from plasmids pJDM1 and pJDM40 as described (Distler and Piepersberg, 1985; Distler et al., 1985; Heinzel et al., 1988). After incubation these were separated from adenine nucleotides by passage over Dowex 1 8 (Serva, Heidelberg, FRG) and yielded specific radioactivities in the range of 10 to 20 mCi/m mol. STR-phosphate-specific phosphatase was partially purified from culture supernatants of S. griseus N2-3-11 by 70% ammonium sulphate precipitation followed by octyl-sepharose (Pharmacia, Freiburg, FRG) column chromatography (the enzyme was bound in the presence of 30% ammonium sulphate and was eluted with salt-free buffer). Also, StrK was cloned into Streptomyces vector pIJ702 (Katz et al., 1983) and expressed from the promoter of the mel gene in pKMWK1/S. lividans 66 strain TK23, then tested in its supernatant in the same way as above. STR-phosphate phosphatase activity first appeared 25 h after inoculation in the culture supernatant of S. griseus N2-3-11 and preceded the first appearance of STR, then paralleling the accumulation of the antibiotic and an increase of the pH to 8.5 in the medium. Maximum relative activity of STR-6-phosphate phosphatase was reached at a value of about 85% hydrolysis in a standard assay (40 ml supernatant; incubation time 30 min). The hydrolysis of STR-6-phosphate was about sixfold faster than that of STR-300 -phosphate. The enzyme was inactivated by temperatures above 60 and pH values below 7.0. Obviously, only very little StrK protein was made, since after enrichment no protein band of the expected size could be detected in polyacrylamide gels. The same enzyme in comparable relative activities was secreted into culture supernatants of S. lividans/pKMWK1 (strK). In contrast, the culture of the control strain (S. lividans/pIJ702) was free of any comparable phosphatase activity (Mansouri et al., 1991). Therefore, StrK was interpreted to be an alkaline phosphatase specific for STR phosphates (EC 3.1.3.39).


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2.9. Myo-inositol-2-dedydrogenase (IDH; scyllo-inosose synthase) ForG (EC 1.1.1.18) from Micromonospora olivasterospora DSM 43868 In the FOR-production gene clusters (Table 19.1) a gene, forG, was found to encode a protein with highest similarity to possible IDH candidates encoded by the genomes of S. coelicolor A3(2) (e.g., 50% aa identity to SCO6255) and other actinomycetes. To test for this possibility we cloned, expressed, and assayed the ForG protein and knocked-out the forG-gene in the producer strain (Schuermann, B., Wehmeier, U. F., and Piepersberg, W., unpublished): The gene was amplified by PCR, using primers that introduced useful single restriction sites at both ends of the reading frame, cloned into various plasmid vectors and expression strains of E. coli and S. lividans 66, which were tested for overproduction of the soluble ForG protein. This goal could be achieved best in E. coli BL21(DE)-Rosetta/ pBSW3-forG (vector pET16b; Novagen, Darmstadt, Germany; forG-ORF N-terminally elongated by His-tag) by demonstrating a new band of approximately 40 kDa (calculated MW of native protein 36.8 kDa) in the supernatant of the crude extract after induction on protein gels (Scha¨gger and von Jagow, 1987). Preparation of enriched His-tag-ForG was achieved by Ni-NTA affinity chromatography. The ForG protein was tested for IDH activity by either (1) monitoring the absorption at 340 nm in 1-ml assay volumes containing 40 mM myo-inositol, 0.5 mM NADþ, 100 mM Tris/ HCl (pH 9.0), and various concentrations of enzyme preparations; or (2) by measuring the absorption at 570 nm in a more sensitive MTT-test (according to Lengeler and Lin, 1972) consisting of 600 ml mix A (3.3 ml 1 M Tris/ HCl, pH 9; 0.7 ml MTT (¼ (3-[4,5-dimethylthiazol-2-yl])-2,5-diphenyltetrazoliumbromide, 0.5 mg/ml); 0.02 ml Triton X-100; 3 ml H2O), 100 ml 0.4 M myo-inositol, 10 ml 0.5 mM NADþ, 100 ml 5-methylphenaziniummethosulfate (1 mg/ml), and 10 to 40 ml enzyme preparation. As controls, assays containing 0.1 U of commercial IDH (from Sigma-Aldrich, Steinheim, Germany) were also run. For direct detection and quantitation of newly formed scyllo-inosose, ion chromatography (DIONEX DX-500 system with electrochemical detector, and column material CarboPac MA1, and Chromelion 6.2 software, Dionex, Idstein, Germany) was employed. The results are summarized as follows: (1) ForG expression significantly enhanced background IDH activity in both E. coli and S. lividans (up to 20-fold); (2) an enrichment factor of 20-fold was again achieved after Ni-NTA chromatography; and (3) the ForG product was identified as scyllo-inosose by ion chromatography. Also, forG knock-out mutants were constructed by replacement of a 200-bp segment internal to the forG gene by the omega-aacC4 APR-resistance cassette (BlondeletRounault et al., 1997) in plasmid pBSW2 and by its introduction by transformation into protoplasts of the FOR-producer M. olivasterospora

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DSM 43868 according to a published method (Hasegawa et al., 1991). APR-resistant transformants generated by double crossing over with the native forG gene could be selected in which the native forG-gene had been knocked out, as proven by CR-experiments. These mutants no longer produced FOR (compared to the wildtype strain DSM 43868) as could be measured by lack of inhibition zone formation around agar plugs cut from fully grown plate cultures on a production medium (ATCC no. 5) and placed on fresh plates inoculated with spores of Bacillus subtilis as a tester strain. Therefore, the forG-gene is essential for FOR-production.

2.10. Glycosyltransferases involved in CAG pathways The glycosyltransferases (GTs) involved in the STR-related (Ca-6DOH) biosynthetic pathways are uncharacterized and practically nonidentified among the gene products of the respective gene clusters (str/sts, spc, and kas). For instance, in the case of the STR pathway of S. griseus, for which two GTs are necessary, StsF is the only candidate protein with some distant similarity to known GTs. The SpcF and KasA proteins are more reliable candidate GTs for the single glycosyltransfer steps involved in the SPC- and KAS-pathways, respectively (Piepersberget al., 2007). In this respect, better data are available for the biosyntheses of 2DOS-containing and related CAGs (see also Chapter 20). These are mostly Cb1 pathways (a few exceptions are made via the Ca pathway, e.g., FOR), in which an enzyme family of GTs, the strongly related ‘‘M-enzymes’’ (e.g., KanM, NeoM, ForM; glycosyltransferases group 1; data base accession: InterPro: IPR001296, pfam PF00534, Glycos_transf_1; e.g., KanM belonging to the cluster COG0438 of RfaG-like GTs), catalyze the first attachment of a sugar residue, 2-NAc-D-glucosaminyl, to position 4 of an aminocyclitol unit. The aminocyclitol is the diaminocyclitol 2DOS in the NEO-, KAN-, and HYG-B-related families; however, it can also be a monoaminocyclitol, scyllo-inosamine or 2-deoxy-scyllo-inosamine, as in the FOR- and ISTpathways, respectively (see Section 1.1). Several members of the ‘‘M’’enzymes and of the deacetylases, deacetylating NAc-glucosaminyl moieties are subsequently used in the respective pathways. Interestingly, the second N-acetylglucosaminyltransferase in the NEO-type pathways, NeoF, and related ‘‘F’’-enzymes in the BTR-, PAR-, and LIV-pathways, are not members of the ‘‘M’’-type subfamily of group 1 GTs, whereas the second GTs used in the KAN/GEN-type pathways (e.g., KanM2, are, though they are transferases for glucosyl- or xylosyl-residues). From the KAN and NEO pathways the following gene products have been studied in a comparative investigation in our laboratory and tested for enzymatic activity (Mandt, C., Wehmeier, U. F., and Piepersberg, W., unpublished data). Generally, for the genes kanM1, kanN, and kanM2 from the KAN producer Streptomyces kanamyceticus DSM 40500, the following cloning


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strategy was employed. Genes were amplified from the genomic DNA of the wildtype CAG-producing strains as their ORFs by PCR using primers introducing unique restriction sites at both ends and ATG start codons, where GTG start codons are naturally present. PCR fragments were cloned into suitable E. coli selection plasmids, such as pUCPU21 (Wehmeier, U. F., unpublished), screened on X-Gal in E. coli DH5a and checked by sequencing. The respective genes were then subcloned into suitable E. coli/ Streptomyces shuttle and expression vectors, such as pUWL201PW (Doumith et al., 2000) or pJOE2775 (Khalameyzer et al., 1999), or into Streptomyces vectors, such as pMACP (Wehmeier, U. F., unpublished), which can be transferred by conjugation from E. coli ET12567/pUZ8002 to Streptomyces spp. and allow expression there. For comparative purposes, the same cloning and expression strategy was used for neoM, neoF, and neoD, from the NEO producer S. fradiae DSM 40063.

2.11. KanM1, KanM2, and KanN of S. kanamyceticus DSM 40500 KanM1 and KanM2 had been postulated to be the first and second step GTs forming a pseudotrisaccharidic intermediate (6-O-Glc-paromamine) via 4-O-NAc-D-glucosaminylation (20 -NAc-paromamine synthase) and 6-OD-glucosylation of 2DOS, using UDP-2-NAc-D-glucosamine and UDPD-glucose as donor substrates, respectively. KanN was assumed to be a monofunctional deacetylase deacetylating 20 -NAc-paromamine in the KAN-pathway; in contrast, the related deacetylase, NeoD, was shown to be a bifunctional N-deacetylase deacetylating both NAc-D-glucosaminyl residues during the formation of 6000 -hydroxy-NEO-C, that is, in 20 -NAc-paromamine, and in 2000 -NAc-6000 -hydroxy-NEO-C (Lewellyn and Spencer, 2006; Yokoyama et al., 2008). Structurally, both KanN and NeoD are members of the LmbE-related protein family (COG2120; pfam02585, PIG-L, GlcNAc-PI de-N-acetylase family). KanM1, KanN, NeoM, NeoF, and NeoD from the producer strains S. kanamyceticus and S. fradiae could be actively overexpressed, whereas KanM2 could only be expressed as insoluble protein aggregates and did not show enzymatic activity under the tested conditions. For testing the KanM1 and NeoM activities in cell-free extracts of the expression strains S. lividans 66 TK64/pCM201PW-kanM1 and S. lividans TK23/pCMW-neoM as compared to the background activity in the recipient strain plus vector plasmid (S. lividans TK23/pUWL201PW), assays of 300 ml of total volume contained 50 mM Tris/HCl, 10 mM MgCl2, 2 mM UDP-2-NAc-D-glucosamine, 1 mM 2DOS, and 30 ml of cell-free extract, pH 7.5 by incubation at 37 for up to 6 h. The result clearly revealed a conversion of 2DOS (RF 0.53) and UDP-NAc-D-glucosamine to 20 -NAcparomamine (RF 0.59); paromamine had an RF of 0.22 in the same TLC separation system (TLC plates were Silica Gel F254, Merck, Darmstadt,

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Germany; running solution was 50 mM KH2PO4); spots were visualized after development of plates by spraying with ninhydrin-solution and subsequent heating with a heat gun. Similarly, NeoF was tested for the third GT activity of the NEOpathway (2-NAc-glucosaminyltransferases II, 300 -NAc-glucosaminylation; cf. structure of NEO-C in Fig. 19.1) in extracts of S. lividans TK23/ pCMW-neoF with ribostamycin (5-ribosylparomamine; RF 0.40) as acceptor and UDP-NAc-D-glucosamine as donor substrates. The result was the detection of a new spot (RF 0.48; interpreted to be 2000 -NAcNEO-C), in the TLC system used (run in a solution of methanol/chloroform/12.5 % (w/v) ammonia in water ¼ 1/0.2/1; visualization of spots was as for NeoM assays). The product obtained in the NeoF assays was used as substrate in deacetylation assays with KanN and NeoD. The deacetylating activities of the proteins KanN and NeoD were tested by adding samples of cell-free extracts of the expression strain E. coli DH5a/ pCMJOE-kanD to various GT-assays, after being fully converted. Enzyme assays were incubated overnight at 37 in a total volume of 300 ml containing 50 mM Tris/HCl, pH 7.6, 10 mM MgCl2, 60 ml of KanM- or NeoFassay and 30 ml of cell-free supernatant with enriched NeoD. Aliquots of 2 ml of this enzyme assay were spotted on TLC plates (silica gel F254, Merck, Darmstadt, Germany) run in a solution of 50 mM KH2PO4, and visualized by spraying with ninhydrin-solution and subsequent heating with a heat gun. NeoD hydrolyzed both acetylated substrates to paromamine and NEO-C, respectively, thereby proving to be at least a bifunctional enzyme with a broader substrate range. KanN deacetylated only 20 -NAc-paromamine, but not 2000 -NAc-NEO-C, proving it to be a monofunctional deacetylase with a more restricted substrate range relative to NeoD. Kudo and Eguchi and collaborators recently published similar data for the same set of glycosyltransferase/deacetylase enzymes from the NEO-pathway (Yokoyama et al., 2008; see Chapter 20 in this volume).

3. Acarbose and Related Metabolites Acarbose (ACB; Fig. 19.1) is a pseudotetrasaccharide composed of the C7-cyclitol valienamine, 4,6-dideoxyglucose and maltose that is found, together with higher homologues (i.e., ACB-containing metabolites elongated up to 30 sugar residues bound to the maltose moiety and/or at C4 of the cyclitol; see Fig. 19.1) in the culture medium of Actinoplanes sp. SE50/110 when grown on starch or malto-oligosaccharides (reviewed in Wehmeier and Piepersberg, 2004). Similar secondary metabolites seem to be made by a wide range of bacteria, including nonactinomycetes (Mahmud et al., 2007). ACB is produced industrially using developed strains of Actinoplanes sp. SE50/110,


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and it is used pharmaceutically (Glucobay/Precose) in the treatment of diabetes patients, to slow down the intestinal release of glucose. The ACB gene cluster from Actinoplanes sp. SE50/110 was identified and sequenced (Wehmeier, 2003; Wehmeier and Piepersberg, 2004; see Table 19.1). The 25 known acb genes from the producer strain Actinoplanes sp. SE50/110 encode various functions, including biosynthesis of a 6-deoxyhexose (6DOH) and a C7-cyclitol (valienol; a Cb2 product), intra- and extra-cellular anabolism and catabolism of alpha-1,4-glucosides, and uptake and export of (oligo)-saccharides. Recently, a second ACB biosynthetic gene cluster from Streptomyces glaucescens (gac cluster) was identified and sequenced (AM409314) (Rockser and Wehmeier, 2008). The differences between the set of encoded proteins in the acb and gac gene clusters indicate that (1) there is a major difference between the routes for C7-cyclitol formation (see Section 3.1) and (2) that a homologue of the acarviosyltransferase (AcbD) (see Section 3.2.1), an important enzyme in the extracellular metabolism of ACB, seems to be lacking in S. glaucescens GLA.0. This indicates not only that the latter strain produces a different metabolite spectrum compared to Actinoplanes sp. SE50/ 110, but also that the gac cluster encodes exporters, extracellular and ACBinsensitive starch-degrading enzymes, an ACB-specific importer, an intracellular amylomaltase, and an ACB kinase. We postulate that all these findings suggest a function of ACB and related compounds as a carbophor, that is, as an extracellular trap for glucose or malto-oligosaccharides, which, when attached to it, can only be imported and metabolically used by the various molecular tools provided by the acb or gac clusters and maybe its analogues, such as the val cluster in S. hygroscopicus. ssp. limoneus IFO 12704 (see Section 3.2).

3.1. The synthesis and modification of C7-cyclitols The Cb2 metabolite 2-epi-5-epi-valiolone is the common C7 cyclitol precursor in ACB (amylostatins), validamycins, and other biosynthetic pathways (Stratmann et al., 1999; reviewed in Wu et al., 2007). 2-epi-5epi-valiolone is synthesized from sedo-heptulose-7P by 2-epi-5-epi-valiolone synthases. The AcbC protein from Actinoplanes was the first member of this enzyme family to be studied (Stratmann et al., 1999). The sequence similarity of AcbC to the AroB-related DHQS proteins, cyclizing 3-deoxy-Darabino-heptulosonate 7-phosphate (DAHP) to dehydroquinate, suggested that this enzyme family catalyzes synthesis of the C7-cyclitol. The enzymatic activity of AcbC was proven using the AcbC protein enriched in extracts from strain S. lividans 1326/pAS8/7 (an overexpressing plasmid clone of acbC in vector pIJ6021) (Takano et al., 1995), which was shown to cyclize sedoheptulose 7-phosphate, but no other heptulose-7-P (Stratmann et al., 1999). The enzyme assay was carried out at 30 in a 100-ml volume containing 20 mM potassium phosphate (pH 7.4), 0.05 mM CoCl2, 2 mM KF, 1 mM NADH, 5 mM sedoheptulose 7-phosphate, and 100 mg of protein.

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The reaction of C7-cyclitol cyclases could be monitored by thin-layer chromatography analysis (solvent system, n-butanol:ethanol:H2O, 9:7:4). The product had the same properties as synthetic 2-epi-5-epi-[6-2H2]valiolone (Stratmann et al., 1999). Using the acbC gene as a probe, a number of gene clusters for C7-cyclitol containing metabolites have been identified, such as the gene clusters for validamycin A (VAL) (Bai et al., 2006; Singh et al., 2006; Wu et al., 2007). Meanwhile, a number of AcbC-like proteins have been identified and active site residues from the available 2-epi-5-epivaliolone synthases revealed striking dissimilarities with DHQ synthases because one-third of the active-site residues are consistently altered in 2-epi-5-epi-valiolone synthases (Wu et al., 2007). The next steps after the cyclization obviously differ remarkably in the various pathways. For the biosynthesis of ACB in Actinoplanes, Zhang et al. (2002) have shown that 2-epi-5-epi-valiolone is phosphorylated to 2-epi-5-epi-valiolone-7-P by the enzyme AcbM (tested in 25 mM Tris-HCl, 10 mM MgCl2, [20 mM NH4Cl] 10 mM 2-epi-5-epi-valiolone adjusted to pH 7.6, using cell free extracts from S. lividans TK64 harboring plasmids pCW4123M) (Zhang et al., 2002) and then 2-epi-5-epi-valiolone-7-P is epimerized to 5-epivaliolone-7-P by AcbO, a new type of epimerase, which is obviously cofactor independent (Zhang et al., 2003). The assays with AcbO were performed under the same conditions described above for AcbM using 2epi-5-epi-valiolone-7-P as substrate and cell-free extract from S. lividans TK64/pMJO7 (Zhang et al., 2003). In the biosynthesis of VAL, 2-epi-5epi-valiolone is first epimerized to 5-epi-valiolone, and then water is eliminated to give valienone. These two steps are catalyzed by the bifunctional enzymes (epimerase/dehydratase) ValK (val gene cluster from Streptomyces hygroscopicus 5008) (Bai et al., 2006) or VldD (vld gene cluster from Streptomyces hygroscopicus var. limoneus IFO 12703) (Singh et al., 2006), respectively. Then valienone is phosphorylated to valienone-7-P by a cyclitol kinase (Bai et al., 2006; Singh et al., 2006). Valienone-7-P is the precursor for both cyclitol moieties of validoxamine A, the unglycosylated precursor of validamycin A. In the S. glaucescens pathway for ACB, obviously a third variant is realized, where 2-epi-5-epi-valiolone is first epimerized by an epimerase GacJ and then water is eliminated, and the keto-group at C1 is reduced by the enzyme GacO (dehydratase/dehydrogenase) to give 1-epi-valienol, which is then phosphorylated by GacM to 1-epi-valienol-7-P (Rockser and Wehmeier, 2008). According to the proposals for ACB biosynthesis, the cyclitol is then activated by nucleotidyl-transferases, AcbR or GacR, respectively. The activation of the cyclitol is obviously conserved in a number of C7-cylitol pathways, as related nucleotidyltransferases are also encoded by the gene clusters for the biosynthetic routes for validamycins (Bai et al., 2006; Singh et al., 2006). NDP-1-epi-valienol-7-P is then transferred to a TDP-4-amino-4,6-deoxy-glucose by the glycosyltransferases (AcbI/GacI) to form acarviosine-7-P (Wehmeier and Piepersberg,


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2004; Rockser and Wehmeier, 2008). TDP-4-amino-4,6-deoxy-glucose is synthesized using the standard biosynthetic routes for deoxyglucose. The genes acbA/gacA and acbB/gacB encode the TDP-glucose synthase (EC 2.7.7.24; AcbA/GacA) and the TDP-glucose-4,6-dehydratase (EC 4.2.1.46; AcbB/GacB), respectively. The enzymes are equivalent to StrD/E described in Section 2.2–2.3. The final modification step in this sugar pathway leading to TDP-4-amino-4,6-dideoxy-glucose is an amination catalyzed by AcbV/GacV. The enzymatic activity of AcbV was shown using dTDP-4,6dideoxy-glucose as a substrate (Diaz-Guardamino, M., and Piepersberg, W., unpublished). The enzymatic reaction was carried out in 100 mM Tris-HCl buffer (pH7.5) containing 4 mM dTDP-4-amino-4,6-dideoxy-glucose, 8 mM L-glutamic acid and 1 mM pyridoxalphosphate. The activities can be measured photometrically at g 340 nm in a coupled assay with the a-ketoglutarat dehydrogenase, which converts a-ketoglutarate in the presence of NAD and coenzymeA to succinyl-CoA. A HPLC-system was established by Chung et al. (2007), who analyzed the aminotransferase GerB, which is highly related to AcbV/GacV. The enzyme GerB also was characterized as a dTDP-4-amino-4,6-dideoxy-glucose-aminotransferase. Finally, two glucose moieties are attached to acarviosine-7-P by a second glycosyltransferase (AcbS/GacS). It is still not clear if AcbS/GacS transfer a glucose molecule twice or just a maltose. The last step is the export of ACB-7-P by the ABC exporters AcbWXY or GacWXY, respectively, and unphosphorylated ACB is released (Wehmeier and Piepersberg, 2004; see Fig. 19.3).

3.2. The biochemistry of carbophors: A unique system for the acquisition of glucose from starch in actinomycetes Since ACB is produced as part of a mixture of compounds differing by the number of a-1,4-bound glucose residues at both the reducing and the nonreducing ends, we hypothesize that despite its activity as an inhibitor of a-glucosidases, the benefits of ACB for the producing organism might stem from another function. We proposed that ACB and related compounds might function as ‘‘carbophors’’ (in analogy to the well-known siderophores): these metabolites are produced in order to prevent competitors in the natural habitat from utilizing starch as a carbon and energy source (Zhang et al., 2002; Wehmeier and Piepersberg, 2004). This could be achieved by their function as potent inhibitors of a-glucosidases and of transport systems mediating the uptake of malto-oligosaccharides. Increasing amounts of extracellular, unbound ACB could be used as acceptor molecules for the covalent binding by transglycosylation of glucose or malto-oligosaccharides by the action of the acarviosyltransferase, AcbD, and/or other extracellular a-glucosidases with high transglycosylation potency. The resulting longer homologues would be transported into the

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P P

NDP-valienol-7-P

Acbl

Acarviose-7-P

dTDP-4-amino-4,6-dideoxy-glucose AcbS P

(n)

Acarbose homologs-7-P (n)

P Acarbose-7-P

W

W

F

Glucose

AcbQ

AcbK Carbophor function

2x

X

G

Cytoplasm

Y

H

Out AcbD

Acarbose homologs

Acarbose (inhibitor function)

(n) AcbE, AcbZ Starch

Figure 19.3 Schematic representation of the‘‘carbophor’’ function of ACB.The figure illustrates schematically the biosynthesis and export of ACB, the extracellular modification, the reimport of higher ACB homologues and the intracellular ACB cycle in Actinoplanes sp. SE50/110 (acb cluster, Acb enzymes). The enzymes involved in the respective steps are indicated; for details see text. A similar carbophor cycle is encoded by the gac cluster from S. glaucescens GLA.O (see text).

cytoplasm by a specific importer system, where they would first become phosphorylated by an ACB kinase (AcbK/GacK) and then glucose units would be released and shuttled into metabolism (Fig. 19.3). Free ACB would be released to the environment again by an ACB exporter. The available biochemical data for Acb/Gac-proteins, which provide evidence for and could be involved in the carbophor cycle, are summarized here. 3.2.1. The acarviosyl transferase AcbD The acarviosyl transferase AcbD (Atase, EC2.4.1.19), encoded by the acb cluster of Actinoplanes sp. SE50/110, is a 76-kDa extracellular enzyme from Actinoplanes sp. SE50/110 that transfers the acarviosyl moiety from ACB to the 4-hydroxyl group of various sugars (Hemker et al., 2001; Leemhuis et al., 2004; Wehmeier, U. F., and Rockser, Y., unpublished). ATase has highest similarity to cyclodextrin glucanotransferases (CGTases), e.g., 42% sequence identity to CGTase from Thermoanaerobacterium thermosulfurigenes (P26827). Sequence alignments of AcbD with various a-amylase family enzymes revealed that the four short-sequence regions (I-IV) characteristic of the a-amylase family (Nakamura et al., 1993; Soogard et al., 1993; Uitdehaag et al., 1999) are present in ATase. Therefore, it was assumed


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that the catalytic site architecture of AcbD is similar to those of other a-amylase family enzymes, and that the preference for ACB is probably the result of relatively small differences (e.g., point mutations) in or close to its active site (Leemhuis et al., 2004). It was shown by site-directed mutagenesis that the replacement of Gly140His and/or Gln327His change the specificity of Atase dramatically: The enzyme loses its ACB resistance and Atase function and gains CGTase and a-glucosidase activities (Leemhuis et al., 2004). From these observations, it was assumed that the presence of the His residues mentioned above at these positions is responsible for ACB sensitivity of a-glucosidic enzymes. However, the ACB resistant a-amylases AcbE and AcbZ encoded by the acb gene cluster and also the a-glucosidases and a-amylases encoded by the gac cluster from S. glaucescens GLA.O all contain His residues at these respective positions, and have the active site almost completely conserved when compared to other a-amylases. Thus, the origin of ACB resistance is still unclear. The acbD gene has been cloned and the secreted ATase AcbD was overproduced well in S. lividans strain TK23 when using the E. coli/ actinomycete shuttle vector pUWL201PW-AT (Leemhuis et al., 2004). Acarviosyl transferase activity was found in the culture medium, demonstrating that the signal sequence of ATase is also recognized by S. lividans. The ATase protein can be purified from the culture medium using a standard His-tag purification protocol, yielding about 4 mg of ATase protein per liter of culture (Leemhuis et al., 2004; Rockser and Wehmeier, unpublished). For the expression and purification of AcbD the culture was inoculated by a single colony directly after transformation and grown in Cerestar medium (Cerestar-malto-oligodextrins, 70 g; (NH4)SO4, 5g; yeast extract, 2 g; K2HPO4, 1 g; KH2PO4, 1 g; tri-sodium citrate, 5 g; MgCL2 6H2O, 1 g; FeCL3 6 H2O, 0.25 g; and CaCl2 2 H2O, 2 g; all components dissolved in 1.2 l distilled H2O and sterilized by filtration) for 72 h at 30 on a rotary shaker at 180 rpm. The mycelium was removed by centrifugation and the supernatant was filtered by gravity through glass fibers. Subsequently, 1 ml NiNTA Agarose suspension (Sigma-Aldrich) was added to 100 ml of supernatant and then filled up with Tris-HCl (pH 7.5, final concentration 20 mM ), NaCl (250 mM ), and imidazole (10 mM). This mixture was incubated at 4 for 4 h with gentle agitation. Then the NiNTA-agarose was washed twice with five volumes of buffer 2 (20 mM Tris-Cl, pH 7.5; 250 mM NaCl; 10 mM imidazole; and 1 mM CaCl2). The AcbD protein was eluted with the same buffer containing 250 mM imidazole. AcbD assays are carried out in Tris-HCl buffer (20 mM, pH 7.5) supplemented with 1 mM CaCl2 at 30 . AcbD activity was determined by measuring the transfer rate of the acarviosyl moiety from ACB (donor substrate) to malto-oligosaccharides (acceptor substrates). Reaction mixtures contained 1 to 20 mM ACB, 10 mM malto-oligosaccharides and

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about 10 mg of enzyme per milliliter. Reactions are started by adding enzyme and incubated for 1 to 4 h, then stopped by boiling samples for 5 min. The reactions can be monitored on Silica HPTLC plates (Merck) and should be run twice in acetonitrile/1-propanol/H2O (9/5/7). Visualization of products is obtained by a methanol solution containing 0.3% (w/v) N-(1naphtyl)ethylenediamine and 5% (v/v) H2SO4, followed by heating with a heat gun for a few minutes. ACB, higher ACB homologues, and malto-oligosaccharides can be separated on an HPLC system according to Leemhuis et al. (2004), or using a CarboPac PA200 anion exchange column (250 4 mm) (Dionex, Idstein, Germany) coupled to a CarboPac1 guard column. With eluent A (sodium hydroxide, 0.1 mM) and eluent B (0.6 M sodium acetate in 0.1 M sodium hydroxide) using the following gradient: 0 min, 3% eluent B; 20 min, 30% B; 30 min, 100% B; 30 to 35 min 100% B; 35 min, 3% B; 35 to 47 min 3% B; and 47 min end of run. Detection was performed with an ED50 electrochemical detector (Dionex) with an Au working electrode and an Ag/AgCl reference electrode with a sensitivity of 300 nC. The following pulse program should be used: þ0.1 volt (0 to 0.40 s); to 2.0 volts (0.41 to 0.42 s); þ0.6 volt (0.43 to 0.44 s); and –0.1 volt (0.44 to 0.45 s). 3.2.2. Starch-degrading enzymes Actinoplanes sp. SE50/110 produces two a-amylases (EC3.2.1.1): AcbE (1038 aa) and AcbZ (1103 aa) which represent a new a-glucosidase subfamily (Wehmeier, U. F., Merettig, N., and Rockser, Y., unpublished). They are long-chain acarbose-insensitive a-amylases. The four conserved sequence motifs in the catalytic domain of a-glucosidases and cyclodextrin synthases converting starch and maltodextrins and the three active site amino acid residues, D268, E302 and D375, the ‘‘catalytic triad’’ (Gilles et al., 1996; Nakamura et al., 1993; Soogard et al., 1993; Uitdehaag et al., 1999), are also conserved in AcbE and AcbZ. The activities of a-amylase can be determined using standard a-amylase tests with soluble starch and photometric detection, or the Dionex system described in Section 3.2.1 for AcbD. When using starch as a substrate, the degradation products produced by a-amylases (maltose, maltotriose, or higher malto-oligosaccharides) can be detected. The activities of ACB-insensitive a-amylases are determined in the presence of ACB (0.1 to 1 mM ) and soluble starch (0.1 to 0.5% in the test). With 1 mM ACB AcbE and AcbZ still exhibit more than 70% catalytic activity while ACB-sensitive enzymes are completely inhibited at these concentrations. Four a-glucosidic enzymes are encoded by the gac gene cluster from S. glaucescens GLA.O: GacE1, GacE2, GacZ1, and GacZ2. The two a-amylases GacZ1 and GacZ2 are related to long-chain a-amylases and both also contain a pullulan-binding domain. GacE2 is a short-chain secreted a-glucosidase, while the protein sequence from GacE1 contains


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no signal peptide and seems to be a cytoplasmic enzyme. It remains to be shown if all these enzymes are ACB resistant. These S. glaucescens enzymes all contain the typical active sites found in all a-amylases or starch-degrading enzymes mentioned above and in Section 3.2.1 and, like AcbE and AcbZ from Actinoplanes, no changes are found within these active sites. We have first indications that GacE1 is a cytoplasmic ACB-resistant a-glucosidase and that the extracellular a-amylytic activities detected in the supernatant of S. glaucescens are insensitive toward ACB (Merettig, N., and Wehmeier, U. F., unpublished results). 3.2.3. Binding protein dependent ACB transporters Within the acb and gac cluster putative gene sets (acbHFG, gacFGH) encoding components of ATP-binding cassette (ABC) import systems were found that consist of a membrane-bound solute-binding protein (AcbH/GacH) and two membrane-spanning subunits (AcbFG/GacFG). As often observed for sugar ABC transport systems of Gram-positive bacteria, a gene encoding an ATPase component is lacking (Hurtubise et al., 1995; Schlo¨sser et al., 1999). However, genes encoding ABC proteins (MsiK, multiple sugar import ATPase) (Hurtubise et al., 1995; Schlo¨sser et al., 1997), which are probably part of the transporter complexes, had also been identified in Actinoplanes (Elvers, D., Scha¨fer, G., Brunkhorst, C., and Schneider, E., unpublished) and S. glaucescens (Rockser, Y., and Wehmeier, U. F., unpublished). The homology to members of the CUT 1 (carbohydrate uptake transporter) subfamily, the members of which predominantly transport di- and oligo-saccharides, made the transporters AcbFGH/GacFGH candidates for ACB uptake systems. We have studied this hypothesis (Wehmeier, U. F., in collaboration with Licht, A., Weidlich, D., Scheffel, F., and Schneider, E., Humboldt University Berlin, Germany). We (Brunkhorst et al., 2005) have shown that (1) the product of the acbH gene is induced under the same conditions as other genes of the acb cluster, (2) AcbH is attached to the cytoplasmic membrane, and (3) the purified His10–AcbH fusion protein binds ACB and its longer derivatives but not maltodextrins via surface plasmon resonance. Thus, the substrate binding site of AcbH has apparently evolved in such a way that it can discriminate maltooligosaccharides from pseudo-oligosaccharides. This is in contrast to maltose/maltodextrin binding proteins from E. coli and Alicyclobacillus acidocaldarius, which also bind ACB (Brunkhorst et al., 1999; Hu¨lsmann et al., 2000) and these data strongly support the carbophor model. Comparable data for GacH are still not available. The putative binding protein–dependent ABCimporter GacFGH from S. glaucescens is more related to maltose uptake systems from streptomycetes (45 to 60% identity to the maltose transporter MalFGH from S. coelicolor [NP626480—NP626482]) than to the relatives from Actinoplanes (only 22 to 25% identity). Therefore, it is still speculative which substrates are predominantly transported by the GacFGH-system. As

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a homologue to the Atase AcbD is not encoded by the gac cluster; the strain apparently produces a differently composed mixture of metabolites, which is obviously also reflected in the importer. 3.2.4. Cytoplasmic enzymes involved in ACB metabolism Two proteins are required to close the cytoplasmic ACB cycle, an amylomaltase and an ACB kinase. Proteins for these functions are encoded by the genes acbQ/gacQ and acbK/gacK, respectively, in the two gene clusters investigated. It was first shown by Pape and coworkers that Actinoplanes sp. SE50/110 produces an enzyme phosphorylating ACB at the 7-position and that 7-phospho-ACB does not inhibit the cytoplasmic and ACBsensitive amylomaltase activity of Actinoplanes sp. SE50/110 (Goeke et al., 1996; Drepper and Pape, 1996). The respective enzyme AcbK is encoded by the acb cluster from Actinoplanes sp. SE50/110 and a homologue GacK by the gac cluster from Streptomyces glaucescens GLA.O. The activities of the ACB kinases were determined in assays in a total volume of 20 ml including 10 mM ACB (or ACB homologues synthesized with AcbD, see Section 3.2.1), 10 mM ATP, 10 mM MgCl2, 20 mM NH4CL, 25 mM Tris-HCl (pH 7.6) using the enzymes overproduced in either E. coli BL21(DE3) or JM109 (DE3) (using pET11 or pET16 derivatives) or S. lividans TK23 or TK64 using the pUWL201 (Doumith et al., 2000) expression system. Both enzymes, AcbK and GacK, have been shown to phosphorylate ACB and also higher ACB homologues with similar rates (Rockser and Wehmeier, 2008, and unpublished results). Finally, the additional glucose residues of the 7-phospho-ACB homologues are hydrolyzed by the intracellular amylomaltases AcbQ or GacQ, respectively, being members of glycoside hydrolase family 77 (EC 2.4.1.25). Although both enzymes share high similarity with a-glucanotransferases involved in starch metabolism (e.g., about 60% identical aa residues with 4-alpha-glucanotransferase of Streptomyces coelicolor A3(2); NP_626885), they obviously are members of a new subgroup within this enzyme family and only small changes are responsible for their different substrate specificity. AcbQ was overproduced in E. coli DH5a using the rhamnose-inducible promoter from pJOE2702 (Volff et al., 1996). Preliminary data indicate that phosphorylated higher ACB homologues are hydrolyzed by AcbQ (Merettig, N., Y. Rockser, U. F. Wehmeier, unpublished). For these assays, AcbQ was incubated in buffer (25 mM Tris-HCl, pH 7.5, 10 mM MgCl2) with 10 mM substrate, and the hydrolysis of the substrates was analyzed by TLC according to Hemker et al. (2001). These data fit perfectly with our carbophor model.


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C H A P T E R

T W E N T Y

Biosynthetic Enzymes for the Aminoglycosides Butirosin and Neomycin Fumitaka Kudo* and Tadashi Eguchi† Contents 1. Introduction 2. General Method to Investigate Functions of the Biosynthetic Enzymes for Aminoglycosides 3. Neamine Biosynthetic Enzymes (Enzymes 17, and 20) 3.1. 2-Deoxy-scyllo-inosose (2DOI) synthase 3.2. L-Glutamine:DOI aminotransferase—a dual-function aminotransferase in 2DOS biosynthesis 3.3. 20 -N-Acetylparomamine synthase (UDP-GlcNAc:2DOS N-acetylglucosaminyltransferase) 3.4. 20 -N-Acetylparomamine deacetylase 3.5. FAD-dependent paromamine 60 -dehydrogenase 3.6. 60 -Dehydro-60 -oxoparomamine aminotransferase 4. Ribostamycin Biosynthetic Enzymes (Enzymes 8 and 9) 4.1. Phosphoribostamycin synthase (PRPP: neamine phosphoribosyltransferase) 4.2. Phosphoribostamycin phosphatase 5. Neomycin Biosynthetic Enzymes (Enzymes 57, 10, and 11) (Addition of Neosamine B to Ribostamycin) 5.1. UDP-GlcNAc:ribostamycin N-acetylglucosaminyltransferase 5.2. 2000 -N-Acetyl-6000 -deamino-6000 -hydroxyneomycin C deacetylase (repetitive function of 20 -N-acetylparomamine deacetylase) 5.3. 6000 -Deamino-6000 -hydroxyneomycin C dehydrogenase (probable repetitive function of paromamine 60 -dehydrogenase, uncharacterized) 5.4. 6000 -Deamino-6000 -dehydro-6000 -oxoneomycin C aminotransferase

* {

494 499 500 500 501 504 505 505 506 507 507 508 508 508 509

509 510

Department of Chemistry, Tokyo Institute of Technology, Tokyo, Japan Department of Chemistry and Materials Science, Tokyo Institute of Technology, Tokyo, Japan


#


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494

Fumitaka Kudo and Tadashi Eguchi

6. (S)-4-Amino-2-Hydroxybutyrate Biosynthetic Enzymes (Enzymes 1219) (Addition of AHBA to Ribostamycin) 6.1. g-L-glutamyl-ACP ligase and the pathway-specific acyl carrier protein 6.2. g-L-glutamyl ACP decarboxylase 6.3. g-L-glutamyl-4-aminobutyryl ACP mono-oxygenase and NAD(P)H:FMN oxidoreductase 6.4. g-L-glutamyl-4-amino-2-hydroxybutyryl ACP:ribostamycin g-L-glutamyl-4-amino-2-hydroxybutyryltransferase 6.5. g-L-glutamyl-butirosin B g-L-glutamyl cyclotransferase 7. Other Related Enzymes in the Biosynthesis of 2DOSContaining Aminoglycoside Antibiotics 8. Concluding Remarks and Future Perspectives References

510 511 512 513 514 514 515 516 516

Abstract Butirosin and neomycin belong to a family of clinically valuable 2-deoxystreptamine (2DOS)-containing aminoglycoside antibiotics. The biosynthetic gene clusters for butirosin and neomycin were identified in 2000 and in 2005, respectively. In recent years, most of the enzymes encoded in the gene clusters have been characterized, and thus almost all the biosynthetic steps leading to the final antibiotics have been understood. This knowledge could shed light on the complex biosynthetic pathways for other related structurally diverse aminoglycoside antibiotics. In this chapter, the enzymatic reactions in the biosynthesis of butirosin and neomycin are reviewed step by step.

1. Introduction The aminoglycosides are a well-known important class of clinically valuable antibiotics that interact with bacterial ribosomal RNA and inhibit protein synthesis. The majority of the aminoglycosides contain a unique aminocyclitol 2-deoxystreptamine (2DOS) moiety as the core, which is decorated with various aminosugars, leading to diverse structural molecules, including kanamycin, tobramycin, gentamicin, ribostamycin, butirosin, neomycin, paromomycin, lividomycin, apramycin, and hygromycin B. Other groups of aminoglycosides contain cyclitols either derived from myo-inosotol-1-phosphate or from sedo-heptulose-7-phosphate as reviewed in Chapter 19 of this volume. A number of biosynthetic studies on bioactive secondary metabolites, including the aminoglycosides, have been carried out not only to improve the productivity of the antibiotics and to modify the structures to combat newly emerging resistant pathogenic strains, but also to understand the molecular details of the enzymology that builds the complex structures. Initial incorporation studies and mutational analyses of

Butirosin and Neomycin Biosynthetic Enzymes

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the producer microorganisms suggested plausible biosynthetic pathways. Also, enzymatic analyses with cell-free systems from the producer strains revealed each enzymatic reaction in the biosynthetic pathways. However, because the purification of secondary metabolic enzymes is generally too hard, due to the low amount of proteins in the cells, only a few enzymatic reactions have been confirmed from the native producer microorganisms. The development of molecular genetic techniques in Escherichia coli around 1980 allowed the cloning of biosynthetic genes and their subsequent heterologous expression to obtain the corresponding recombinant proteins. This strategy was applied to the 2DOS biosynthetic studies. In 1999, a key biosynthetic enzyme, 2-deoxy-scyllo-inosose (2DOI) synthase, which catalyzes the first step of 2DOS biosynthesis, was purified from the butirosin producer, Bacillus circulans (Kudo et al., 1999a), and its corresponding gene was subsequently cloned in E. coli (Kudo et al., 1999b). This discovery boosted the rapid identification of many 2DOS-containing aminoglycoside biosynthetic gene clusters, including those for butirosin, neomycin, kanamycin, tobramycin, gentamicin, ribostamycin, paromomycin, lividomycin, apramycin, istamycin, and hygromycin B, because microbial secondary metabolic biosynthetic genes are usually clustered in the genomes of the producing microorganisms. Consequently, during the last decade, most of the enzymes encoded by the butirosin and neomycin biosynthetic gene clusters have been enzymatically characterized, revealing almost all the biosynthetic steps in the pathways. The identification of the aminoglycoside biosynthetic gene clusters and bioinformatic analysis have been summarized in recent excellent reviews (Flatt and Mahmud, 2007; Piepersberg et al., 2007). Biosynthetic studies on 2DOS-containing aminoglycoside antibiotics were also recently reviewed (Llewellyn and Spencer, 2006). Thus, in this chapter, we focus step by step on the enzymatic reactions in the biosynthesis of butirosin and neomycin. Before we go through each enzyme in the biosynthesis of butirosin and neomycin, all biosynthetic enzymes encoded in the gene clusters are summarized in Table 20.1. Comparative analysis of the putative biosynthetic enzymes and chemical structures of 4,5-disubsituted 2DOS aminoglycosides (butirosin and neomycin) and 4,6-disubstituted 2DOS aminoglycosides (kanamycin, tobramycin, and gentamicin), clearly suggested a set of seven conserved enzymes (Table 20.1, enzymes 17) for the biosynthesis of the common neamine structure. Also, four proteins (Table 20.1, enzymes 8, 9, 21, and 22) were specifically encoded in the butirosin and neomycin biosynthetic gene clusters, indicating their involvement in the ribosylation of neamine to give ribostamycin. By comparing the other 4,5-disubsituted 2DOS aminoglycoside (paromomycin and lividomycin) biosynthetic gene clusters, two unique enzymes for neomycin-related structures (Table 20.1, enzymes 10 and 11) and nine unique enzymes for butirosin (Table 20.1, enzymes 1220) were classified with a reasonable degree of certainty.

496 Table 20.1 Butirosin and neomycin biosynthetic enzymes encoded in the gene clusters

Characterized function or putative function

Butirosin biosynthetic enzymes (aa, GI number)

Neomycin biosynthetic enzymes (aa, GI number)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

368 (70720843), BtrC 418 (70720837), BtrS (R) 349a (70720845), BtrE 412c (70720839), BtrM 275 (70720844), BtrD 504 (70720855), BtrQ 432 (70720842), BtrB 604 (70720840), BtrL 213 (70720853), BtrP — — 87 (70720849), BtrI 419 (70720850), BtrJ 428 (70720851), BtrK 341 (70720852), BtrO 205c (70720834), BtrU (V) 302 (70720848), BtrH

430 (62857283) 424 (62857282) 340 (62857281) 421 (62857284) 279 (62857292) 541b(62857287) 416 (62857294) 660 (62857293) 233 (62857289) 366 (62857291) 299 (62857288) — — — — — —

2DOI synthase L-Gln:2DOI aminotransferase 2DOIA dehydrogenase (NADþ dependent) UDP-GlcNAc:2DOS GlcNAc transferase 20 -N-Acetylparomamine deacetylase Paromamine 60 -dehydrogenase L-Gln:60 -Oxoparomamine aminotransferase PRPP:neamine 5-phosphoribosyltransferase 500 -phosphoribostamycin phosphatase UDP-GlcNAc:ribostamycin GlcNAc transferase Neomycin 5000 -epimerase? (radical SAM enzyme) Acyl carrier protein (ACP) g-L-Glu-ACP ligase g-L-Glu-ACP decarboxylase g-L-Glu-GABA-ACP monooxygenase NAD(P)H:FMN oxidoreductase g-L-Glu-AHBA-ACP:ribostamycin g-L-Glu-AHBA transferase

18 19 20 21 22 23 24 25 26 27 28 29 30 a

g-L-Glu-butirosin g-L-glutamyl cyclotransferase Butirosin 300 -epimerase? 2DOIA dehydrogenase (radical SAM enzyme) Unknown Hypothetical protein ABC transporter ABC transporter Aminoglycoside N-acetyltransferase (resistant) ABC transporter ABC transporter Transcriptional regulator Transcriptional regulator Hypothetical protein

156 (70720847), BtrG 232 (70720846), BtrF 250 (70720838), BtrN 1225 (70720841), BtrA 87 (70720854), BtrV — — — 585 (70720856), BtrW 458< (70720857), BtrX 217 (70720833), BtrR1 (W) 380 (70720835), BtrR2 (U) 98c (70720836), BtrT

— — — 1321c (62857295) 83 (62857290) 646c (62857285) 594 (62857286) 287 (62857296) — — — — —

No enzymatic activity was observed, even though several reaction conditions were examined. Another probable repetitive function is not detected. c Several ORFs are deposited as different sizes. Notes: Uncharacterized proteins are indicated as italic and underlined. The hyphens indicate no homological ORFs encoded in the gene clusters. The symbols of ORFs encoded in the neomycin biosynthetic gene cluster were omitted to prevent confusion, since many same gene products are deposited as different symbols in the database. As one of reference, the GI numbers are listed from the accession code AB097196 for butirosin biosynthetic gene cluster (btr) and AB211959 for neomycin biosynthetic gene cluster (neo), which we deposited. G6P, D-glucose-6-phosphate; 2DOI, 2-deoxy-scyllo-inosose; 2DOIA, 2-deoxy-scyllo-inosamine; amino-DOI, 3-amino-2,3-dideoxy-scyllo-inosose; 2DOS, 2-deoxystreptamine; Gln, L-glutamine; Glu, L-glutamate; GABA, g-aminobutyrate; AHBA, (2S)-4-amino-2-hydroxybutyrate; ACP, acyl carrier protein; UDP, uridine 50 -diphosphate; GlcNAc, N-acetyl-D-glucosamine; PRPP, 5-phosphoribosyl-1-diphosphate;. NAD(H), nicotinamide adenine dinucleotide (and its reduced form); FAD(H2), flavin adenine dinucleotide(and its reduced form); FMN(H2), flavin mononucleotide(and its reduced form). b

497

498


Considering the putative functions from homology analysis with known functional enzymes, the enzymatic functions in each biosynthetic step were plausibly presumed and the expected enzymatic reactions were investigated with the recombinant proteins expressed in E. coli. Consequently, the overall biosynthetic pathways for butirosin and neomycin have been almost completely understood, as shown in Fig. 20.1, except for the epimerizations at the last steps and an oxidation reaction of 6000 -deamino-6000 -hydroxyneomycin C indicated as broken arrows. Although probable ABC transporters and regulatory proteins are not characterized, it is noteworthy that most of the proteins encoded in the gene clusters were found to be required for the construction of the relevant molecules. We also note that relatively large numbers of enzymes are repetitively used in the biosynthesis of the aminoglycosides. This fact may

2-

Common pathway

OPO3 O

HO HO

1

HO D-glucose-6-phosphate (G6P)

HO HO

5

AcHN H2N O HO

NH2 OH 2⬘-N-acetylparomamine

UDP-GIcNAc

2-deoxy-scyllo-inosamine (2DOIA)

HO HO

6

H2N HO HO

2

3-amino-2,3-dideoxyscyllo-inosose (amino-DOI) H

OH O H2N H2N O HO

NH2

O O H2N H2N O HO

NH2 HO

2-deoxystreptamine (2DOS)

HO HO

7 NH2

NH2 O H2N O HO

H2N

OH

Neamine

NH2 OH

NH2 O

HO HO

O

H2N H2N O NH2 O 2-O PO 3 O OH

8

H2N HO HO HO O

3 or 20 OH HO

OH Paromamine

NH2 HO HO

H2N HO HO

2 OH HO

2-deoxy-scyllo-inosose (2DOI)

OH O

HO HO

4

O

HO HO

OH

9 HO

H2N H2N O NH2 O O OH

PRPP HO OH Ribostamycin

HO OH

In S. fradiae

Ribostamycin

NH2 NH2 NH2 NH2 O O O O HO HO HO HO HO HO H2N H2N H2N H2N H2N H2N H2N H2N OO OO OO OO NH NH NH2 NH 2 7 2 6? HO 2 HO HO 11? HO O O O O OH OH OH OH

NH2 O H2N H2N OO NH2 HO O OH

HO HO

HO HO 10

O OH

O OH O

UDP-GlcNAc

5

OH OH

AcHN

OH

In B. circulans

H2N

OH

NH2 H2N H2N H O O N HO O OH

17

g -L-Glu-AHBA-ACP

13

NH+ 3 S

-O C 2

O L-Glu

OH

O OH

H2N

NH2 OH

O

OH

O

CO2-

N H

O

18

NH+ 3

NH2 H2N

H2N OO HO O

S

+H

3N

ACP

O

13

O

O HO HO H2N OH H2N 19? H OO N HO NH2 O OH OH O

Butirosin B

OH

O -O C 2

NH+ 3 L-Glu

OH

H N OH

HO OH

14 ACP

NH2

O

OH OH Neomycin B

H2N

OH Neomycin C

O

HO HO

HO OH

12 ACP-SH

O OH

O OH

O

NH2

O

HO HO Ribostamycin

O OH OH OH

O

H2N

N H

S O

15 and 16 ACP

O2

O

-O

2C

NH2

Butirosin A

OH N H

S

ACP

O NH3+ g -L-Glu-AHBA-ACP

Figure 20.1 Biosynthetic pathways for neomycin (in Streptomyces fradiae) and butirosin (in Bacillus circulans). The solid arrows indicate the enzymatically characterized reaction steps and the broken arrows show the uncharacterized reaction steps. The numbers over the arrows correspond to the enzymes listed inTable 20.1.The shunt pathways and reverse reactions are not shown.


499

mean that the producer microorganisms somehow minimize the set of enzymes to save energy or for the purposes of regulation. We now show each biosynthetic enzyme involved in the biosynthesis of butirosin and neomycin. Here, the biosynthetic steps are divided into those for neamine formation, ribostamycin formation, neomycin formation, and AHBA biosynthesis in the butirosin pathway. Due to space limitations, we omit mechanistic insights into the enzyme reactions and will discuss these issues elsewhere.

2. General Method to Investigate Functions of the Biosynthetic Enzymes for Aminoglycosides Chromosomal DNAs are first prepared from the cells of the producer microorganisms according to standard protocols such as those in Practical Streptomyces Genetics (Kieser et al., 2000). The aminoglycoside-producing microorganisms are often purchased from a stock center such as NBRC, JCM, or ATCC. Information on many aminoglycoside antibiotic biosynthetic genes is now widely available from public databases. Target genes are next amplified by standard PCR with designed primers having appropriate restriction sites and chromosomal DNA as a template. Usually, high-fidelity DNA polymerases such as PrimeSTAR HS DNA polymerase (TAKARA) and KOD plus DNA polymerase (TOYOBO) are used to obtain target DNA fragments. The amplified target DNAs are subcloned once into a commonly used standard plasmid vector such as pUC18, pT7Blue or pLITMUS28 according to a standard protocol (Sambrook et al., 2001). After confirmation of the sequences of the cloned DNAs, the target genes are inserted into the appropriate site of a suitable expression plasmid such as pET30, pET28 or pColdI. The expression plasmids with target genes are then introduced into an appropriate E. coli host such as BL21(DE3) for overexpression of the genes. The culture conditions (temperature, concentration of inducer compound such as isopropyl-b-D-thiogalactoside (IPTG), and media) depend on the target proteins. The E. coli cells expressing the target proteins are homogenized by use of a French Pressure Cell Press or sonic homogenizer to obtain the cell-free extract containing the recombinant proteins. His-tagged recombinant proteins are purified with Ni or Co affinity (such as Ni-NTA and TALON) chromatography. Also, conventional purification methods are applicable. Preparation of appropriate substrates is usually a difficult problem. Degradation of the parent antibiotics and chemical synthesis have been generally used up to now. However, once the sequenced biosynthetic enzymes have been functionally characterized, enzymatic synthesis could be used for the purpose.

500


The enzyme reaction products could be detected by LC-ESI-MS directly or HPLC after derivatization, for example, with dansyl chloride or dinitrofluorobenzene, which attach UV-absorbing functional groups to the amino groups. The enzyme reaction products can also be purified by cation exchange chromatography. After the salt forms of the aminoglycosides have been exchanged for hydrochloride or sulfate, NMR analysis should be performed. See original papers for the details of the experiments in this section.

3. Neamine Biosynthetic Enzymes (Enzymes 17, and 20) 3.1. 2-Deoxy-scyllo-inosose (2DOI) synthase 2-Deoxy-scyllo-inosose (2DOI) synthase catalyzes the first step in the biosynthesis of 2-deoxystreptamine (2DOS). The conversion of D-glucose6-phosphate (G6P) to 2DOI was first observed in a cell-free extract of neomycin-producing S. fradiae (Yamauchi and Kakinuma, 1992). The enzymatic conversion was then found to be an NADþ-dependent reaction. The corresponding enzyme, 2DOI synthase, was purified from the butirosin producing B. circulans SANK72073 (Kudo et al., 1999a). The native 2DOI synthase derived from B. circulans was isolated as a heterodimer, composed of a 40-kDa subunit and a 20-kDa subunit. The corresponding genes, btrC and btrC2, were separately cloned by reverse genetic methodology on the basis of the N-terminal amino acid sequences of the native enzyme (Kudo et al., 1999b; Tamegai et al., 2002a) Eq. (20.1). OPO32– HO HO

O

OH

HO D-glucose-6-phosphate (G6P)

2-deoxy-scyllo-inosose synthase NAD+ Co2+

O HO HO

OH HO 2-deoxy-scyllo-inosose (2DOI)

Pi

ð20:1Þ

The encoded protein sequences for the large subunit (BtrC) showed significant similarity to dehydroquinate (DHQ) synthase, which catalyzes carbocyclic formation from 3-deoxy-D-arabino-heptulosonate-7-phosphate in the shikimate pathway. The recombinant BtrC is composed of 368 amino acids, 40,746 Da and has 2DOI synthase activity with NADþ and Co2þ. The optimum pH is 7.5 to 8.0. Under the optimum reaction conditions (pH 7.7 and 46 C), the kinetic constants, kcat 1.0 s–1, Km (G6P) 2.1 10–4 M, Km (NADþ) 2.3 10–5 M, were determined. Except for the kinetic constants and stability, the biochemical properties are the same as those of the native heterodimeric enzyme (kcat 7.3 10–2 s–1, Km (G6P) 9.0 10–4 M, Km (NADþ) 1.7 10–4 M, at pH 7.7, 46 C). In addition


501

to G6P, 2-deoxy-G6P, 3-deoxy-G6P, 2-fluoro-G6P (Km 2.1 10–3 M) and 3-amino-G6P (Km 1.4 10–3 M) are accepted as substrates, although 4-deoxy-G6P, 2-amino-G6P and 3-fluoro-G6P were not accepted (Eguchi et al., 2002; Iwase et al., 1998). The crystal structure of BtrC was solved and showed significant similarity to DHQ synthase (Nango et al., 2008). Extensive mechanistic studies on 2DOI synthase have also been carried out and its similarity and dissimilarity to DHQ synthase were clearly elucidated (Hirayama et al., 2007; Huang et al., 2005b). The 2DOI synthases derived from the neomycin producer S. fradiae NBRC12773 (Kudo et al., 2005), the tobramycin/apramycin producer Streptoalloteichus hindustanus JCM3268 (Hirayama et al., 2006), the ribostamycin producer S. ribosidificus ATCC21294 (active enzyme was only expressed in S. lividans) (Subba et al., 2005), the kanamycin producer S. kanamyceticus ATCC12853 (Kharel et al., 2004b), and the tobramycin producer S. tenebrarius ATCC17920 (Kharel et al., 2004a) were shown to have activity. The kinetic constants for 2DOI synthase from S. hindustanus JCM3268 were found to be kcat 0.075 s–1, Km (G6P) 6.9 10–4 M (Hirayama et al., 2006). Among the 2DOI synthases, BtrC seems to be the most active enzyme thus far characterized. The 2DOI synthase homologous gene is a useful biomarker for the identification of the biosynthetic gene clusters for 2DOS-containing aminoglycoside antibiotics. 2DOI itself is thought to be a valuable material for the production of fine chemicals such as carbaglucose and aromatic compounds (Kakinuma et al., 2000). Thus, the 2DOI production system using the 2DOI synthase in E. coli has been greatly improved (Kogure et al., 2007). The small subunit of 2DOI synthase from B. circulans (BtrC2) was found to be an L-glutamine amidotransferase homologue involved in the newly identified pyridoxal-50 -phosphate (PLP) pathway in Bacillus (Burns et al., 2005). This small subunit suppresses the 2DOI synthase activity but significantly stabilizes the protein complex. Thus, it was speculated that this small subunit controls the 2DOI synthase activity in the cells of microorganisms to keep a certain concentration of the aminoglycoside and PLP, which is required for many transamination reactions in aminoglycoside biosynthesis.

3.2. L-Glutamine:DOI aminotransferase—a dual-function aminotransferase in 2DOS biosynthesis The two transaminations with L-glutamine as an amino donor in 2DOS biosynthesis Eqs. (20.2) and (20.3) were extensively studied by the Walker group with the native enzymes derived from the producer strains. They found that the transamination activity could not be separated, indicating that these two transaminations were catalyzed by the same enzyme in the strains (Lucher et al., 1989).

502


L-glutamine:

2-deoxy-scyllo-inosose aminotransferase∗

O HO HO

OH

PLP L-glutamine

HO

H2N HO HO

L-glutamine:

2-deoxy-scyllo-inosose aminotransferase∗

HO O

PLP L-glutamine

∗same enzyme

H2N HO HO

2-ketoglutaramate OH HO 2-deoxy-scyllo-inosamine (2DOIA)

ð20:2Þ

H2N HO 2-ketoglutaramate NH2 HO HO 2-deoxystreptamine (2DOS)

ð20:3Þ

Among the biosynthetic genes for butirosin, one open reading frame (ORF), BtrS (also called BtrR), showed significant homology to L-glutamine:scyllo-inosose aminotransferase in the biosynthesis of streptomycin (Ahlert et al., 1997), indicating that it could catalyze the amination of 2DOI to give 2-deoxy-scyllo-inosamine (2DOIA). In 2002, BtrS was found to be L-glutamine:2DOI aminotransferase and also the reverse reaction from 2DOS with pyruvate as an amino acceptor was detected, indicating that BtrS catalyzes two transaminations in DOS biosynthesis (Huang et al., 2002; Tamegai et al., 2002b). The structure of the reverse reaction product from 2DOS was confirmed as chiral 3-amino-2,3-dideoxy-scyllo-inosose (aminoDOI), indicating stereospecific recognition of two enantiotopic amino groups in 2DOS (Yokoyama et al., 2005). Substrate recognition studies showed that the a-positions of the ketones of the substrates are not so important, but functional groups at the b-position are critical for recognition by this dual-function enzyme. The forward reaction of the second transamination was also confirmed with the neomycin biosynthetic enzyme (Kudo et al., 2005). The crystal structure of this aminotransferase derived from B. circulans was solved and showed the tertiary structure of the enzyme, including its active site with PLP or PMP cofactors (Popovic et al., 2006). The substrate-docking model displayed some possible recognition amino acid residues, but the precise recognition mechanism of the distinguishable enantiotopic amino groups of 2DOS on the amino acid level is still unclear. Thus, detailed studies, including mutational studies, are needed to understand the recognition mechanism. In streptomycin biosynthesis, two different aminotransferases are known to be involved in the construction of the streptidine moiety (Ahlert et al., 1997; Walker and Walker, 1969). Thus, the recognition mechanism of this family of aminotransferase is quite interesting. The homologous aminotransferase derived from the neomycin producer S. fradiae NBRC12773 (Huang et al., 2005a; Kudo et al., 2005), and the tobramycin producer S. tenebrarius ATCC17920 (only the first transamination) (Kharel et al., 2005), have been shown to have the activity.

503


3.2.1. NAD-dependent 2-deoxy-scyllo-inosamine dehydrogenase This enzymatic reaction Eq. (20.4) was initially observed with a cell-free system from the neomycin producer S. fradiae and found to be an NADþdependent reaction (Suzukake et al., 1985). Thus, an NADþ-dependent dehydrogenase encoded in the biosynthetic gene cluster was postulated in this oxidation reaction. Only one NADþ-dependent dehydrogenase homologous protein is conserved in the biosynthetic gene clusters for similarly structured aminoglycoside antibiotics. The corresponding ORF from the neomycin producer S. fradiae NBRC12773 was then expressed in E. coli and the expected NADþ-dependent oxidation of 2DOIA was observed in the presence of zinc ion (Kudo et al., 2005). The product, amino-DOI, was indeed converted to 2DOS by the dual-function aminotransferase as mentioned above. However, this recombinant enzyme has not been purified yet to investigate the specific recognition mechanism. Although no other enzyme has been characterized so far, the other actinomycete enzymes seem to catalyze this oxidation reaction. Heterologous expression of several genes for kanamycin and gentamicin biosynthesis in Streptomyces clearly revealed that this type of NADþ-dependent dehydrogenase is involved in the biosynthesis (Kurumbang et al., 2008; Park et al., 2008). H2N HO HO

2-deoxy-scyllo-inosamine dehydrogenase OH HO

NAD+ in Streptomyces fradiae

H2N HO HO

HO O 3-amino-2,3-dideoxyscyllo-inosose (amino-DOI)

NADH

ð20:4Þ

On the other hand, the corresponding enzyme derived from B. circulans did not show the activity at all, since the enzyme lacks the catalytically important zinc-binding motif, which is required for the activity of this class of dehydrogenase. So, the true function of the protein in B. circulans is unclear at the moment. It may be just a relic during evolution or may function at another biosynthetic step, such as the epimerization at C-3’’ of butirosin. 3.2.2. Radical SAM 2-deoxy-scyllo-inosamine dehydrogenase A radical SAM dehydrogenase was found to catalyze the oxidation of 2DOIA to give amino-DOI in the biosynthesis of butirosin in B. circulans Eq. (20.5) (Yokoyama et al., 2007). It was really unprecedented that this type of enzyme should be involved in 2DOS biosynthesis. The enzyme BtrN encoded in the butirosin biosynthetic gene cluster possesses a CXXXCXXC motif conserved within the radical S-adenosyl-L-methionine (SAM) superfamily, which is an emerging group of enzymes that catalyze a wide range of radical reactions (Wang and Frey, 2007). The common mechanism found in all radical SAM enzymes is a reductive

504


cleavage of SAM to generate the 50 -deoxyadenosyl radical, which is subsequently used for radical reactions such as activation of other enzymes through generating a protein radical, sulfur insertion, C–C bond rearrangement, and amino group rearrangement. Disruption of btrN in the butirosin producer B. circulans blocked the biosynthetic pathway between 2DOIA and 2DOS. Further, in vitro assay of the overexpressed enzyme revealed that BtrN catalyzed the oxidation of 2DOIA under strictly anaerobic conditions along with consumption of an equimolar amount of SAM to produce 50 -deoxyadenosine, methionine, and amino-DOI. The BtrN reaction with [3-2H]2DOIA generated deuterated 50 -deoxyadenosine, while no deuterium was incorporated by incubation of unlabeled DOIA in the deuterium oxide buffer. These results indicated that the hydrogen atom at C-3 of 2DOIA was directly transferred to 50 -deoxyadenosine to give the radical intermediate of 2DOIA. The 2DOIA radical intermediate was also detected by EPR spectroscopy, clearly revealing the radical reaction mechanism (Yokoyama et al., 2008a). H2N HO HO HO

2-deoxy-scyllo-inosamine dehydrogenase OH

S-adenosyl-L-methionine in Bacillus circulans

H2N HO HO

HO O

+ methionine and 5´-deoxyadenosine

ð20:5Þ

3.3. 20 -N-Acetylparomamine synthase (UDP-GlcNAc:2DOS N-acetylglucosaminyltransferase) This glycosyltransfer reaction was presumed to be catalyzed by a retention-type of glycosyltransferase with nucleotidylyl sugars Eq. (20.6). Initially, NDPglucosamine was thought to be a sugar donor in paromamine synthesis. However, the existence of the following pathway-specific deacetylase (Truman et al., 2007) indicated that UDP-N-acetyl-D-glucosamine (UDPGlcNAc) is a most likely sugar donor. In the butirosin and neomycin biosynthetic gene clusters, only one obvious glycosyltranferase is encoded, suggesting that this type of enzyme catalyzes the glycosyl transfer reaction at this step. Although B. circulans-derived recombinant enzyme was not obtained as a soluble protein (unpublished), S. fradiae–derived recombinant enzyme was obtained in soluble form by coexpression with GroEL and GroES. The expected enzymatic reaction with 2DOS and UDP-GlcNAc in the presence of Mg2þ showed the production of 2’’-N-acetylparomamine, which was clearly confirmed by NMR (Yokoyama et al., 2008b). This enzyme has unfortunately not yet been purified to investigate the precise substrate specificity. Thus far, the impure enzymatic solution showed high specificity to UDPGlcNAc over UDP-glucose, suggesting a specific GlcNAc transfer enzyme.

505


H2N

HO HO

HO

NH2

UDP-N-acetylglucosamine: 2-deoxystreptamine N-acetylglucosaminyltransferase HO (2´-N-acetylparomamine synthase) HO UDP-GlcNAc Mg2+

OH O H2N O HO

+ UDP

AcHN

NH2

OH 2´-N-acetylparomamine

ð20:6Þ 3.4. 20 -N-Acetylparomamine deacetylase The functional characterization of this 20 -N-acetylparomamine deacetylase Eq. (20.7) was achieved with B. circulans- and S. fradiae–derived recombinant enzymes (Fan et al., 2008; Truman et al., 2007; Yokoyama et al., 2008b). Also, this enzyme was found to catalyze the deacetylation of 2000 -N-acetyl-6000 -deaminoneomycin C in the biosynthesis of neomycin (Fan et al., 2008; Yokoyama et al., 2008b). Thus, this family of enzyme seems to recognize only the N-acetylglucosamine moiety over the other moieties in aminoglycosides. This enzyme does not require any cofactor for its activity. Under neutral pH between 7.0 and 8.0, removal of the acetyl group from N-acetylated substrates is catalyzed. HO HO

OH O H2N O HO

2´-N-acetylparomamine deacetylase

AcHN

NH2 OH

HO HO

OH O H2N + acetic acid O NH2 HO OH Paromamine

H2N

ð20:7Þ

The gentamicin biosynthetic gene cluster was initially identified without this deacetylase homologous ORF, but recent reanalysis of the sequence revealed a deacetylase homologue, and the involvement of the gene in the biosynthesis was clearly confirmed by heterologous expression of all the enzymes required for gentamicin A2 construction (Park et al., 2008). Thus, a sequential pathway from G6P to paromamine has been verified as a common pathway to this class of antibiotics.

3.5. FAD-dependent paromamine 60 -dehydrogenase The remaining two steps to the most advanced common intermediate, neamine, in the 2DOS-containing aminoglycosides were presumed to be sequential oxidation and transamination reactions Eqs. (20.8) and (20.9). From the comparison of the gene clusters, two completely conserved common ORFs were found, encoding a FAD-dependent dehydrogenase and a PLP-dependent class III aminotransferase. The S. fradiae–derived

506


dehydrogenase was expressed in E. coli and the presence of the FAD cofactor in the protein was confirmed (Huang et al., 2007). The expected oxidation reaction of paromamine was successfully detected under aerobic conditions. On the other hand, removal of oxygen by flushing with argon gas caused a significant reduction in enzymatic activity, indicating reoxidation of the reduced FAD cofactor with molecular oxygen for the next catalytic cycle. The aldehyde product was converted to neamine by the following aminotransferase, suggesting that this FAD-dependent enzyme is involved in the oxidation step in neamine formation. This enzyme did not exhibit oxidation activity toward the possible biosynthetic intermediates UDP-GlcNAc, 20 -Nacetylparomamine, and paromomycin indicating the specific recognition of the paromamine molecule. However, this enzyme is presumed to catalyze the oxidation at C-6000 of 6000 -deamino-6000 -hydroxyneomycin C. In fact, our preliminary study of the corresponding enzymatic reaction showed the expected activity. This family of enzyme is encoded in the paromomycin biosynthetic gene cluster, although the paromamine moiety remains in the final product. Thus, the recognition mechanism of this possibly dual functional FAD-dependent dehydrogenase is interesting. HO HO

OH O H2N H2N O HO

Paromamine 6´-dehydrogenase

OH

NH2

FAD

H HO HO

O O H2N O HO

H2N

OH

NH2

ð20:8Þ Inactivation of the corresponding gene in tobramycin biosynthesis caused the production of 60 -deamino-60 -hydroxy-kanamycin C, supporting the idea that this enzyme is involved in the oxidation reaction in the neamine moiety of in the biosynthesis of kanamycin/tobramycin (Yu et al., 2008).

3.6. 60 -Dehydro-60 -oxoparomamine aminotransferase As mentioned above, this enzymatic activity was observed by coupled transformation from paromamine with the FAD-dependent enzyme and the PLP-dependent aminotransferase Eq. (20.9) (Huang et al., 2007). This aminotransferase exhibited the characteristic absorptions for enzyme-bound pyridoxal-50 -phosphate (PLP) and pyridoxamine-50 -phosphate (PMP). This aminotransferase preferred to use L-glutamine and L-glutamate as an amino donor in this reaction; L-leucine, L-lysine, L-arginine, L-aspartate, and S-adenosylmethionine were also accepted with reduced efficiency (Huang et al., 2007). The B. circulans-derived aminotransferase preferred to use L-glutamine rather than L-glutamate (unpublished).

507


H HO HO

O O H2N O HO

L–glutamine:6´-oxoparomamine aminotransferase

H2N

OH

NH2

HO HO

PLP L-glutamamine

NH2 O 2-ketoglutaramate H N H2N 2 O NH2 HO OH

ð20:9Þ

Neamine

In the gentamicin biosynthetic gene cluster, a total of four homologous aminotransferases are encoded and might be responsible for the aminotransfer reactions at specific carbons. So, precise characterization of substrate specificity of this type of aminotransferase should be important.

4. Ribostamycin Biosynthetic Enzymes (Enzymes 8 and 9) 4.1. Phosphoribostamycin synthase (PRPP: neamine phosphoribosyltransferase) Four conserved ORFs among the butirosin, neomycin, ribostamycin, paromomycin, and lividomycin biosynthetic gene clusters were presumed to be involved in the transformation of neamine to ribostamycin Eq. (20.10). All of the enzymes were expressed in E. coli, and all combination of the enzymes were examined to detect the expected ribosylation reaction with 5-phosphoribosyl-1-diphosphate as a possible ribosyl donor. As a result, one of the proteins was characterized as 5-phosphoribosyl-1-diphosphate:neamine 5-phosphoribosyltransferae and another was found to be 5’’-phosphoribostamycin phosphatase to give ribostamycin (Kudo et al., 2007). The other two proteins remain to be characterized, although they may be involved in the ribostamycin related chemistry in the biosynthesis. NH2 O HO HO H2N H2N O HO

5-phosphoribosy-1-diphosphate:neamine 5-phosphoribosyltransferase

OH

NH2

PRPP Mg2+

NH2 O H2N H2N O NH2 2–O PO O 3 O OH HO HO

PPi

ð20:10Þ

HO OH

Only the B. circulans–derived enzyme has been functionally characterized so far. Due to the instability of the protein, the substrate specificity was investigated with cell-free extracts of E. coli expressing this enzyme. As a result, paromamine was accepted as a substrate, but not kanamycin A. Thus, this enzyme seems to recognize the pseudodisaccharide structure of

508


2DOS-containing aminoglycosides. The enzyme was also found to have optimum catalytic activity at pH 7.5, at a temperature between 27 and 32 C, and required an essential divalent metal ion: optimally Mg2þ, Ni2þ, or Co2þ. However, further research is needed to solve the precise chemistry of this unique 5-phosphoribosyltransferase.

4.2. Phosphoribostamycin phosphatase As mentioned above, this enzymatic reaction was detected with a coupled reaction with PRPP:neamine 5-phosphoribosyltransferase (Kudo et al., 2007) Eq. (20.11). Only the B. circulans–derived enzyme has been characterized so far. The recombinant enzyme was purified and no metal requirement for the phosphatase activity was observed. The substrate specificity of the enzyme was also investigated with PRPP, ribose 5-phosphate, fructose 1,6-bisphosphate, fructose 6-phosphate, AMP, GMP, CMP, UMP, and TMP in addition to p-nitrophenyl phosphate, since authentic 500 -phosphoribostamycin was not available. p-Nitrophenyl phosphate was well hydrolyzed by this enzyme, and fructose 1,6-bisphosphate was also hydrolyzed, although the regiospecificity was not elucidated. The other possible substrates including PRPP were not hydrolyzed by this phosphatase at all. More studies are needed to solve the recognition mechanism of 500 -phosphoribostamycin phosphatase. HO HO

NH2 O HN H2N 2 OO O

2–O PO 3

5˝-phosphoribostamycin phosphatase NH2 OH

HO OH

NH2 O H2N H2N OO NH2 HO O OH

HO HO

HO OH Ribostamycin

Pi

ð20:11Þ

5. Neomycin Biosynthetic Enzymes (Enzymes 57, 10, and 11) (Addition of Neosamine B to Ribostamycin) 5.1. UDP-GlcNAc:ribostamycin N-acetylglucosaminyltransferase As shown in Table 20.1, two unique ORFs were identified in the neomycin, paromomycin, and lividomycin biosynthetic gene clusters. Among them, one ORF was found to encode a radical SAM enzyme, which was presumed not to be involved in the glycosylation event. Another ORF was found to show low similarity to some glycosyltransferases, especially at the C-terminal end. Thus, the enzyme derived from the neomycin producer S. fradiae was expressed in

509


E. coli and the glycosylation of ribostamycin with UDP-GlcNAc Eq. (20.12) was examined (Yokoyama et al., 2008b). The product, 2000 -N-acetyl-6000 deamino-6000 -hydroxyneomycin C, was clearly confirmed by NMR. This enzyme specifically recognized ribostamycin as glycosyl acceptor rather than 2DOS, paromamine, 20 -N-acetylparomamine, and neamine, and UDP-GlcNAc as glycosyl donor rather than UDP-Glc. This enzyme requires an essential divalent metal ion: optimally Mg2þ, Mn2þ, or Co2þ. NH2 O HO HO HN H2N 2 O NH2 HO OO OH HO OH

UDP-N-acetylglucosamine:ribostamycin N-acetylglucosaminyltransferase UDP-GlcNAc Mg2+

NH2 O HN H2N 2 O NH2 O HO O OH

HO HO

UDP

ð20:12Þ

O OH OH O AcHN OH OH 2˝Ń-acetyl-6˝´-deamino-6˝´-hydroxyneomycin C

5.2. 2000 -N-Acetyl-6000 -deamino-6000 -hydroxyneomycin C deacetylase (repetitive function of 20 -N-acetylparomamine deacetylase) In the neomycin-related antibiotic biosynthetic gene clusters, only one obvious deacetylase functioning as 20 -N-acetylparomamine deacetylase is encoded, suggesting the repetitive use of the deacetylase Eq. (20.13). As expected, the enzyme also catalyzed the deacetylation of 2000 -N-acetyl-6000 deamino-6000 -hydroxyneomycin C to give 6000 -deamino-6000 -hydroxyneomycin C, which was confirmed by NMR (Yokoyama et al., 2008b). Thus, the enzyme seems to recognize only the N-acetylglucosamine moiety in aminoglycosides. NH2 O HO HO H N H2N 2˝´-N-acetyl-6˝´-deamino-6˝´-hydroxyneomycin C 2 O NH2 deacetylase HO O O OH

NH2 O H2N H2N O NH2 HO O O OH

HO HO

*Same enzyme as 2´-N-acetylparomamine deacetylase

O OH OH O OH AcHN OH

O OH OH O OH H2N OH

acetic acid

ð20:13Þ

5.3. 6000 -Deamino-6000 -hydroxyneomycin C dehydrogenase (probable repetitive function of paromamine 60 -dehydrogenase, uncharacterized) This oxidation reaction Eq. (20.14) was presumed to be catalyzed by paromamine 60 -dehydrogenase, since the reaction scheme appears to be similar to the oxidation of C-6 of the glucosamine moiety, and also other candidates are not found in the gene clusters. In fact, our preliminary study showed the oxidation activity of the enzyme (unpublished).

510

Fumitaka Kudo and Tadashi Eguchi NH2 O

HO HO

HN H2N 2 OO NH2 HO O OH

NH2 O

HO HO HO

6˝´-deamino-6˝´-hydroxyneomycin C 6˝´-dehydrogenase

H2N H2N OO O

OH

FAD

O OH H2N

ð20:14Þ

O OH OH OH

O

O OH

O

H2N

OH

NH2

OH

5.4. 6000 -Deamino-6000 -dehydro-6000 -oxoneomycin C aminotransferase This reverse reaction Eq. (20.15) was confirmed to be catalyzed by 60 -oxoparomamine aminotransferases derived from S. fradiae and B. circulans (Huang et al., 2007). Our preliminary result showed a sequential oxidation and transamination of 6000 -deamino-6000 -hydroxyneomycin C to neomycin C (unpublished). We are now in the process of confirming the structure of the product as neomycin C, not neomycin B, since an epimerization reaction was suspected during the sequential enzymatic reaction. HO HO

NH2 O H2N

HO

H2N OO NH2 O OH

NH2 O H2N H2N OO HO O

HO HO L-glutamine:6˝´-oxoneomycin C aminotransferase∗

OH

NH2 + 2-ketoglutaramate

PLP L-gluatamine

O OH H2N

O OH

O OH

∗ Same enzyme as L-glutamine: 6˝´-oxoparomamine aminotransferase

O OH H2N

O

NH2 OH

ð20:15Þ

OH Neomycin C

The last step of neomycin B biosynthesis appears to be the epimerization reaction at C-5000 of neomycin C to neomycin B, although the timing of the epimerization is not clear at present. A putative radical SAM enzyme, which is also encoded in the paromomaycin and lividomycin biosynthetic gene clusters, remains to be solved. This protein may catalyze the epimerization at C-5000 of neomycin C with a radical mechanism. Functional analysis of this enzyme derived from S. fradiae is in progress.

6. (S)-4-Amino-2-Hydroxybutyrate Biosynthetic Enzymes (Enzymes 1219) (Addition of AHBA to Ribostamycin) The (S)-4-amino-2-hydroxybutyryl (AHBA) moiety at 1-N of the 2DOS of butirosin is a structurally unique feature. It is well known that AHBA attachment on kanamycin A and dibekacin lead to the clinically valuable amikacin and arbekacin (Kondo and Hotta, 1999). Labeling studies

511


to seek the attachment of AHBA with several amino acids such as L-glutamate, g-aminobutyric acid (GABA), and AHBA were performed, showing that L-glutamate and GABA are direct precursors to the AHBA moiety of butirosin but not AHBA itself (Yukita et al., 2003). Thus, a certain activation reaction of the carboxyl group was presumed before the loading of the amino acid substituent. The butirosin biosynthetic gene cluster contains seven unique contiguous genes compared with the neomycin biosynthetic gene cluster (Table 20.1, 1215, 1719). The Spencer group expressed six of the seven enzymes and one probable NADH:FAD oxidoreductase encoded in the butirosin biosynthetic gene cluster and elegantly figured out the unique AHBA biosynthetic pathway and transfer reaction onto ribostamycin to give butirosin, as described in this section (Li et al., 2005; Llewellyn et al., 2007).

6.1. g-L-glutamyl-ACP ligase and the pathway-specific acyl carrier protein First of all, the pathway-specific acyl carrier protein (ACP) was characterized, since acyl-activated enzymology was presumed, as in the polyketide synthase and non–ribosomal peptide synthetase reactions. Initially, certain acyl ACP intermediates derived from L-glutamate or GABA were presumed from the previous incorporation study. The ACP was expressed in E. coli and the majority of the recombinant protein was obtained as the apo form of ACP. Therefore, the gene was then coexpressed with the wide-spectrum 40 -phosphopantetheine transferase (PPTase, Sfp) derived from B. subtilis to obtain the holo form of the ACP, which was confirmed by LC-ESI-MS. NH3+

g-L-glutamyl-ACP ligase∗

ACP-SH

–O C 2

ATP

S

ADP and Pi

ACP

ð20:16Þ

O

L-glutamate 2+, Mn2+

Mg

+H N 3

S O

g-L-glutamyl-g-aminobutyryl-ACP ligase∗

ACP

O –O

ATP L-glutamine Mg2+, Mn2+

∗same enzyme as the ligase at the first step

2C

NH3+

N H

S

ACP

ADP and Pi

O

ð20:17Þ

The second enzyme was g-L-glutamyl–ACP ligase, which shows weak local similarity to biotin carboxylase and carbamoyl phosphate synthase. These enzymes belong to the growing ATP grasp superfamily, which includes D-alanine:D-alanine ligase, glutathione synthetase, glycinamide ribonucleotide transformylase, and succinyl-CoA synthetase. These enzymes

512


exhibit a characteristic ATP-binding motif and possess ATP-dependent carboxyl-amine or carboxyl-thiol ligase activity. The reaction mechanism of the enzymes was proposed to involve the generation of acyl phosphate intermediate via hydrolysis of ATP to ADP. The enzyme was expressed in E. coli and the recombinant protein was found to require DTT in the buffer solution to prevent aggregation. The recombinant enzyme reacted with ATP and various amino acids including L-glutamate, D-glutamate, GABA, and AHBA in the presence of MgCl2, MnCl2, and KCl. Then, a possible ATPase activity was demonstrated by detecting ADP, which was coupled to the reaction of pyruvate kinase and NADH-dependent lactate dehydrogenase (LDH). As a result, only in the presence of L-glutamate, a decrease of NADH was observed, suggesting L-glutamate-dependent ATPase activity of the enzyme. Also, this observation implied the involvement of a possible L-glutamylphosphate intermediate, which can be transferred onto the holoACP. In fact, addition of the holo-ACP to the enzyme reaction gave L-glutamyl-S-ACP as product, which was detected by LC-ESI-MS Eq. (20.16). Since L-glutamate should be converted to AHBA through decarboxylation of the a-carboxy group, the g-carboxy group was presumed to be attached to the ACP. Obviously L-glutamate was found to be the best substrate for the reaction rather than GABA and AHBA, and consistent with the incorporation experiments. The incorporation of GABA into butirosin could be due to a weak activity of the enzyme.

6.2. g-L-glutamyl ACP decarboxylase The enzyme shows significant similarity to a PLP-dependent ornithine/arginine/diaminopimerate decarboxylase family and was thus thought to be involved in the decarboxylation of the a-carboxyl moiety of the L-glutamate portion Eq. (20.18). To investigate the expected decarboxylation activity, the expressed purified enzyme was added to the above-mentioned g-L-Glu–ACP ligase reaction mixture to detect the probable GABA-ACP intermediate. However, surprisingly, g-L-glutamyl-g-aminobutylyl–ACP was clearly observed by LC-ESI-MS. No g-L-Glu-g-L-Glu–ACP or GABA–ACP was observed in the reaction mixture, suggesting that after the decarboxylation of g-L-Glu–ACP to GABA–ACP by the decarboxylase, the resulting GABA– ACP was further glutamylated by g-Glu–ACP ligase to afford g-L-GluGABA–ACP. In fact, when the g-Glu–ACP ligase reaction was quenched by the addition of EDTA after g-L-Glu–ACP formation, and then the decarboxylase was added to the solution, the expected GABA-ACP was observed by LC-ESI-MS. This decarboxylase weakly accepted free amino acids, suggesting specific recognition of the acyl ACP. Thus, a series of transformations from the holo-ACP to g-L-Glu-GABA–ACP Eq. (20.16), (20.18), then (20.17) have been clearly demonstrated.

513


NH3+

S

–O

2C

g-L-glutamyl ACP decarboxylase

ACP

S

+H

3N

ACP

CO2

O

PLP

O

ð20:18Þ

6.3. g-L-glutamyl-4-aminobutyryl ACP mono-oxygenase and NAD(P)H:FMN oxidoreductase The third and fourth enzymes are involved in a two-component monooxygenase reaction, which is the stereospecific hydroxylation at C-2 of g-LGlu-GABA–ACP to give g-L-Glu-AHBA–ACP Eq. (20.19). One enzyme is g-L-Glu-GABA–ACP mono-oxygenase encoded in the butirosin-specific gene cassette. This enzyme shows sequence similarity to alkanesulfonate mono-oxygenase, which is involved in sulfur metabolism. This enzymatic reaction system utilizes two proteins to hydroxylate the substrate; one is an oxidoreductase that catalyzes the reduction of FMN with NAD(P)H as an electron donor, and the other is a mono-oxygenase that catalyzes the hydroxylation of the substrate in the presence of FMNH2 and molecular oxygen. In this two-component system, the oxidoreductase provides FMNH2 to the mono-oxygenase to reduce molecular oxygen to water and to attach a hydroxy group to the substrate. This sort of enzyme is encoded 20 kb upstream of the mono-oxygenase gene in the butirosin biosynthetic gene cluster. Thus, the enzyme was proposed to be the oxidoreductase to supply FADH2 to the mono-oxygenase. g-L-glutamyl-g-aminobutyryl-ACP monooxygenase O2 H2O

O –O

2C

NH3+

N H

S O

O

–O C 2

ACP FMNH2

NADH + H+

FMN

NH3+

OH N H

S

ACP

O

ð20:19Þ NAD+

NADH:FMN oxidoreductase

The purified recombinant oxidoreductase expressed in E. coli was found to contain FMN as the cofactor rather than FAD. Kinetic studies on this oxidoreductase were also conducted to characterize it as an NAD(P)H: FMN oxidoreductase, which reduces FMN to FMNH2 with NAD(P)H, presumably for the FMN-dependent mono-oxygenase. The mono-oxygenase was also expressed in E. coli and purified in colorless form, suggesting no bound flavin cofactor in the protein. Thus, free FMNH2 from the above-mentioned NADH:FMN oxidoreductase seemed to be used for the mono-oxygenase reaction. In fact, in the presence of the NADH:FMN oxidoreductase system, the recombinant monooxygenase catalyzed the hydroxylation of g-L-Glu-GABA–ACP to afford g-L-Glu-AHBA–ACP, which was detected by LC-ESI-MS. Indeed, FMN and molecular oxygen were required for the hydroxylation.

514


6.4. g-L-glutamyl-4-amino-2-hydroxybutyryl ACP: ribostamycin g-L-glutamyl-4-amino2-hydroxybutyryltransferase Two unpredictable functional enzymes encoded in the butirosin-specific biosynthetic gene cassette were then proposed to be involved in the AHBA acyl-transfer reaction to complete butirosin B biosynthesis Eqs. (20.20) and (20.21). Both enzymes were expressed in E. coli and purified to investigate the expected reaction with the above-mentioned g-L-Glu-AHBA–ACP, which was prepared by Sfp with synthetic g-L-Glu-AHBA-CoA, and ribostamycin as the acceptor substrate. The LC-ESI-MS analysis of the reaction clearly confirmed the production of butirosin B. When either of two enzymes was used for the reaction, only one of the reactions gave a new product, which corresponded to g-L-Glu-butirosin B. Thus, the enzyme was characterized as g-L-Glu-AHBA–ACP:ribostamycin g-L-Glu-AHBA transferase. Subsequently, another enzyme was found to be involved in the cleavage of the glutamyl moiety to give butirosin B. NH2 HO HO

NH2

O

H2N H2N O O HO O

g-L-glutamyl-4-amino-2-hydroxybutyryl ACP:ribostamycin g-L-glutamyl-4-amino-2-hydroxybutyryltransferase NH2 OH

-O

2C

HO OH

OH

O

NH+ 3

N H

S O

HO HO

O

H2N H2N O O HO O

OH

H N OH O

ACP HO OH

+ ACP

O N H

CO3NH+ 3

ð20:20Þ

In this two-enzyme reaction, 2DOS and paromamine were not converted to the corresponding AHBA derivatives. Neamine was less effectively converted to AHBA neamine than ribostamycin. Thus, as illustrated, ribostamycin is the only true substrate in butirosin biosynthesis. On the other hand, the related 4,5-disubstituted aminoglycosides, paromomycin and neomycin, were converted to the corresponding AHBA derivatives but not the 4,6-disubstituted aminoglycoside gentamicin C and kanamycin. However, interestingly, when a high concentration of the g-L-Glu-AHBA– ACP analog, g-L-Glu-AHBA–S-N-acetylcysteamine thioester (5 mM) was used for the g-L-Glu-AHBA transferase reaction, various aminoglycosides, including neomycin, paromomycin, 2DOS, paromamine, neamine, apramycin, kanamycin, and gentamicin C, were all acylated. Since the g-L-GluAHBA–SNAC thioester was also hydrolyzed by the following g-L-glutamyl cyclotransferase, stepwise addition of the two enzymes was required to complete the synthesis of the AHBA derivative (Llewellyn and Spencer, 2008). This chemoenzymatic method could be a promising approach to create a range of AHBA-containing aminoglycosides.

6.5. g-L-glutamyl-butirosin B g-L-glutamyl cyclotransferase As the above-mentioned results show, the enzyme responsible for the final step of AHBA attachment has been clarified as g-L-Glu cyclotransferase. Another product of the reaction, in addition to butirosin B, was found to be

515


pyroglutamate. Thus, this enzyme was characterized as a novel g-L-Glu-butirosin g-L-Glu cyclotransferase. The reaction mechanism was proposed as the direct nucleophilic transamidation of the a-amino group of the g-L-glutamyl moiety to give butirosin B and pyroglutamate Eq. (20.21). An alternative mechanism involving initial formation of a g-L-Glu-enzyme intermediate followed by pyroglutamate formation cannot be excluded. Further detailed biochemical studies are necessary to elucidate the reaction mechanism. NH2 HO HO HO

O H2N H2N O O O

HO OH

OH

H N OH

O

O N H

CO2– NH3+

γ-L-glutamyl-butirosin B γ-L-glutamyl-cyclotransferase

NH2 O

HO HO

H2N HO

H2N OO

O

H N OH

OH NH2

O + pyroglutamate

HO OH Butirosin B

ð20:21Þ

The only remaining butirosin-specific enzyme encoded in the cluster is a putative NADPþ-dependent dehydrogenase, which might be involved in the epimerization at C-300 of butirosin B and ribostamycin to butirosin A and xylostacin, respectively. Or the above-mentioned inactive enzyme as a NAD-dependent 2DOIA dehydrogenase might be involved in this reaction. At the present time, even the timing of the epimerization of the ribose moiety is not clear. Further investigation is therefore required to fully understand the epimerization reaction in the butirosin biosynthetic pathway.

7. Other Related Enzymes in the Biosynthesis of 2DOS-Containing Aminoglycoside Antibiotics Thus far, not so many enzymatic studies on kanamycin, tobramycin and gentamicin have been reported. Only several of the same enzymes in the biosynthesis of 2DOS, paromamine, and neamine have been characterized by enzymatic analyses and gene inactivation studies. Some of the enzymes from the kanamycin biosynthetic pathway are covered in Chapter 19 in this volume. Thus far, the basic enzymatic reactions from G6P to 2DOS, paromamine, and neamine are believed to be the same as characterized in butirosin and neomycin biosynthesis. Although several branching enzymatic reactions such as deamination, deoxygenations, methylations, and glycosylations should be involved, no specific enzyme for such reactions has been enzymatically characterized yet. One important study by heterologous expression of several enzymes encoded in the gentamicin biosynthetic gene cluster in Streptomyces venezuelae showed the involvement of a novel type of glycosyltransferase, which catalyzes xylose transfer onto paromamine in the biosynthesis of gentamicin (Park et al., 2008). Although detailed studies are required, this is definitely a branching enzyme among structurally diverse 2DOS-containing aminoglycoside antibiotics. Another recent important finding by gene

516


inactivation is that a radical SAM enzyme is involved after gentamicin A2 formation (Kim et al., 2008). Further enzymatic analysis will show more detailed enzymatic reaction mechanisms in this class of antibiotics.

8. Concluding Remarks and Future Perspectives The sequential biosynthetic pathways leading to butirosin and neomycin have been mostly enzymatically characterized. This knowledge shed light on the other related aminoglycoside biosynthetic enzymology. These researches also provided fundamental insights into unexpected enzymatic functions, such as the AHBA biosynthetic enzymes and radical SAM dehydrogenase. Thus, characterizations of secondary metabolite biosynthetic enzymes expand general knowledge of enzymatic catalysis, in addition to understanding complex secondary metabolism. The epimerization steps in both butirosin and neomycin biosynthetic pathways are still unclear. These enzymatic reactions should be quite unique to each biosynthetic pathway and thus expand the molecular diversity of aminoglycoside antibiotics. Although structurally related kanamycin-, tobramycin- and gentamicinspecific biosynthetic enzymes are not enzymatically characterized, the branching enzymes will be functionally characterized based on the knowledge of the butirosin and neomycin biosynthetic enzymes. Then, in future, combinatorial enzymatic synthesis of aminoglycoside antibiotics, which might produce significant biological activities, should be possible. For the purpose, in addition to characterization of enzymes, further detailed biochemical studies on the biosynthetic enzymes will be needed.

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C H A P T E R

T W E N T Y- O N E

Enzymatic Synthesis of TDP-Deoxysugars Jessica White-Phillip,* Christopher J. Thibodeaux,* and Hung-wen Liu*,†,‡ Contents 1. Introduction 2. Enzymatic Synthesis of TDP-a-D-glucose 2.1. Preparation of enzymes required for in vitro synthesis of TDP-a-D-glucose (5) 2.2. Enzymatic synthesis of TDP-a-D-glucose (5) 2.3. Purification of TDP-a-D-glucose (5) 3. Generation of TDP-4-keto-6-deoxy-a-D-glucose (6) 3.1. Generation of TDP-2,6-dideoxysugars 4. In vitro Reconstitution of Entire Deoxysugar Biosynthetic Pathways 4.1. One-pot synthesis of TDP-a-D-mycaminose (12) 4.2. Two-stage one-pot synthesis of TDP-b-L-mycarose (13) 4.3. Multistep enzymatic synthesis of TDP-a-D-forosamine (14) 4.4. TDP-a-D-desosamine (11) 5. Synthesis of Deoxysugars In Vivo by Metabolic Pathway Engineering 6. Summary Acknowledgment References

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Abstract Many biologically active bacterial natural products contain highly modified deoxysugar residues that are often critical for the activity of the parent compounds. Most of these deoxysugars are secondary metabolites that are biosynthesized in the form of nucleotide diphosphate (NDP) sugars prior to their transfer to natural product aglycones by glycosyltransferases. Over the past decade, many biosynthetic pathways that lead to the formation of these

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Institute for Cellular and Molecular Biology, University of Texas-Austin, Austin, Texas, USA College of Pharmacy, University of Texas-Austin, Austin, Texas, USA Department of Chemistry and Biochemistry, University of Texas-Austin, Austin, Texas, USA


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Jessica White-Phillip et al.

unusual sugars have been unraveled, and the mechanisms of many key enzymatic transformations involved in these pathways have been elucidated. However, obtaining workable quantities of NDP-deoxysugars for in vitro studies is often a difficult task. This limitation has hindered an in-depth investigation of the substrate specificity of deoxysugar biosynthetic enzymes, many of which are promiscuous with respect to their NDP-sugar substrates and are, thus, potentially useful catalysts for natural product glycoengineering. Presented in this review are procedures for the enzymatic synthesis and purification of a variety of NDP-deoxysugars, including some early intermediates in NDP-deoxysugar biosynthetic pathways, and highly modified NDP-deoxysugars that are late intermediates in their respective biosynthetic pathways. The procedures described herein could be used as general guidelines for the development of specific protocols for the synthesis of other NDP-deoxysugars.

1. Introduction Glycosylation is important for the biological activity of macrolide, peptide, and aminoglycoside antibiotics as well as numerous anticancer, antiparasitic, and antifungal agents of diverse biosynthetic origin (Lamb and Wright, 2005; Mendez and Salas, 2001; Walsh et al., 2003). These sugar residues play crucial biological roles in many natural products and their removal oftentimes results in the loss of biological activity (Mendez and Salas, 2001; Thorson et al., 2001; Weymouth-Wilson, 1997). The most chemically diverse group of carbohydrate moieties found in secondary metabolites are 6-deoxyhexoses, which are produced by a variety of organisms, but are most prevalent in actinomycetes, a group of soil bacteria that are a rich source of biologically active secondary metabolites (Salas and Mendez, 2007). For many of the natural product biosynthetic pathways found in these organisms, a combination of experimental evidence and database comparisons has been used to identify the deoxysugar biosynthetic genes and to delineate the corresponding biosynthetic pathways (reviewed in Thibodeaux et al., 2008). Altering and/or exchanging the sugar structures and points of aglycone attachment in natural products is a feasible route to enhance or vary the physiological properties of these compounds. Importantly, it has been discovered that many natural product glycosyltransferases (or GTs enzymes which link activated sugar donors to aglycone acceptors) exhibit relaxed substrate specificity. This discovery has resulted in an explosion in the use of these GTs to engineer natural products with altered glycosylation patterns, and has generated several broad and complementary glycoengineering strategies (Thibodeaux et al., 2007, 2008). For in vivo glycodiversification, deoxysugar biosynthetic pathways can be altered within a producing bacterial strain using gene disruption and/or heterologous gene

Enzymatic Synthesis of TDP-Deoxysugars

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expression methods in order to reroute sugar biosynthetic intermediates to new final products. Alternatively, new biosynthetic pathways can be assembled in hosts that do not normally produce glycosylated natural products. Furthermore, these genetically engineered bacteria can be fed with non-native aglycones or they can be transformed with additional plasmids in order to produce novel compounds in a combinatorial fashion. In vitro, purified wildtype or engineered sugar biosynthetic enzymes can be used to synthesize specific natural product glycoforms, as well as to prepare libraries of novel glycoforms. A detailed understanding of the organization of deoxysugar biosynthetic machinery and of the biochemical properties of the enzymes involved in synthesizing and coupling these sugars to their aglycones is critical to the success of these enzymatic glycoengineering approaches. A significant limitation to in vitro glycodiversification efforts is the availability of activated sugar donor substrates for the promiscuous glycosyltransferases. There are several methods by which activated deoxysugar donors (i.e., NDP-deoxysugars) are generally obtained. Total chemical synthesis, though feasible, often requires significant technical expertise and can be plagued by low yields. Another approach for obtaining highly modified NDP-deoxysugars is to hydrolyze the desired reducing sugar from the natural product, and then to chemically synthesize the NDP sugar from the reducing sugar (Chang et al., 2000; Chen et al., 2002). The recently discovered in vitro reversibility of GT-catalyzed reactions may provide a new synthetic route to some NDP-deoxysugars (Bode and Muller, 2007; Melancon et al., 2006; Minami et al., 2005; Zhang et al., 2006), but it is not yet clear whether this will be a generally applicable method for NDP-deoxysugar synthesis. Thus, we envision that multistep enzymatic and chemoenzymatic synthesis will continue to play an important role in the production of structurally complex, NDP-activated deoxysugar donors for glycobiology and glycoengineering studies. Enzymatic synthesis is appealing for several reasons. Enzymes generally catalyze reactions with well-defined regio- and stereo-chemical preference and enzymes are a readily renewable resource. Furthermore, when manipulated in vitro, it is possible to present these enzymes with substrate analogues that contain chemically incorporated non-natural functional groups. Introduction of certain types of functional groups can allow further chemoselective derivitization to enhance the structural diversity of glycoform libraries (Fu et al., 2003). Here, we provide detailed protocols for the enzymatic synthesis and purification of several thymidine diphosphate (TDP)–activated deoxysugar intermediates that are common to many TDP-deoxysugar biosynthetic pathways in bacteria. These methods should be useful for researchers interested in obtaining workable quantities of a desired TDPdeoxysugar. We also highlight several successful examples of in vitro multistep enzymatic syntheses of highly modified TDP-deoxysugars and discuss how biosynthetic machinery can be manipulated in vivo to generate

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desired deoxysugar structures. For more information on NDP-deoxysugar synthesis, bacterial deoxysugar biosynthesis, mechanistic studies of deoxysugar biosynthetic enzymes, and the application of these enzymes in glycoengineering efforts, the reader is directed to several recent and comprehensive reviews (He et al., 2000; He and Liu, 2002; Langenhan et al., 2005; Luzhetskyy et al., 2008; Mendez et al., 2008; Rupprath et al., 2005; Salas and Mendez, 2007; Thibodeaux et al., 2008).

2. Enzymatic Synthesis of TDP-a-D-glucose Most of the unusual deoxysugars produced by bacterial biosynthetic pathways are derived from a-D-glucose-1-phosphate (1) which is, in turn, derived from D-glucose (2) by direct anomeric phosphorylation, or from glucose-6-phosphate (3) by a phosphohexose mutase-catalyzed reaction (Scheme 21.1). A nucleotide monophosphate (NMP) moiety from a nucleotide triphosphate (NTP) is then coupled to (1) by a nucleotidylyltransferase to generate NDP-glucose (4). While some deoxysugars are derived from primary metabolites and can be activated with other NDP groups, the vast majority of bacterial deoxysugars used in glycosylation of secondary metabolites are TDP-sugars that are biosynthetically derived from TDP-a-Dglucose (5) (Thibodeaux et al., 2007, 2008). Because TDP-a-D-glucose (5) is rather costly, efficient methods for its preparation are desirable. Towards this end, we have developed an efficient and facile one-pot, two-step enzymatic synthesis for 5 (Scheme 21.2) using readily available enzymes and inexpensive substrates (Takahashi et al., 2006). In the first step of this reaction, thymidine is converted to thymidine triphosphate (TTP) by the

HO HO

OH O HO

D-glucose

HO HO

Anomeric kinase OH

(2)

OPO=3 O HO

HO HO Phosphohexose mutase

OH

OH O

= HO OPO3 Glucose 1-phosphate (1)

Nucleotidylyltransferase HO HO NTP

PPi

OH O HO OTDP

NDP-a-Dglucose (4) NDP = TDP (5)

Glucose 6-phosphate (3)

Scheme 21.1 Activation of the glycolytic intermediates D-glucose and glucose6-phosphate. D-Glucose (2) and glucose-6-phosphate (3) can be converted to aD-glucose-1-phosphate (1) by anomeric kinase and phosphohexose mutase, respectively. A nucleotide monophosphate (NMP) moiety is then tranferred from the corresponding NTP to 1 by a nucleotidylyltransferase to form NDP-a-D-glucose (4). Most deoxysugars produced in bacterial secondary metabolism are derived fromTDP-a-D-glucose (5).

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Step 1 O H

N

Me

NDK

TMK TMP

O N O

HO

TK

ATP

ADP

TDP ATP

ADP

P O

ATP

ADP

O

O

_ O_

H

O

O O P _ O_ P O O O

HO PK

PK

N

Me

O N O HO

PK TTP

Pyr

PEP

Pyr

Pyr

PEP

PEP OH O

Step 2 O O O P O _ O O_ P O P O _ O O

_

HO HO

O H

N

HO OPO=3

Me

1

HO HO

O N O

HO OTDP

RfbA TTP

HO

OH O

PPi

TDP-a-D-glucose (5)

Scheme 21.2 Enzymatic synthesis of TDP-a-D-glucose (5). In the first step of TDP-aD-glucose (5) synthesis, thymidine triphosphate (TTP) is synthesized from thymidine by three successive ATP-dependent phosphorylations catalyzed by thymidine kinase (TK), thymidylate kinase (TMK), and nucleotide diphosphate kinase (NDK). ATP is only needed in catalytic amounts, due to a pyruvate kinase (PK) ATP regeneration system, which transfers a phosphate group from phosphoenol pyruvate (PEP) to ADP, yielding pyruvate (Pyr) and ATP. Removal of the enzymes and addition of a-D-glucose-1-phosphate (1) and RfbA (a thymidylyltransferase from S. enterica) leads to the synthesis of TDP-a-D-glucose (5).

sequential action of three separate ATP-dependent kinases (thymidine kinase [TK], thymidylate kinase [TMK], and nucleotide diphosphate kinase [NDK]). ATP is continually regenerated by pyruvate kinase at the expense of phosphoenol pyruvate (PEP). In the second step of the synthesis, a TMP moiety from TTP is coupled to a-D-glucose-1-phosphate (1) by RfbA, an a-D-glucose-1-phosphate thymidylyltransferase from Salmonella enterica. The resulting TDP-a-D-glucose (5) can then be purified or used directly for subsequent enzymatic transformation.

2.1. Preparation of enzymes required for in vitro synthesis of TDP-a-D-glucose (5) The genes encoding thymidine kinase (TK), thymidylate kinase (TMK), and nucleotide diphosphate kinase (NDK) can be amplified from the genomic DNA of Escherichia coli strain HMS174 and cloned in tandem into a single pET28b (þ) plasmid, such that each gene has its own ribosome binding site and a His6-tag. The three enzymes can then be coexpressed from the same plasmid in E. coli BL21(DE3) using standard growth and induction conditions. Typically, we inoculate LB medium (supplemented

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with 50 mg/ml kanamycin) with a 1:500 dilution from an overnight culture, grow the cells at 37 , induce with 0.3 mM isopropyl a-thiogalactoside (IPTG) at OD600 readings of 0.4 to 0.6, and continue to grow for 18 h at 25 . Cells are then harvested by centrifugation, lysed by sonication, and centrifuged to remove cellular debris. The recombinant TK/TMK/NDK enzymes are then purified from the resulting supernatant using Ni-NTA affinity chromatography. We generally use 150 mM NaCl in our lysis, wash, and elution buffers, as opposed to the 300 mM recommended in the QIAexpressionist protocol. The mixture of purified TK/TMK/NDK enzymes is dialyzed against 50 mM potassium phosphate buffer (pH 7.5) containing 15% glycerol. In parallel, rabbit muscle pyruvate kinase from Sigma (purchased as a 400 to 800 units/mg ammonium sulfate precipitate) is dissolved in water to a concentration of 2500 units/ml, dialyzed against buffer (50 mM NaH2PO4, 300 mM NaCl, pH 8.0) to remove the ammonium sulfate, dispensed in 500-ml to 1-ml aliquots, flash frozen in liquid nitrogen, and stored at –80 . The rfbA gene is PCR-amplified from S. enterica serovar Typhimurium LT2 genomic DNA with an upstream ribosome binding site (RBS)/translational spacer element (TSE) and is inserted into a pUC18 vector. A (His)5-tag is added to the C-terminus of RfbA by PCR amplification of the rfbA insert of plasmid rfbA/pUC18. The newly synthesized gene is then cloned into the same vector, and the resulting plasmid, rfbA-(His)5/pUC18, introduced by transformation into BL21(DE3) for protein expression. Expression and purification conditions are identical to those for the TK/TMK/NDK enzymes, except that 100 mg/ml ampicillin is used for selection. Induction with IPTG is unnecessary. The approximate amount of RfbA purified from an average 6 l culture is 140 mg. Each liter of culture produces enough RfbA to perform one large-scale TDP-a-D-glucose synthesis reaction.

2.2. Enzymatic synthesis of TDP-a-D-glucose (5) In the first step of TDP-a-D-glucose synthesis, a 15-ml reaction mixture containing phosphoenol pyruvate (PEP, 85.6 mM ), thymidine (27 mM ), ATP (1.8 mM ), and MgCl2 (30 mM ) in 50 mM TrisHCl buffer (pH 7.5) is prepared. After the addition of these reagents, the pH of the solution is adjusted to 7.5 using NaOH or HCl prior to the addition of enzymes. Pyruvate kinase (PK, 1000 units) and 30 mg of the TK/TMK/NDK enzyme mixture are added to give final concentrations of 25 to 30 mM for each of the TK/TMK/NDK enzymes. The reaction mixture is incubated at 37 for 4 h, and then filtered through an Amicon ultrafiltration cell unit (YM-10 membrane) under nitrogen at 4 to remove the enzymes. The flowthrough is collected in a 50-ml conical tube and the pH adjusted to 7.5. This flowthrough contains TTP and will be used in the next step.


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To synthesize TDP-a-D-glucose (5), a mixture containing TTP, a-Dglucose-1-phosphate (1, 4 mM ), MgCl2 (30 mM ), and recombinant RfbA (47 mM ) is prepared. This reaction mixture is incubated at 30 to 37 for 12 to 16 h. It is important to keep the Mg2þ ion concentration similar to that of TTP, as a high molar excess of Mg2þ (over TTP) is known to adversely affect the activity of RfbA (Amann et al., 2001). After the reaction is completed, any precipitate is removed by centrifugation at 5000g for 10 min at 4 . RfbA is then removed by filtering the supernatant through either a YM-10 Amicon stirred cell unit or a 50-ml conical centrifugal filter unit at 4 . TDP-D-glucose (5) present in the flowthrough can now be purified or used directly for the following enzymatic reactions.

2.3. Purification of TDP-a-D-glucose (5) TDP-a-D-glucose (5) can be purified with either anion exchange or sizeexclusion chromatography. When using the size-exclusion method, the sample containing 5 should be frozen and lyophilized to reduce the volume to 1 to 2 ml. It is important to note that many TDP-sugars degrade significantly if lyophilized to dryness without removing salts. For this reason, if the stability of a particular NDP-sugar is not known, it is always best to lyophilize to half-volume, dilute with the appropriate solvent, and lyophilize to half-volume again. This cycle can be repeated as necessary to exchange solvents or to remove volatile substances. When the concentrated TDP-a-D-glucose is thawed, the sample typically appears as a slightly yellowish syrup. If purifying by FPLC using a MonoQ column, the TDP-a-D-glucose sample does not need to be lyophilized/ concentrated before purification. 2.3.1. Purification using size-exclusion chromatography A P2 biogel (Biorad) column (2.5 100 cm) is packed following the manufacturer’s instructions and equilibrated with 1 l of filtered ddH2O at 4 . After washing, the solvent above the column bed is removed and the concentrated (1-2 ml) TDP-a-D-glucose-containing reaction mixture is carefully applied to the top of the resin bed. The sample is introduced into the resin by gravity flow, ensuring that the resin bed does not dry. After the sample has been loaded onto the column, water is used to elute the TDP-sugar product. An adjustable pneumatic pump is used to maintain the flow rate at about 6 to 12 ml per hour. Under these conditions, 5 will generally be eluted from the column around 24 to 48 h after loading. Fractions eluted from the P2 column can be analyzed by UV-Vis spectroscopy for the presence of 5 (e267 = 9600 M–1cm–1). Usually, there will be a sharp rise in absorbance in the first fraction containing 5, followed by 10 to 12 fractions with high absorbance readings, and then a sharp decrease in absorbance to baseline. The fractions comprising the absorbance plateau are individually frozen and lyophilized until their purity can be verified by HPLC or NMR spectroscopy.

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To verify the purity of the fractions from the P2 size-exclusion column, HPLC analysis is performed using a Dionex Carbopac PA1 column and a 20-ml sample injection volume. With water as solvent A and 500 mM NH4OAc (adjusted to pH 7.0 with aqueous NH3) as solvent B, the following gradient elution is typically used: 5 to 20% B over 15 min, 20 to 60% B over 20 min, 60 to 100% B over 2 min, 3-min wash at 100% B, 100 to 5% B over 5 min, and re-equilibration at 5% B for 15 min. The flow rate is 1 ml/min and the detector is set at 267 nm. Under these conditions, the retention times for compounds are as follows: TDP-a-D-glucose (31 to 33 min), TMP (25 min), and TDP (41 to 42 min). In samples of high concentration, TDP-a-D-glucose (5) may be eluted anywhere from 30 to 34 min, but will still appear as a very sharp, well-defined peak in the HPLC trace. Fractions are pooled based on purity either before or after lyophilization. The final concentration of 5 in pooled fractions can be determined spectrophotometrically (e267 = 9600 M–1cm–1). The theoretical yield for this reaction is 228 mg (based on a 15 ml reaction and the substrate concentrations described in 1.2), but the actual yields range from 150 to 200 mg, depending on one’s experience with the methodology. Most commonly, one will find that 10% of the product will be greater than 90% pure, 40% will be 90 to 80% pure, 40% will be 80 to 70% pure, and the remaining 10% will be less than 70% pure. The most abundant contaminants using P2 size-exclusion chromatography as the method of purification are TMP and TDP, both of which can be removed by further FPLC purification. However, we have found that further purification is usually unnecessary if 5 is to be used in enzymatic reactions with high yield (i.e., >90% conversion). 2.3.2. FPLC Purification of TDP-a-D-glucose (5) As mentioned above, P2 chromatography will separate the majority of unreacted a-D-glucose-1-phosphate (1) from the desired TDP-a-D-glucose product (5). However, only about 10% of 5 will be greater than 90% pure, with the remainder being contaminated by TDP and TMP. The majority of TDP and TMP in these samples can be removed if the P2 fractions are subjected to anion exchange chromatography. It should be noted that, in many cases, the enzymatically synthesized TDP sugars can be purified directly by FPLC, bypassing the P2 purification step. Purification of 5 can be achieved by FPLC using a Mono Q 10/10 or 16/10 column with a gradient, where HPLC-grade water is buffer A and 400 mM NH4HCO3 in water (pH 8.2) is buffer B. Flow rates of 1 or 4 ml/min are used for the 10/10 and 16/10 columns, respectively. To elute 5, the following gradient is used: from 0 to 10% B over 0.5 column volumes, from 10 to 40% B over four column volumes (which will elute the TDP-sugar), and from 40 to 100% B over 0.5 column volumes. The column is then washed with 100% B for two-column volumes, followed by reduction to 0% B over 0.5 column volumes, and re-equilibration at 0% B over


529

three-column volumes. TDP-a-D-glucose (5) will be eluted at approximately 30 min. Other important peaks include TMP and TDP, which have retention times of 29 min and 34 min, respectively. Fractions containing the major peak from each injection should be lyophilized individually, redissolved in water, and lyophilized again to remove NH4HCO3. Alternatively, fractions can be desalted using a G-10 column (see Section 2.3.3), lyophilized, redissolved in water, and lyophilized again. The purities of these fractions should be analyzed by 1H and 31P NMR spectroscopy and typically range from 50 to 90%. If one begins with pre-purified 5 (from P2 column), the purity will be much higher, ranging from 75 to 95%. The most common residual contaminant is TMP. The resolution of the Mono-Q column is not sufficient to completely separate TMP from TDP-a-D-glucose (5) in cases of large injection volumes or high concentrations of samples. From this method, one can typically obtain an average of 25 mg of 90% pure TDP-a-D-glucose from 150-200 mg of 5. 2.3.3. Desalting FPLC fractions To desalt the FPLC fractions, a Sephadex G-10 desalting column (25 mm 50 cm) is prepared according to manufacturer’s instructions. After loading the FPLC fraction, the column is washed with 1 l HPLC-grade water using gravity flow (24 ml/h) at 4 . Fractions of 2 ml are collected over a period of 12 to 15 h. TDP-a-D-glucose is typically eluted over 5 to 10 fractions within 8 h. Fractions displaying significant absorbance at 267 nm are combined, lyophilized to near dryness, resuspended in HPLC-grade water, and lyophilized again to remove the remaining NH4HCO3 from the TDP-sugar. After desalting, TDP-a-D-glucose is quite stable and can be stored for months at – 80 , although multiple freeze–thaw cycles should be avoided. For the majority of enzymatic reactions, we have found that the presence of small amounts of contaminating TMP or TDP do not significantly affect subsequent enzymatic reactions. Thus, TDP-a-D-glucose of greater than 85% purity obtained after P2 size-exclusion chromatography or FPLC purification is generally sufficient. For applications that require a much higher substrate purity, such as kinetic analysis or in situ 1H NMR spectroscopic assays to determine the activity of sugar biosynthetic enzymes, the complete purification sequence of P2, FPLC, and G-10 chromatography should be followed to achieve greater than 95% purity.

3. Generation of TDP-4-keto-6-deoxy-aD-glucose (6) TDP-4-keto-6-deoxy-a-D-glucose (6) is produced from TDP-a-Dglucose (5) by the action of TDP-glucose-4,6-dehydratase (4,6-DH, Scheme 21.3). TDP-4-keto-6-deoxy-a-D-glucose is a common intermediate in many

530


OH O

HO HO

4,6-DH

O

Me

NAD+

HO OTDP

O

HO HO

OTDP

TDP-4-keto-6-deoxya-D-glucose (6)

TDP-a-D-glucose (5)

2-DH

O

Me

HO

3-KR Equatorial

O

O

Me

Me

O

O HO

O 8

OTDP

7

O maltol

3-KR Equatorial O

OTDP

3-AT

Me O HO

O OTDP

9

Me O

H2N 10

OTDP

Scheme 21.3 EarlyTDP-deoxysugar biosynthetic intermediates. The key intermediate in TDP-deoxysugar biosynthesis is TDP-4-keto-6-deoxy-a-D-glucose (6), which is synthesized from 5 by a TDP-glucose-4,6-dehydratase enzyme (4,6-DH). This intermediate is a branching point for the biosynthetic pathways of TDP-deoxysugars. For 2,6-dideoxyhexoses, 6 is converted to the unstable intermediate 7 by a 2-dehydratase (2-DH).This intermediate can then be reduced by a 3-ketoreductase (3-KR) to give 8 or 9, or it can be transaminated by a 3-aminotransferase (3-AT) to give10.

sugar biosynthetic pathways. The 4-keto group provides a chemically versatile handle that can be manipulated by deoxysugar biosynthetic enzymes to produce structurally diverse products. Due to such versatility, the ready availability of this sugar is important for the investigation of many interesting deoxysugar biosynthetic enzymes. Not surprisingly, several useful methods for the synthesis of 6 have been developed (Elling et al., 2005; Oh et al., 2003; Rupprath et al., 2005; Stein et al., 1998). To enzymatically synthesize this sugar, we cloned rfbB (the 4,6-DH from S. enterica serovar Typhimurium LT2) into the pUC18 vector and expressed the recombinant RfbB-(His)5 protein in E. coli BL21(DE3) in a manner similar to that described in Section 1.1 for the RfbA-(His)5 construct (Melancon et al., 2007; Takahashi et al., 2006). Recombinant RfbB was purified by affinity chromatography using Ni-NTA resin (Qiagen) according to the manufacturer’s instructions, with the exception that 10% glycerol was included in the lysis, wash, and elution buffers. RfbB was then dialyzed against 50 mM NaH2PO4 buffer, 300 mM NaCl, 15% glycerol (pH 8.0), concentrated, flash frozen in liquid nitrogen, and stored at –80 until use. The yield of RfbB is generally 80 mg per liter of growth culture.


531

A typical RfbB reaction contains 5 (25 mg) and RfbB (5 mg) in 2 ml of 50 mM TrisHCl buffer (pH 7.5). Incubation is carried out at 30 for 3 to 4 h. The reaction is slightly slower at 30 , but less degradation of 5 occurs at this temperature. Some published procedures (Amann et al., 2001; Stein et al., 1998) use alkaline phosphatase (2 units/ml) to suppress any competitive inhibition by TDP, but we have found its inclusion to be unnecessary. The progress of the reaction can be monitored by HPLC using a CarbopacPA1 column (Dionex). To assess the progress of the reaction, 20- to 30-ml reaction aliquots are removed at appropriate time intervals and subjected to a prewashed YM-10 microcon centrifugal filter unit to remove RfbB. The flowthrough is then injected into HPLC and the TDP-sugars (5 and 6) are eluted using the conditions described in Section 2.3.1. The retention times for 5 and 6 are 31 to 33 min and 33.5 to 35 min, respectively. The peak for 6 is slightly broader than that for 5. For maximum yields, it is best to use fresh RfbB enzyme that has not been subjected to multiple freeze–thaw cycles. Using this method, we have achieved a reproducible yield of 95% before purification. After the conversion of 5 to 6 is completed, RfbB can be removed by filtration though either an Amicon stirred cell unit equipped with a YM-10 membrane or a larger centrifugal filter unit. The filtrate, which contains 6, can be directly used in subsequent biosynthetic reactions or divided into smaller aliquots and frozen at –80 for future use. Desalting is necessary if the sample is lyophilized and stored in dry form.

3.1. Generation of TDP-2,6-dideoxysugars Another useful TDP-deoxysugar is TDP-4-keto-2,6-dideoxy-a-D-glucose (8), which is produced from TDP-4-keto-6-deoxy-a-D-glucose (6) at an early stage in the biosynthesis of 2,6-dideoxy sugars (Scheme 21.3). The 2-deoxygenation reaction is catalyzed by a group of enzymes called 2-dehydratases (2-DH) that employ a metal ion to convert 6 into a 3,4diketo sugar intermediate 7 (Chen et al., 1999; Draeger et al., 1999). The 3-keto group of 7 has one of two fates in different biosynthetic pathways: it can be reduced by a 3-ketoreductase (3-KR) to a hydroxyl group with either equatorial (8) or axial (9) stereochemistry, or it can be transaminated by a 3-aminotransferase (3-AT) to give a 3-aminosugar (10). Our group employs TylX3 (a 2-DH) and TylC1 (an axial 3-KR from the tylosin biosynthetic pathway of Streptomyces fradiae) to synthesize 9 from 6 (Chen et al., 1999; Takahashi et al., 2006), and TylX3 and KijD10 (an equatorial 3-KR from the kijanimicin biosynthetic gene cluster of Actinomadura kijaniata) to synthesize 8 from 6 (Zhang et al., 2007). In our experience, the generation of the equatorial 3-OH product (8) from 6 is straightforward, while synthesis of the axial 3-OH sugar (9) from 6 is more problematic. In both cases, excess reductase (5 or 2 molar excess, respectively of either TylC1 or KijD10 as compared to TylX3) is used to drive the reaction forward.

532


Cloning, expression and purification of TylX3, TylC1, and KijD10 have been described (Chen et al., 1999; Takahashi et al., 2006; Zhang et al., 2007). In general, these proteins are purified following the protocols given above for RfbA and RfbB. Importantly, 20% glycerol should be included in all purification buffers for TylX3, and 10% glycerol should be present in all reactions performed with TylX3. Due to the instability of the TylX3 product (Scheme 21.3, compound 7), TylX3 should be the last component added to initiate the coupled reactions. Both TylC1 and KijD10 are NADPH-dependent, but many other 3-KR enzymes utilize NADH. 3.1.1. Synthesis of TDP-4-keto-2,6-dideoxy sugars with equatorial C3-OH stereochemistry To synthesize the TDP-4-keto-2,6-dideoxy sugar (8) with equatorial C3-OH stereochemistry, a 3 ml reaction containing 6 (21.8 mM ), NADPH (53 mM ), and 10% glycerol in 50 mM TrisHCl buffer (pH 7.5) is prepared. KijD10 (360 nM ) and TylX3 (160 nM ) are added to initiate the reaction. The reaction mixture is incubated at 25 and the progress of the reaction is monitored by HPLC using the conditions described in section 1.3.1 (the retention times of 6 and 8 are 33 to 34 min and 32 to 33 min, respectively). A degradation product of 7 (maltol, see Scheme 21.3) eluted at 1.7 to 1.8 min is always observed. When freshly purified TylX3 and KijD10 are used, this reaction is complete within 2 to 3 h (after repeated freeze–thaw cycles, KijD10 loses activity). The TylX3/KijD10 coupled reaction is quite efficient and, on average, affords approximately 85% conversion to product (or 20 mg of 8 from 24 mg of 6). With freshly prepared enzyme, we have observed nearly quantitative conversion within 6 h. Prior to its use in any subsequent enzymatic reactions, compound 8 should be purified due to the presence of NADPþ and NADPH in the reaction. The sample (containing 8) should be filtered to remove enzymes and lyophilized to a yellow syrup. Importantly, the frozen sample must be removed from the lyophilizer as soon as it begins to thaw. If lyophilization is allowed to proceed to dryness, there is often substantial (>50%) degradation of the product. To purify 8 using P2 size-exclusion chromatography, the procedure provided in Section 2.3.1 should be followed, and 25 mM NH4HCO3 instead of ddH2O should be used to wash and equilibrate the column, and to elute the sample. A typical flow rate is 6-8 ml/h. Samples with absorbance at 267 nm begin to elute after 24 to 30 h, and continue for the next 12 to 20 h. As described for TDP-a-D-glucose (5), a similar absorbance plateau is commonly observed. However, 8 should be eluted towards the end of the plateau region ( just prior to the reduction of A267 back to baseline) over only 2 to 4 fractions. The fractions at the beginning of the plateau region contain NADPþ and NADPH. Generally, two of the four fractions are greater than 85% pure (as determined by HPLC), while one or two are of lower purity. The desired fractions are combined and


533

lyophilized. Care should be taken not to lyophilize to dryness, as this will cause significant degradation, as observed by 1H-NMR spectroscopy. Usually, approximately 10 mg of 8 (from 24 mg of 6) is recovered from this purification process (Zhang et al., 2007). 3.1.2. Synthesis of TDP-4-keto-2,6-dideoxy sugars with axial C3-OH stereochemistry To generate the TDP-4-keto-2,6-dideoxy sugar 9 with axial C3-OH stereochemistry, we use TylC1 in place of KijD10. A typical 3-ml reaction mixture contains 6 (21.8 mM ), NADPH (53 mM ), and 10% glycerol in 50 mM TrisHCl buffer (pH 7.5). TylC1 (40 mM ) and TylX3 (8 mM ) are added to initiate the reaction. The incubation is carried out at 25 and product formation is monitored by HPLC. Under the HPLC conditions described in Section 2.3.1, the retention times of substrate (6) and product (9) are 33.6 to 34 min and 32.3 to 33 min, respectively. Generation of 9 is not as facile as the C-3 equatorial product (8). We obtain approximately 50% conversion of 6 ! 9, even after overnight incubation. Gel filtration using a P2 column to remove NADPþ and NADPH can be carried out by the method described in Section 3.1.1 for compound 8, and typical yields are about 5 mg of purified sugar (from 24 mg of 6). Compound 9 can often be generated in situ in sufficient quantities to be processed by downstream biosynthetic enzymes.

4. In vitro Reconstitution of Entire Deoxysugar Biosynthetic Pathways Up to this point, we have summarized methodology to synthesize and purify several common (early) intermediates (5, 6, 8, and 9, Scheme 21.3) produced in deoxysugar biosynthetic pathways. These intermediates can be used as starting materials for more elaborate enzymatic syntheses. To date, only a handful of TDP-deoxysugar biosynthetic pathways have been fully reconstituted in vitro using native pathway enzymes. Some of these highly modified TDP-sugars (Scheme 21.4) that have been enzymatically prepared include TDP-a-D-desosamine (11), TDP-a-D-mycaminose (12) (Melancon et al., 2007), TDP-b-L-mycarose (13) (Takahashi et al., 2006), TDP-a-Dforosamine (14) (Hong et al., 2008), TDP-b-L-epivancosamine (Chen et al., 2000), and TDP-b-L-digitoxose (Zhang et al., 2007). Due to the variation in the reactivity and stability of different TDP-sugar intermediates, and to the differences in catalytic efficiencies of the various biosynthetic enzymes involved, there is no general protocol that can be used to synthesize all TDPsugars. Instead, syntheses must be optimized on a case-by-case basis. The fact that few in vitro sugar biosyntheses are known reflects the dearth of biochemical information available for deoxysugar biosynthetic enzymes, and clearly

534

H2N HO


Me

O

Me

DesII O

HO OTDP

O

Me

DesV

H2N

HO OTDP

21

O

DesVI

HO OTDP 23

22

Me Me2N

O

HO OTDP

11

DesI O

Me

O

HO

Me

Tyl1a

HO O

HO OTDP 6

O

Me

HO H2N

TylB

HO OTDP

O

HO

15

16

TylM1

HO Me2N

OTDP

Me

O

HO 12

OTDP

TylX3 SpnO Me

O

O

O

O

TylC1 OTDP

7

Me

HO

9

TylC3

O

O

Me

HO Me

OTDP

TylK

O

17

OH

OH O

Me

OTDP TylC2

O Me

OTDP

Me HO

O

OTDP

Me

18

13

SpnN O

Me

HO 8

O OTDP

SpnQ

O

Me

19

O

SpnR OTDP

H2N

Me

20

O

SpnS

OTDP

Me Me2N 14

O OTDP

Scheme 21.4 In vitro enzymatic synthesis of highly modified TDP-deoxysugars. Biosynthetic pathways for several highly modified TDP-deoxysugars (11 to 14) that have been synthesized in vitro using native pathway enzymes. See text for details.

suggests that more thorough kinetic investigations of these important enzymes are warranted. Here, we present procedures for the enzymatic synthesis of several TDP-deoxysugars (11 to 14) that have been investigated in our laboratory.

4.1. One-pot synthesis of TDP-a-D-mycaminose (12) The synthesis of TDP-a-D-mycaminose (TDP-3-N,N-dimethylamino3,6-dideoxy-a-D-glucose, 12, Scheme 21.4) from TDP-4-keto-6-deoxya-D-glucose (6) can be carried out in a single reaction using three enzymes (Tyl1a, TylB, and TylM1) from the tylosin biosynthetic pathway of S. fradiae (Scheme 21.4). Tyl1a and TylB catalyze the conversion of 6 to TDP-3-amino-3,6-dideoxy glucose (16) through a TDP-3-keto-6-deoxy sugar intermediate (15) (Melancon et al., 2007). TylM1 is an N, N-dimethyltransferase that converts 16 to TDP-a-D-mycaminose (12) (Chen et al., 1998, 2002). Genes encoding Tyl1a, TylB, and TylM1 can be cloned into either pET24b or pET28b, expressed in E. coli as the His6tagged fusion proteins, and purified using standard protocols. For the synthesis of TDP-a-D-mycaminose (12), compound 6, prepared as described in Section 3, is lyophilized and resuspended in 50 mM potassium phosphate buffer, pH 7.5. The reaction mixture contains 6 (1 mM ), 0 L-glutamate (30 mM ), pyridoxal 5 -phosphate (PLP, 150 mM ), S-adenosylL-methionine (SAM, 2 mM ), TylB (30 mM ), and TylM1 (60 mM ) in a total


535

volume of 2 ml of 50 mM potassium phosphate buffer (pH 7.5). Tyl1a (3 mM ) is added to initiate the reaction. It is best to use newly prepared enzymes and a high level of glycerol in the reaction mixture should be avoided. After 12 h at 25 , 40 to 60% conversion of 6 to 12 should be observed by HPLC. Using the HPLC protocol described in Section 2.3.1, retention times for TDP-sugars 12, 16, 6, and 15 are 7 to 8 min, 13 min, 35 to 36 min, and 39 min, respectively. In addition to the TDPsugar peaks, peaks for TDP (41.9 min), (2R,3R)-2-methyl-3,5-dihydroxy4-keto-2,3-dihydropyran, a degradation product of 15 (1.8 min), SAM and its degradation products (3.2 min, 21 min, and 23 min), and S-adenosyl-Lhomocysteine (2.1 min) are also visible. An Adsorbosphere SAX column can also be employed to purify 12 (Chen et al., 2002). This column is especially useful for isolating TDP-amino sugars, which are significantly more polar than most other TDP-deoxysugars (see Section 4.3). After the TDP-a-D-mycaminose (12) peak is collected, the sample can be readily desalted using HPLC for small-scale reactions, or by FPLC for large-scale preparations. When using HPLC, the sample is applied to an analytical C18 column and H2O is used to elute 12 (retention time is 5 to 10 min). For larger-scale preparations, FPLC purification can be carried out using the buffers and column described in Section 2.3.2, applying a linear gradient from 0 to 100% B over 25 min. Under these conditions, TDP-a-D-mycaminose (12) will elute at approximately 12.5 min. During FPLC purification, most of the unused substrate, TDP-4-keto-6-deoxy-aD-glucose (6, retention time 15 min), can be recovered and desalted using the G-10 column. The average yield from a 2-ml reaction mixture for the enzymatic synthesis of TDP-a-D-mycaminose is 11% (0.11 mg of 12 from 1 mg of 6), which is a threefold improvement over the chemical preparation method (Chen et al., 2002).

4.2. Two-stage one-pot synthesis of TDP-b-L-mycarose (13) The functions of each of the enzymes in TDP-b-L-mycarose (13) biosynthesis in the tylosin producer, S. fradiae, have been verified in vitro and are shown in Scheme 21.4 (Chen et al., 1999, 2001; Takahashi et al., 2005). From TDP-4-keto-6-deoxy-a-D-glucose (6), TylX3 and TylC1 catalyze the formation of 9. Following SAM-dependent C3 methylation of 9 by TylC3, compound 17 is converted to TDP-b-L-mycarose (13) by the sequential action of TylK, a 5-epimerase, and TylC2, an NADþ-dependent 4-ketoreductase. Two features make a one-pot synthesis of 13 attractive. First, as noted above, the instability of the TylX3 product (7) requires the presence of a 3-ketoreducatse (TylC1) to drive the reaction forward and to avoid the decomposition of 7 into TDP and maltol (Chen et al., 1999).

536


Second, the presence of the 4-ketoreductase (TylC2) is required in order to drive the formation of 13 from the epimerized TylK product (18), which is in equilibrium with 17 (Takahashi et al., 2005). Thus, a two-stage, one-pot synthesis of TDP-b-L-mycarose (13) was developed, wherein compound 6 is first generated by procedures similar to those outlined in Sections 2.2 and 3 with slight modifications (Takahashi et al., 2006). Following the completion of TTP synthesis from thymidine and ATP, the TM/TMK/NDK/PK enzymes are removed by ultrafiltration. Next, a-D-glucose-1-phosphate (1, 3 mM ) and RfbA (57 mM ) are added to the reaction mixture to convert 1 to 5. After a 30-min incubation period at 30 , RfbB (28 mM ) is added and the mixture is allowed to incubate for 1 h at 37 to convert 5 to 6. At this point, it is not necessary to remove either RfbA or RfbB from the incubation mixture. In the next stage of the reaction, the five TDP-b-L-mycarose (13) biosynthetic enzymes (TylX3/C1/C3/K/C2, 30 mM each) are added along with 6 mM NADPH and 3 mM SAM. The reaction is allowed to proceed for 1 h at room temperature to convert 6 to 13. The cloning, expression, and purification of the mycarose biosynthetic enzymes have been previously described (Chen et al., 1999, 2001; Takahashi et al., 2005). TDP-b-L-mycarose (13) is purified from the reaction mixture by FPLC using a MonoQ 10/10 column that is eluted with H2O over 2-column volumes, followed by a linear gradient from 0-280 mM NH4HCO3 buffer (pH 7.0) over 2-column volumes at a flow rate of 1 ml/min. Following FPLC purification of 13, desalting is carried out using a Sephadex G-10 column (see Section 2.3.3). The identity of 13 is confirmed by 1H NMR spectroscopy and high-resolution MS analysis (Takahashi et al., 2005). The final yield of TDP-b-L-mycarose (13) is 16% from glucose-1-phosphate (1) (Takahashi et al., 2006).

4.3. Multistep enzymatic synthesis of TDP-a-D-forosamine (14) TDP-a-D-forosamine is a highly modified tetradeoxy sugar produced during spinosyn biosynthesis in Saccharopolyspora spinosa (Scheme 21.4). Unlike the preparation of TDP-mycarose (13) from 6, a one-pot synthesis of 14 from 6 is impractical. This is mainly because the SpnQ-catalyzed reaction (8 ! 19) is sensitive to O2 and, therefore, must be conducted using deoxygenated buffers under anaerobic conditions or in the presence of sodium dithionite. We have recently demonstrated the activities for each of the TDP-a-D-forosamine biosynthetic enzymes in vitro (Hong et al., 2008). The activities of SpnO and SpnN (6 ! 7 ! 8) were established in a coupled assay by following the consumption (loss of absorbance at 340 nm) of NADPH by SpnN. Formation of the SpnN product (8) was verified by HPLC analysis as described in Hong et al., 2008. However, the yield of compound 8 is not as high as that obtained in the TylX3/KijD10 reaction


537

described in Section 3.1.1. Thus, for preparative purposes, we recommend using the optimized TylX3/KijD10 system to synthesize 8. SpnQ catalyzes the 3-deoxygenation of 8 to give 19. SpnQ is a [2Fe-2S] cluster containing pyridoxamine 50 -phosphate (PMP)–dependent enzyme, which is a homologue of E1—a mechanistically well-characterized 3-dehydrase from Yersinia pseudotuberculosis (reviewed in He et al., 2000). These 3-dehydrase enzymes also require a reductase component to complete their catalytic cycles, which involve single electron transfer radical chemistry. Because of the sensitivity of the [2Fe-2S] center in SpnQ to O2, the reaction must be carried out anaerobically. A typical SpnQ reaction contains compound 8 (0.7 mM ), PMP (250 mM ), SpnQ (30 mM ), NADPH (0.7 mM ), and a physiological reductase system comprised of either flavodoxin/flavodoxin reductase (30 mM each) or ferredoxin/ferredoxin reductase (30 mM each) in 50 mM potassium phosphate buffer (pH 7.5). Sodium dithionite (0.6 mM ) can also be used as the reductant. Production of 19 is monitored by HPLC (see below). The next step in the pathway is catalyzed by a PLP-dependent 4-aminotransferase, SpnR. The substrate of this enzyme, 19, can be generated in situ by SpnQ using the conditions described above. Following a 3-h incubation period at 24 to convert 8 ! 19, SpnQ, ferridoxin, and ferridoxin reductase are removed by ultrafiltration through a YM-10 membrane. SpnR (37 mM ), PLP (305 mM ), L-glutamate (12.2 mM ), and MgCl2 (1.2 mM ) are then added and the reaction mixture is incubated at 24 for an additional 2 h. The formation of the SpnR product (20) can be monitored by HPLC analysis (see below). The final N-methylation reaction to afford TDP-a-D-forosamine (14), catalyzed by the methyltransferase SpnS, is accomplished by the incubation of the SpnR product (20) with SpnS (10 mM ), SAM (2.0 mM ), MgCl2 (2.0 mM ), and DTT (2.0 mM ) in 50 mM potassium phosphate (pH 7.5) at 37 . Both monomethylated and dimethylated products are observed. The HPLC conditions used to resolve the TDP-a-D-forosamine biosynthetic intermediates are as follows. For the SpnO/N, SpnQ, and SpnQ/R reactions, a Dionex CarboPac PA1 analytical column (4 250 mm) equipped with a CarboPac PA1 guard column (4 50 mm) is employed, and the elution conditions are identical to those given in Section 2.3.1. Using a 1 ml/min flow rate and detection at 267 nm, the HPLC retention times for TDP-sugars 8, 19, and 20 are 33.4, 36.2, and 9.2 min, respectively. The above HPLC conditions cannot resolve the mono- and di-methylated SpnS-catalyzed reaction products from the other assay components. Thus, to purify these sugars, an Adsorbosphere SAX column (5 mm, 4.6 250 mm) is used. Here, a linear gradient from 0 to 20% buffer B (500 mM KH2PO4 buffer, pH 3.5) in buffer A (50 mM KH2PO4 buffer, pH 3.5) over 20 min is applied.

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4.4. TDP-a-D-desosamine (11) TDP-a-D-desosamine is a 4,6-dideoxysugar found in several macrolide antibiotics, including erythromycin, oleandomycin, mycinamicin, methymycin/pikromycin, and megalomicin. The biosynthetic pathway for 11 (6 ! 21 ! 22 ! 23 ! 11, Scheme 21.4) has been fully established through biochemical studies of the pathway enzymes. DesI is a PLP-dependent 4-aminotransferase that converts 6 to 21 (Zhao et al., 2001). DesII, a member of the radical SAM enzyme superfamily, catalyzes the oxidative deamination of 21 to produce 22 (Szu et al., 2005). The PLP-dependent 3-aminotransferase, DesV, transaminates 22 to generate 23 (Szu et al., 2005; Zhao, 2000). The final step, N,N-dimethylation, is catalyzed by DesVI (Chang et al., 2000; Chen et al., 2002) to complete the biosynthesis of 11. Starting from 6, the DesV product (23) can be prepared in a one-pot, two-step reaction from 6 (Szu et al., 2005). In the first step of this synthesis, 6 is converted into 22 by the combined action of DesI and DesII. Due to the sensitivity of the [4Fe-4S] cluster of DesII to O2, and to the fact that this cluster must be reduced for activity, this coupled reaction must be performed anaerobically. Prior to the enzymatic synthesis, the inactive [4Fe-4S]2þ cluster of DesII (190 mM ) must be reduced under anaerobic conditions to the [4Fe-4S]1þ state by sodium dithionite (1.2 mM ) in 100 mM TrisHCl buffer (pH 8.0) for 40 min. The reduction of the [4Fe-4S]2þ cluster can be monitored by following the decrease in absorbance at 420 nm. A 1-ml reaction mixture containing compound 6 (0.6 mM ), L-glutamate (0.5 mM ), PLP (0.14 mM ), DesI (26 mM ), the abovereduced DesII (100 mM ), SAM (0.1 mM ), and DTT (2 mM ) in 100 mM TrisHCl buffer (pH 8.0) is carried out in an anaerobic chamber using degassed buffers. The reaction mixture is incubated at 25 for 3 h. At this point, DesV (100 mM ), along with additional L-glutamate (10 mM ) and PLP (0.8 mM ), are added to the reaction mixture and incubated at 25 for another 30 min. The DesI, DesII, and DesV enzymes are removed by ultrafiltration through a YM10 membrane, and the reaction progress is analyzed by HPLC using a Dionex anion exchange column (4 250 mm). Using a flow rate of 0.6 ml/min, detection at 267 nm, and a linear gradient from 20 to 35% of eluent B (1 M NH4OAc, pH 7.0) in eluent A (H2O) over 30 min, the retention times for compound 6 (the DesI substrate), compound 22 (the DesII product), and compound 23 (the DesV product) are 13.8, 28.2, and 3.7 min, respectively. It should also be noted that a preparative-scale enzymatic synthesis of the DesI product (compound 21) has been reported (Zhao et al., 2001). For the final step of TDP-a-D-desosamine (11) synthesis, a small-scale reaction (50 ml) containing 23 (1.2 mM ), DTT (2 mM ), SAM (10 mM ), and N,N-dimethyltransferase DesVI can be carried out for 3 h at 25 in 50 mM potassium phosphate buffer (pH 7.5). These conditions yield roughly


539

80% conversion of 23 ! 11 (Chen et al., 2002). The reaction products can be purified using an Adsorbosphere SAX column and the elution conditions given in Section 4.3. Under these conditions, the retention times for 23 and 11 are 6.5 and 17 min, respectively. A small amount of an N-monomethylated DesVI reaction intermediate may also be present (retention time 9.3 min). A preparative-scale synthesis of 11 from 23 (Chen et al., 2002) can be performed by incubating 23 (5.7 mM ), SAM (30 mM ), DTT (2 mM ), and DesVI (1.8 mg) in 1 ml of 50 mM potassium phosphate buffer (15% glycerol, pH 7.5) for 3 h at 25 . After removal of DesVI by ultrafiltration (YM-10 membrane), TDP-a-D-desosamine (11) is isolated by sizeexclusion chromatography using a P2 column (2 100 cm) with a 0.5 M NH4HCO3 solution as the eluent. Fractions containing 11 (identified by absorbance at 267 nm) are concentrated and further purified with an FPLC MonoQ HR (10/10) column using a flow rate of 3 ml/min and a linear gradient of 0-0.15 M NH4HCO3 over 15 min to elute compound 11 (retention time 8 min).

5. Synthesis of Deoxysugars In Vivo by Metabolic Pathway Engineering While using purified biosynthetic enzymes to synthesize TDP-deoxysugars in vitro is desirable, this approach may not always be feasible if the sugar biosynthetic enzymes required for a particular synthesis cannot be expressed and purified in suitable quantities, or are not stable. In these cases, an in vivo biosynthetic approach may be useful, wherein a glycosylated natural product is isolated from cultures of a producing bacterial strain, and the deoxysugar of interest is then recovered from the glycosylated product either through reverse GT catalysis or by hydrolysis and chemical derivitization. One method is to heterologously express the deoxysugar biosynthetic gene(s) of interest in a host that naturally produces glycosylated natural products with similar structures. In this case, it may be advantageous to disrupt some of the host’s deoxysugar biosynthetic genes to increase the intracellular concentration of the substrate(s) for the heterologously expressed biosynthetic enzyme(s). Another approach involves the expression of entire biosynthetic pathways in a non-producing, but tolerant strain such as Streptomyces lividans or Streptomyces albus. In these organisms, the heterologously expressed sugar biosynthetic enzymes will not have to compete with endogenous enzymes for their substrate, and the glycosylated products will be excreted from the host cells, limiting their toxicity. It should be noted that synthesis of TDP-deoxysugars in large quantities is not the primary focus of many of the in vivo pathway engineering studies reported in the literature, but the biological systems used in these studies

540


could potentially serve as good sources for TDP-deoxysugar production. Below we highlight some of our recent in vivo studies on deoxysugar biosynthesis and pathway engineering in Streptomyces venezuelae (Thibodeaux and Liu, 2007), an organism that produces several macrolide derivatives bearing D-desosamine moieties. The native biosynthetic pathways for production of D-desosaminylated 12and 14-membered ring macrolactones in S. venezuelae are shown in Scheme 21.5 (6 ! 21 ! 22 ! 23 ! 11 ! 24). In an attempt to generate TDP-a-Dmycaminose (12) in S. venezuelae, we constructed an S. venezuelae double knockout mutant (termed KdesI/desVII) lacking the 4-aminotransferase (DesI) that catalyzes 6 ! 21, and the endogenous desosaminyltransferase (DesVII) that catalyzes 11 ! 24 (Borisova et al., 1999; Melancon et al., 2005). This mutant was designed to enable the accumulation of 6, which could then be processed by heterologously expressed genes from the D-mycaminose pathway of S. fradiae (Scheme 21.5, 6 ! 15 ! 16 ! 12) in order to make 12—a deoxysugar that is not normally produced by S. venezuelae. The tyl1a, tylB, and tylM1 genes were heterologously expressed in this S. venezuelae mutant along with the genes required for mycaminosyltransfer H2N HO

Me

O Me

DesI

O

HO

HO OTDP 21

6

DesII Me

Me

O

HO O

H2N

Me

KdesI/FdtA

O

HO OTDP 23

HO H2N

O O

Me

R2 Me O O

R3

H2N

Me

O

OH

Me O

HO

16

Me

Me O

Me

O Me

Me

DesV

OTDP

Me Me O

Me

O

HO OTDP 27

HO Me

O

HO

Me

Me

Me

O

TylB/DesV Me

O Me

O

R1

HO Me

HO OTDP 15

DesV Me

O

KdesI/Tyl1a

HO OTDP 22

O

O

HO OTDP

OTDP

HO Me O Me HO HO DesVII 2N Me Me HO DesVIII O O DesVI Me2N MeHN HO Me O HO HO OTDP OTDP Me2N DesVII DesVII HO DesVIII DesVIII DesVI

DesVI Me Me2N

TylM1/DesVI

O

HO OTDP

Me

HO Me2N

HO

11

Me2N

O

O

Me

O

HO 24

KdesVII TylM2 TylM3

R1 HO Me2N

HO Me2N

OTDP

12 DesVII DesVIII

Me

O

O

R2

Me

O

HO 25

O

Me HO Me2N

R3

O

R1

HO Me

HO Me2N

HOMe O

O

MeHN

O

O

R1

R2

28 O

R1

HO 29

O

O

R2

HO 26

Scheme 21.5 Manipulating deoxysugar structures by metabolic pathway engineering. A combination of gene knock-out, heterologous gene expression, and precursor feeding experiments allowed the production of several novel compounds in Streptomyces venezuelae. See text for details.


541

(tylM2 and tylM3). When this strain was fed with tylactone (the native aglycone substrate of TylM2/M3), D-mycaminosylated tylactone (25) was generated (Melancon et al., 2005), indicating that the heterologously expressed tyl genes indeed converted 6 into 12. Interestingly, when the tyl1a gene alone was heterologously expressed in a separate S. venezuelae mutant that lacked only the desI gene (Scheme 21.5), novel D-mycaminosylated macrolide derivatives (26) were obtained (Borisova et al., 1999; Melancon et al., 2005). Thus, heterologous expression of a single S. fradiae gene (tyl1a) was sufficient to convert the native D-desosamine biosynthetic pathway into a D-mycaminose pathway in S. venezuelae. From this experiment, it is clear that several of the desosamine biosynthetic enzymes (DesV, DesVI, and DesVII/DesVIII) are capable of processing alternative TDP-deoxysugar substrates (each containing an equatorial 4-OH group that is not present in the natural substrates for DesV-VIII). The above example not only illustrates how deoxysugar biosynthesis can be manipulated in a producing strain by metabolic pathway engineering, but it also reveals the inherent relaxed substrate specificity of many deoxysugar biosynthetic enzymes—a property that should be useful for the synthesis or engineering of deoxysugar structures in other pathways. Indeed, we have exploited the relaxed substrate specificity of the desosamine biosynthetic enzymes to synthesize deoxysugars that have not yet been identified from natural sources (Melancon and Liu, 2007). For example, we expressed fdtA (from the Gram-negative Aneurinibacillus thermoaerophilus), a 3,4-ketoisomerase homologue of tyl1a that produces 27, the C4 epimer of compound 15 (Davis et al., 2007), in the S. venezuelae KdesI mutant. The mutant cultures were found to produce macrolide derivatives bearing either 4-epi-D-mycaminose (28) or 3-N-monomethyl-3-deoxy-Dfucose (29)—two unnatural deoxysugars.

6. Summary The deoxysugar moieties of many glycosylated bacterial secondary metabolites are often essential for the biological activity of these compounds. Changing the structure of these sugar moieties has the potential to generate new glycoforms with altered or improved biological activity. Thus, convenient methods for synthesizing highly modified deoxysugars with defined structures are desirable. Chemical synthesis of deoxysugars is feasible, but is often tedious, technically demanding, and typically suffers from low overall yields. On the other hand, in vitro and in vivo (chemo) enzymatic synthesis of activated nucleotide sugars exploits nature’s biosynthetic machinery and has several benefits over chemical synthesis methods. First, the enzymes used in these methods are a readily available, relatively

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cheap, and renewable resource. Second, enzymes also typically enable more stringent control over the regio- and stereo-chemical outcome of a reaction. Third, these enzymatic reactions are performed under mild, biological conditions and nucleotide sugar purification usually requires only a few simple and familiar chromatographic steps. Here, we have outlined detailed synthesis and purification procedures for several common TDP-sugar intermediates as well as for several specific highly-modified TDP-deoxysugars. It is hoped that the procedures outlined here will provide useful guidelines for the development of synthetic protocols for other unusual sugars. This, in turn, should facilitate the development of workable quantities of glycoforms with defined (and potentially novel) sugar structures.

ACKNOWLEDGMENT We thank the National Institutes of Health (GM35906, GM54346) and the Welch Foundation (F-1511) for their generous support of our research work, and Christian P. Whitman from the College of Pharmacy, University of Texas, Austin, for his critical reading of the manuscript.

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Author Index

A Abe, I., 279 Aboshanab, K., 460, 464, 466, 468, 473, 476, 495 Adam, A., 260 Adams, P. D., 18, 19 Admiraal, S. J., 345, 358, 372 Adusumilli, S., 10 Agnihotri, G., 263, 275, 276, 531, 532, 535, 536, 537 Agusti, G., 272 Ahlbrecht, W., 167, 170, 183 Ahlert, J., 99, 101, 472, 473, 502, 522 Aiba, S., 473 Aidoo, K. A., 230 Airas, K., 386 Ajaz, A. A., 188, 195 Akey, D. L., 21, 27, 31, 37, 39, 264, 271 Alarco, A. M., 11 Albermann, C., 523 Albersmeier, A., 412 Albertini, A. M., 170 Alberts, A. A., 421 Alberts, A. W., 403 Alderwick, L. J., 264 Aldrich, T., 50, 62 Alekseyev, V. Y., 21, 36, 37 Alhamadsheh, M. M., 32 Ali, A., 368, 380, 414 Allen, S., 170 Allmansberger, R., 246 Alon, G., 170, 171 Alt, D., 278 Alt, S., 447 Altabe, S., 410 Altenbuchner, J., 477, 486 Altincicek, B., 358 Altschul, S. F., 192 Alvarez, M. A., 32 Amann, S., 527, 531 Ames, B. D., 17, 21, 35, 374, 381 Amonsin, A., 278 An, J. H., 377 Anand, S., 11, 263, 272, 273, 283 Ancelin, M. L., 188 Anderle, C., 441, 447, 448, 453 Andersen, C., 485 Anderson, C., 170 Anderson, M. S., 404 Anderson, R. J., 260, 265

Andreu, N., 272 Ansari, M. Z., 263, 269, 270, 281 Antoń, N., 216, 217, 218, 223, 224, 225, 237 Aparicio, J. F., 23, 24, 215, 216, 217, 218, 220, 222, 223, 224, 225, 226, 227, 234, 235, 236, 237, 244, 245 Apel, A. K., 237 Appleyard, A. N., 22 Arakawa, K., 171 Arand, M., 199 Aravind, L., 34 Archer, D. B., 50 Arendrup, M., 254 Arikan, F., 167, 170, 183 Arison, B., 145 Armitage, I. M., 424 Arora, P., 263, 268, 269, 270, 276, 282 Arthanari, H., 37 Arthur, C. J., 9, 21, 37, 56, 380, 403 Arya, D. P., 460 Asano, K., 167 Aschauer, H., 407, 411 Ashani, Y., 251 Ashley, G., 30, 322, 329, 344, 348, 350 Ashley, G. W., 23, 24, 119, 329 Asselineau, C. P., 260, 261, 273, 274, 277 Asselineau, J., 261 Astier, J.-P., 484 Astola, J., 276 Asturias, F. J., 18, 19, 21, 23, 24, 25, 32, 39 Asturias, J. A., 226, 227, 237 Atrih, A., 198 Auclair, K., 62, 63 August, P. R., 12, 124 Aune, R., 224 Ausina, V., 277 Austin, M. B., 8, 24, 279, 345 Azad, A. K., 266, 268 Azevedo, V., 170 Azuma, I., 261 B Bachmann, B. O., 11, 99 Backbro, K., 263, 276 Badcock, K., 266, 268, 272, 278, 279, 282 Bae, J., 99 Baerga-Ortiz, Z., 30

545

546 Bai, L., 144, 145, 203, 229, 233, 234, 235, 237, 356, 480 Bai, Y., 264 Bailey, A., 72 Bailey, A. M., 54, 63, 65, 66, 67, 72 Bailey, J. E., 32 Baker, P. J., 398, 419, 425 Balague´, M. M., 217 Balbach, J., 18, 23, 24, 32 Baldock, C., 398, 419, 425 Balibar, C. J., 451 Ballou, C. E., 265 Baltz, R. H., 350 Ban, N., 19, 24, 25, 27, 37, 39, 268 Banchio, C., 346, 355 Banerjee, S., 11, 272, 273, 283 Banerji, N., 278 Bangera, M. G., 403, 422 Bannantine, J. P., 278 Bao, K., 193, 201, 202 Bao, W., 381, 382 Barber, M., 217 Barczak, A. K., 268 Bard, J., 18 Bardou, F., 274, 276 Bardshiri, E., 60 Barends, T. R., 209 Bari, A., 56, 64 Barilone, N., 268 Barnathan, G., 80, 81, 93 Barona-Gomez, F., 345 Barr, P. J., 52, 69 Barreiro, C., 227 Barrow, K. D., 249 Barry, C. E. III, 262, 266, 268, 272, 274, 276, 278, 279, 281, 283 Basak, A., 188, 191, 195, 197 Basnet, D. B., 501 Basset, G. J., 233 Bate, N., 237, 356, 357 Bateman, A., 30, 34, 193, 345, 346 Batist, G., 368 Bau, K., 229 Baumann, S., 358 Beadling, L. C., 50 Beau, J. M., 218 Bechthold, A., 6, 350, 368, 389, 524 Beck, B. J., 136, 222 Beck, E., 358 Beck, J., 50, 51, 145 Bedford, D. J., 52, 296, 344 Beeby, M., 488 Beeman, T. A., 98 Begley, T. P., 501 Bekel, T., 412 Belisle, J. T., 264 Bell, A. A., 57 Bell, R. M., 403 Beltran-Alvarez, P., 9, 56, 377, 379, 380

Author Index

Benhamou, B., 375, 447, 477, 486 Bennett, A. E., 37 Bensadoun, A., 418 Bentley, S. D., 193, 345, 346 Beppu, T., 226, 227 Berge´, J.-P., 80, 81, 93 Berger, J. M., 262, 267 Bergfors, T., 199 Bergler, H., 407, 411 Bergner, A., 473 Berlan, J., 274 Berry, A., 222 Bertero, M. G., 170 Bertozzi, C. R., 262, 263, 264, 271, 272 Bes, M., 170 Besra, G. S., 260, 261, 260, 261, 262, 263, 274, 271, 272, 274, 275, 276, 277, 278, 282, 419, 421, 422 Bessieres, P., 170 Betlach, M. C., 52, 69, 310, 322, 329, 344, 345, 347, 348, 375 Betts, J., 263, 275 Beveridge, T. J., 260 Bevitt, D. J., 10, 191 Beyer, S., 487 Bhatnagar, D., 53 Bhatt, A., 193, 197, 198, 199, 200, 262, 264, 271, 272, 276 Bhatt, K., 262, 271, 276 Bibb, M. J., 197, 202, 245, 246, 249, 346, 347, 348, 349, 355, 373, 375, 381, 413, 414, 441, 442, 445, 447, 472, 479, 499 Bierman, M., 247, 353 Biesiadka, J., 34 Biggins, J. B., 522 Bililign, T., 381 Bingle, L. E. H., 48 Birch, A. J., 47 Birch, H. L., 264 Birch, R. G., 170 Birge, C. H., 406, 422, 423 Bisang, C., 11, 123, 191, 193, 197, 278 Bischoff, N., 412 Bittman, R., 276 Blackaby, A. P., 272 Blanchard, J. S., 276 Blanco, G., 34 Bloch, H., 261, 275 Bloch, K., 408, 409, 420, 421, 422 Blocker, H., 272 Blondelet-Rouault, M. H., 197, 475 Blundell, T. L., 199, 502 Boddy, C. M., 222 Boddy, C. N., 313, 327, 330, 359 Bode, H. B., 341, 523 Bo¨hm, G., 218, 234 Bo¨hm, G. A., 251 Bohm, I. U., 196, 272, 346

547

Author Index

Bolanos-Garcia, V., 199, 200, 201, 204, 205, 206, 209 Bolard, J., 244 Bolotin, A., 170 Bolton, R. C., 273 Bondi, S. M., 69 Bonnefoy, A., 438 Booker-Milburn, K. I., 385 Borchert, S., 170 Bordat, Y., 270, 277, 278, 281 Borgaro, J. G., 415 Borgos, S. E. F., 220, 224, 236, 244, 247, 254, 255, 256, 258 Borisova, S. A., 538, 540, 541 Bornscheuer, U. T., 477 Borovinskaya, M. A., 464 Borowski, E., 229 Borowy-Borowski, H., 229 Borriss, R., 170, 171 Bottova, I., 268 Boursier, L., 170 Bowman, J., 9 Bowman, M. E., 276 Boyd, D. H., 262 Boyne, M. T. II, 145, 156, 160, 167 Braberg, H., 209 Bradford, M. M., 175, 177 Bradley, K. A., 161 Branã, A. F., 11, 387 Branny, P., 197, 475 Brau, B., 246 Braun, C., 464 Braun, L., 9, 84, 95, 99 Brautaset, T., 220, 236, 244, 246, 247, 254, 255, 256, 258 Brayer, G. D., 464 Breen, C., 250 Breiteneder, H., 34 Brendel, N., 125, 127, 132, 172 Brennan, P. J., 261, 263, 264, 273, 275, 276, 277 Brenner, M. B., 272 Breukink, E., 217 Brian, P., 345, 350 Briken, V., 272 Brikun, I. A., 347, 356 Bringmann, G., 447 Broadhurst, R. W., 11 Brodersen, D. E., 464 Brokl, O., 270 Brosch, R., 10, 266, 268, 272, 278, 279, 282 Brown, A. K., 264, 276 Brown, K. A., 345, 350, 353 Brown, M. J., 196, 320, 322, 341 Brown, M. P., 53, 230 Brown, S., 193, 345, 346 Brown, S. D., 412 Brown-Jenco, C. S., 53

Browse, J., 9, 80, 81, 93, 408, 409, 413 Bruheim, P., 224, 254, 255, 256 Brune, I., 412 Bruner, S. D., 37 Brunger, A. T., 18, 19 Brunkhorst, C., 485 Bruno, W. J., 173 Bruton, C. J., 245, 249, 373, 443, 444, 472 Buchholz, F., 12, 13 Bugie, E., 460 Buglino, J., 263, 270 Bujacz, G., 34 Buko, A., 355 Bu’Lock, J. D., 275 Bummer, P., 254 Burkart, M. D., 380, 400, 414, 533 Burns, K. E., 501 Burns, R. N., 253 Burston, S. G., 21, 380, 403 Bushby, N., 65 Buss, A. D., 272 Butcher, P. D., 262, 278, 282 Butcher, R. A., 127, 129, 132, 170 Butler, A. R., 237, 356, 357 Butler, M. J., 356 Butler, M. S., 3, 47 Buttner, M. J., 197, 198, 202, 245, 246, 345, 346, 347, 348, 349, 373, 442, 445, 447, 499 Butts, C. P., 63, 65, 66 Butzke, D., 171, 172 Bycroft, M., 327, 329 Byers, D. M., 419, 425 Byrne, B., 223, 235, 236, 250, 252, 254 Byrne, K. M., 145 Byrom, K. J., 61, 403 Bzymek, K. P., 460, 464 C Caffrey, P., 23, 30, 114, 216, 217, 218, 220, 222, 223, 224, 234, 235, 236, 243, 244, 245, 249, 250, 252, 254 Cai, X., 172, 387, 388 Calderone, C. T., 129, 130 Camacho, L. R., 262, 268, 282 Cambillau, C., 270, 484 Camey, J. R., 69, 321, 323, 327, 330 Campelo, A. B., 229, 233, 235 Campillo, N., 30 Camus, J. C., 278 Canaan, S., 270 Cane, D. E., 10, 21, 22, 23, 24, 25, 27, 32, 36, 37, 39, 60, 64, 69, 152, 167, 177, 187, 188, 195, 196, 202, 203, 204, 205, 206, 209, 214, 272, 323, 327, 329, 330, 341, 345, 348, 356, 358, 372 Capdevila, C., 375, 447, 477, 486 Cardona, P. J., 272

548 Carey, M. R., 398, 404, 416 Carmody, M., 223, 235, 236, 250, 252, 254 Carney, J., 30, 321, 332, 335, 359 Carney, J. R., 145, 321, 325, 326, 329, 331, 345, 358, 359 Carreras, C. W., 380 Carrere, S., 276 Carriuolo, J., 420 Carroll, B. J., 145 Carroll, P., 262, 263, 276 Carter, A. P., 464 Carter, G. T., 133 Carvalho, R., 127, 131, 132, 133, 170 Cary, J. W., 53 Casabona, N. M., 277 Casqueiro, J., 224, 235 Castonguay, R., 117 Cate, J. H. D., 464 Catravas, G. N., 251 Ceballos, E., 218, 220, 234 Celmer, W. D., 188, 195 Cerdeno-Tarraga, A. M., 193, 345, 346 Cernota, W. H., 347, 356 Chadick, J. Z., 18, 19, 23, 24, 32, 39 Chakraborty, T., 412 Chakravarty, B., 19, 37 Chalen, M. P., 72 Challis, G. L., 145, 193, 202, 203, 345, 346, 443, 444 Chalut, C., 269 Chamberlin, J. W., 188 Chamberlin, L., 198, 345 Chami, M., 260, 276 Champney, W. S., 464 Chan, C. H., 418, 419 Chan, J., 276 Chan, Y. A., 143, 144, 145, 156, 160, 167, 490 Chandra, G., 443, 444 Chandran, S. S., 116, 300, 321, 323, 327, 330 Chang, C., 523, 538 Chang, C. W., 523, 534, 535, 538, 539 Chang, P. K., 53 Chang, Y. C., 60 Chang, Z., 125, 230 Charpentier, X., 263, 276 Chater, K. F., 193, 197, 202, 203, 245, 246, 249, 250, 345, 346, 347, 348, 349, 356, 373, 441, 442, 443, 444, 445, 447, 452, 472, 499 Chen, A. Y., 10, 21, 22, 24, 25, 27, 37, 39, 272 Chen, B., 268, 276 Chen, H., 523, 531, 532, 533, 534, 535, 536, 538, 539 Chen, J., 224 Chen, J. C., 21, 37, 39 Chen, S., 188, 229, 233, 234, 235, 237 Chen, X., 531, 532, 535, 536 Chen, X. H., 170, 171 Chen, Y. H., 387

Author Index

Chen, Y. M., 472, 473, 501 Cheng, Q., 34, 375, 380, 381, 382, 384, 388, 389, 414 Cheng, T. Y., 272 Cheng, Y. Q., 11, 127, 131, 165, 166, 167, 168, 170, 171, 182, 183 Cheung, I. K., 18, 19, 23, 24, 32, 39 Chhabra, A., 263, 266 Chiang, Y. M., 266 Chirala, S. S., 19, 37 Chirgadze, D. Y., 502 Chng, C., 251, 253 Choi, C. Y., 32 Choi, K. H., 275, 404, 410, 419, 421, 422 Choi, S. U., 227 Chopra, I., 368 Chopra, T., 11, 259, 261, 263, 272, 273, 276, 283, 411, 412, 413 Choudhry, A. E., 24, 25, 27 Christenson, S. D., 99 Chuck, J.-A., 249, 402 Chung, G. A., 262, 278, 282 Chung, J. Y., 387 Chung, L. M., 144, 145, 356 Chung, Y. S., 481 Churcher, C., 266, 268, 272, 278, 279, 282 Ciorbaru, R., 260 Clade, D., 144 Clardy, J., 170, 340 Clark, D. P., 403, 405, 407, 408, 409 Clark-Lewis, I., 380, 403 Cleland, W. W., 160 Clemens, S., 34 Clemons, W. M., 464 Cleveland, T. E., 53 Clore, G. M., 18, 19 Clough, J. M., 47 Cole, S. T., 266, 268, 272, 278, 279, 282 Coleman, C. M., 270 Coleman, R., 276, 409 Colina, A. J., 218, 220, 226, 227, 234 Collard, J., 438 Constant, P., 262, 268, 270 Converse, S. E., 271 Cooper, D. J., 272 Cooper, H. N., 264 Cooper, S. M., 171 Coppel, R. L., 262, 276 Corre, C., 145, 345 Cortes, J., 10, 11, 22, 114, 191, 193, 196, 197, 218, 234, 278, 296, 320, 322, 327, 329, 341, 345 Costa, A. M. S. B., 72 Costello, C. E., 272 Costet, L., 170 Coughlin, J. M., 165, 172, 183 Cousin, A., 170 Covarrubias, A. S., 263, 276

549

Author Index

Cowtan, K., 19 Cox, J. S., 262, 268, 270, 272 Cox, R. J., 9, 22, 37, 45, 46, 47, 50, 51, 54, 56, 61, 63, 65, 66, 67, 72, 177, 274, 377, 379, 380, 382, 403, 414 Cramer, K. D., 345 Crawford, J. M., 9, 36, 54, 56, 57, 60, 61 Crellin, P. K., 262, 276 Criado, L. M., 229, 233 Cronan, J. E., 395, 398, 399, 400, 399, 403, 406, 407, 408, 409, 410, 411, 412, 416, 418, 419, 420, 421, 423, 425 Cronan, J. E., Jr., 276, 396, 398, 400, 402, 403, 405, 407, 410, 411, 416, 417, 418, 419, 420, 421, 422, 423, 424, 425 Cropp, T. A., 222, 327, 404 Crosby, J., 21, 22, 27, 28, 34, 36, 37, 51, 61, 177, 278, 377, 379, 380, 382, 403, 414 Cruaud, P., 277 Crump, M. P., 21, 27, 28, 34, 36, 37, 63, 65, 66, 380, 403 Cummings, R. D., 264 Cundliffe, E., 237, 356, 357 Cusick, J. K., 420, 423 Cynamon, M. H., 278 Czerwinski, E. W., 188 Czisny, A., 99 Czisny, A. M., 381 D Daffe, M., 260, 261, 260, 263, 268, 270, 274, 276, 277 D’Agnolo, G., 404, 405 Dai, A. Z., 276 Daines, R. A., 24, 25, 27 Dairi, T., 476, 479, 480 Dakrop, O., 414 Dale, G. E., 403 Dalton, E., 262 Damais, A., 170 Dancy, B. C. R., 36, 54 Daniel, J., 262, 263 Daniel, R., 13 D’Arcy, A., 403 Das, B. C., 270 Da Silva, N. A., 69 Dassa, E., 224 Dauter, Z., 19, 22, 30, 55, 403 David, H. L., 277 Davidson, A. L., 224 Davies, J., 350 Davies, J. K., 10 Davis, C. R., 50, 62 Davis, F. P., 209 Davis, M. L., 541 Dawes, S. S., 265 Dawson, M. J., 50, 272

Day, L. E., 188 Dayem, L. C., 321, 325, 358 Deacon, M., 346 DeBont, J. A., 199 Degano, M., 34 Dehesh, K., 405, 421 DeHoff, B. S., 119 de Kruijff, B., 217 Delaney, R., 419 DeLano, W. L., 18, 19 de la Roche, M. A., 425 Delaumeny, J. M., 265, 266 De Lay, N. R., 398, 399, 425 Delort, A. M., 188 Del Vecchio, F., 22 de Meijere, A., 453 de Mendoza, D., 276, 405, 408, 409, 410 Dempsey, C. E., 21, 34, 36, 37 Demydchuk, Y., 11, 191, 193, 197, 198, 199, 200 Deng, Z., 11, 156, 172, 183, 187, 191, 192, 193, 201, 202, 203, 204, 205, 206, 209, 210, 229, 233, 234, 235, 237, 480 Denoya, C. D., 123, 404 Derewenda, U., 30 Derewenda, Z. S., 18, 19, 22, 30, 55, 403 Desai, R. P., 344 De Sousa-D’Auria, C., 276 Dessen, A., 276 Dessoy, M. A., 446 Deutsch, E., 386 De Voss, J. J., 281 de Winde, J. H., 50 Dewolf, W. E., Jr., 411, 423 Dezeny, G., 442 Dharamsi, A., 30, 407 Diacovich, L., 346, 355 Dickens, M. L., 386 Dickens, S., 192, 193, 197, 198, 199, 201 Dijkhuizen, L., 464, 482, 483, 484 Dijkstra, B. W., 209, 482, 484 Dillon, S. C., 30, 34 Diltz, S., 83 DiMare, M., 195 Dimroth, P., 50 Distler, J., 466, 472, 473, 474, 479, 480, 502 Dittmann, E., 11, 360 Dittrich, F., 9, 81, 83, 99, 413 Dolot, R., 34 Domenech, P., 268, 271, 278 Domergue, F., 9, 81, 83, 99, 413 Donadio, S., 10, 114, 166, 174, 177, 191, 266, 296, 305, 306, 341 Dong, H., 11, 191, 192, 201 Dong, S. D., 353, 356, 357, 366 Donovan, F. W., 47 Dorrestein, P. C., 100, 156, 381 dos Reisa, M. C., 68 Doumith, M., 375, 447, 477, 486

550

Author Index

Dover, L. G., 264, 276 Dowd, C. S., 272, 278 Dowding, J. E., 474 Doyle, T. W., 98 Draeger, G., 531 Drager, G., 527, 531 Dra¨ger, G., 472, 530 Draper, P., 260, 261 Dreier, J., 22, 379 Drepper, A., 486 Drost, G., 217 Du, L., 175, 179, 182 Du, L. C., 55, 92, 101, 107 Duartea, R. T. D., 68 Dubendorff, J. W., 490 Dubey, V. S., 270, 276, 277, 278, 281 Dubochet, J., 260 Dubos, R. J., 270 Ducasse, S., 263, 276 Duncan, K., 262 Dunn, J. J., 490 Dupuis-Hamelin, C., 438 Duthoy, S., 278 Dutler, H., 29 Dyer, U. C., 195, 198 E Eads, J. C., 488 Ebeling, A., 407, 411 Eberl, M., 358 Ebert, A., 473 Ebert-Khosla, S., 34, 340, 344, 345, 368, 373, 375, 413 Ebizka, Y., 60 Ebizuka, Y., 52, 53, 58, 59, 171, 194, 345, 385, 387 Edwards, D. J., 55, 92, 101, 107, 126, 127, 129, 175, 179, 182 Edwards, P., 405, 421 Eggeling, L., 264, 276 Eguchi, T., 477, 478, 493, 501, 502, 503, 504, 505, 507, 508, 509, 511 Ehrlich, K. C., 36, 56, 60, 61 Eichenseer, C., 486 Eiglmeier, K., 266, 268, 272, 278, 279, 282 Elass, E., 260 Eley, K. L., 65, 67 Elling, L., 472, 524, 527, 530, 531 Ellingsen, T. E., 220, 224, 236, 244, 246, 247, 248, 254, 255, 256, 258 Ellison, E., 268 El-Sayed, A. K., 133, 171 Emorine, L., 263, 276 Emsley, P., 19 Enrı´quez, L. L., 224 Ensergueix, D., 262, 268, 282 Eramian, D., 209

Ercolani, K., 18 Eshed, R., 68 Essen, L. O., 37 Eswar, N., 209 Eusta´quio, A. S., 441, 452 Evans, D. A., 195 Evans, S. E., 21, 380, 403 Eveland, S. S., 406 Everett, M. J., 262, 278, 282 Eyles, S. J., 272 Eynard, N., 263, 276 F Facciotti, D., 9, 81, 83, 99, 413 Fahey, R. C., 460, 464 Famet, C. M., 99 Fan, Q., 505 Farnet, C. M., 11, 99, 170, 171 Fecik, R. A., 21, 27, 31, 37, 39 Fernandes, N. D., 266, 268 Fernandez, E., 34, 387 Ferreira, F., 34 Ferrer, J. L., 279 Ferreras, J. A., 263, 268, 270 Ferroud, D., 438 Fice, D., 419 Fichtlscherer, F., 99 Fiebig, K. M., 30, 407 Fiedler, H.-P., 145, 438 Fillmore, J. P., 65 Findlow, S. C., 21, 36, 37, 61, 380, 403 Finking, R., 340, 360, 399, 414 Finnan, S., 249, 250 Fischbach, M. A., 10, 170, 296 Fischer, C., 8, 388 Fischer, I., 477 Fischl, A. S., 400 Fish, S., 356 Fitzgerald, M. C., 57 Fitzmaurice, A. M., 266 Fjaervik, E., 244, 256 Flardh, K., 345 Flatman, R. H., 438 Flatt, P. M., 478, 479, 480, 495 Flett, F., 246, 247, 347, 348 Flood, E., 249, 250 Floriano, B., 441 Florova, G., 327, 422 Floss, H. G., 36, 144, 145, 203, 236, 356, 466, 479, 480, 488, 531 Flugel, R. S., 52, 151, 175, 179, 182, 373, 398, 399, 414, 425 Fonstein, L., 99 Forenza, S., 98 Fortman, J. L., 341 Foster, G. D., 72, 272 Foster, P. G., 21, 37, 39

551

Author Index

Foster, S. J., 198 Fotso, S., 254 Fouces, R., 218, 223, 224, 225 Fowler, C. B., 200 Fowler, K., 202, 203, 443, 444 Franco-Domıńguez, E., 227, 237 Frank, M. W., 412 Franke, P., 170, 171 Fredrick, K., 464 Freel Meyers, C. L., 450, 522 Freitag, A., 449 Frey, P. A., 503 Fridman, M., 389, 451 Frigui, W., 10 Frimodt-Mller, N., 254 Fritsch, E. F., 499 Fritsche, E., 473 Fritzler, J. M., 172 Fritzsche, K., 33 Frost, E., 11, 191, 193, 197 Frueh, D. P., 37 Frutos, R., 170 Fryhle, C. B., 236 Fu, H., 22, 23, 24, 32, 34, 322, 329, 344, 350, 351, 356, 357, 366 Fu, J., 349, 360 Fu, Q., 523 Fu, X., 523 Fuchs, K., 407 Fuchsbichler, S., 407, 411 Fucini, P., 464 Fujii, I., 52, 53, 58, 59, 60, 171, 194, 387 Fujii, T., 478, 507, 508 Fujiwara, N., 276 Fujiwara, T., 495, 500 Funa, N., 59, 263, 279 Fungarob, M. H. P., 68 Furibata, K., 145 Furlaneto, M. C., 68 Furlanetoc, L., 68 Fusetani, N., 171, 172 G Gabriel, D. W., 170 Gafoor, I., 72 Gaisser, S., 272, 345 Gaitatzis, N., 126, 134, 360 Gajhede, M., 34 Gallimore, A. R., 197, 198, 199, 200 Galloway, I. S., 23, 24, 222, 327, 329 Galm, U., 3, 98, 441, 446 Gande, R., 264, 276 Gandecha, A. R., 237, 356, 357 Gao, Q., 99 Gao, Q. J., 99, 101 Garcia-Barcelo, M., 273 Garcia-Bernardo, J., 22

Gardella, T., 416 Garneau-Tsodikova, S., 448, 451 Garnier, T., 10, 263, 264, 268, 274, 275, 278 Garwin, J. L., 404, 405, 417, 418, 419, 421, 424 Gas, S., 266, 268, 272, 278, 279, 282 Gastambide Odier, M., 265, 266 Gatto, G. J., Jr., 389 Gawron, L. S., 98 Gebhardt, K., 145 Geders, T. W., 129 Gehring, A. M., 52, 151, 161, 175, 179, 182, 373, 399, 414 Gelmann, E. P., 408, 422 Georgopapdakou, N. C., 50 Gerth, K., 145, 360 Geurra, S. M., 224 Gewain, K. M., 442 Ghoorahoo, H. I., 193 Ghosal, A., 19, 21, 25, 32 Ghosh, R., 263 Gibbons, P. H., 442 Gibert, I., 272 Gibson, E., 236, 252, 254 Gibson, K. J., 264, 276, 278, 282 Gibson, M., 388 Gibson, T., 350 Gibson, T. J., 174, 177 Gicquel, B., 262, 266, 268, 277, 278, 281, 282 Gil, J. A., 220, 223, 224, 228, 229, 230, 233, 235, 236 Gilles, C., 484 Gillig, J. R., 19, 30, 34, 408, 409 Gillois, M., 277 Gilroy, J., 398, 419, 425 Giner, J. L., 98 Giovannoni, J. J., 233 Giquel, B., 272 Giraldes, J. W., 21, 27, 31, 37, 39 Giri, R., 511 Glass, S. N. L., 50 Gleeson, C., 262 Glod, F., 65, 414 Glushka, J., 277 Goeke, K., 482, 486 Goesmann, A., 412 Goff, R. D., 451 Gokhale, R. S., 11, 21, 23, 25, 37, 39, 152, 167, 206, 259, 261, 263, 266, 268, 270, 271, 272, 273, 276, 279, 281, 283, 323, 327, 329, 372, 411, 412, 413 Goldberg, I. H., 98 Golding, B. T., 217 Golik, J., 98, 229 Gomi, K., 52, 58 Gonzalez, M. C., 347, 356 Goodfelllow, M., 274 Gordee, E. Z., 188 Gordon, S. V., 266, 268, 272, 278, 279, 282

552 Goren, M. B., 260, 265, 270, 273 Gorman, M., 188 Gosain, A., 263 Goss, R. J., 313 Goss, R. M., 199, 201, 204, 205, 206, 209 Gottschalk, D., 360 Gottschalk, G., 170, 171 Gould, S. J., 345 Graham, I. A., 80, 93 Graham, S., 263, 276 Gramajo, H., 170, 319, 321, 332, 335, 339, 345, 346, 355, 356, 358, 359, 372, 414 Gramajo, H. C., 346 Grammel, N., 170, 171 Grana, D., 416 Grant, E., 272 Grasdalen, H., 224 Greenwell, L., 387 Gregory, J. F. III, 233 Gregory, M. A., 253 Griesinger, C., 453 Griffin, R., 268 Griffith, B. R., 523, 524 Griffiths, G., 260 Grimaldi, C., 276 Grisebach, H. G., 469, 472 Grogan, D. W., 396 Grond, S., 145, 167, 170, 183 Grondin, S., 278 Grooms, M., 24, 25, 27 Gros, P., 18, 19 Gross, F., 349, 360 Grosse-Kunstleve, R. W., 18, 19 Grubelnik-Leiser, A., 346 Gru¨nanger, C., 145 Gu, L., 125 Gu, Z., 19, 37 Gubernator, J., 279 Guerardel, Y., 260 Guerra, D., 83 Guerra, D. J., 408, 409 Guerra, S. M., 218, 225, 237 Guilhot, C., 262, 268, 270, 276, 282 Guirado, E., 272 Gulder, T., 447 Gulliksen, O. M., 244, 256 Gumieniak, J., 229 Gumila, C., 188 Gunawardana, G., 355 Gu¨nther, N., 472, 530 Guo, Z., 531, 532, 534, 535, 536 Gupta, S., 11, 263, 272, 273, 283 Gurcha, S. S., 263, 264, 272, 272, 276 Gurgui, C., 11 Gurvitz, A., 263, 276 Gust, B., 202, 203, 441, 442, 443, 444, 447, 448, 452, 453 Gustafsson, C., 322, 329, 344

Author Index H Haas, M. J., 474 Hadfield, A. T., 21, 27, 28 Hafner, E. W., 404 Haga, K., 482, 484 Hager, M. H., 3, 98 Hain, T., 412 Hakala, J., 386 Hallberg, B. M., 199 Hallis, T. M., 535, 536 Halo, L. M., 63, 65, 66, 67, 72 Halpern, A. L., 173 Hamill, R. L., 188 Han, L., 404, 421 Hanamoto, A., 193, 225 Hanawa-Suetsugu, K., 464 Handa, S., 236 Handelsman, J., 145, 156, 160, 167 Handschuh, L., 34 Hanefeld, U., 327, 329 Hans, A., 170 Hanson, A. D., 233 Hanson, J. R., 47 Hara, M., 167 Hara, O., 226 Hara, S., 58 Harada, S., 387 Harbacheuski, R., 268 Harding, J. R., 65 Hardt, I. H., 133 Harley, K., 54, 66 Harper, D., 193, 345, 346 Harris, B., 278 Harris, D., 266, 268, 272, 278, 279, 282 Harris, D. E., 193, 345, 346 Harris, J. M., 464 Harsulkar, A., 81 Hartsel, S., 244 Harvey, B. M., 11, 191, 192, 193, 197, 198, 199, 200, 201, 204, 205, 206, 209, 210 Hase, A., 83 Hasegawa, M., 476 Haser, R., 490 Hashimoto, E., 476 Hasler, H., 188, 195 Hattori, M., 193, 225 Haugan, K., 246, 248 Hauvermale, A., 83 Hayashi, H., 503 Haydock, S. F., 10, 11, 23, 24, 119, 191, 192, 193, 197, 198, 199, 201, 210, 218, 222, 234, 237, 301, 502 Haygood, M., 170 He, J., 11, 229, 230, 233, 235, 237 He, W., 206 He, X., 524, 537, 538

Author Index

Heath, R. J., 32, 400, 401, 404, 406, 407, 408, 409, 411, 420, 421, 422, 423 Heathcote, M. L., 199, 201, 204, 205, 206, 209 Heckman, K. L., 302 Heide, L., 437, 438, 441, 442, 446, 447, 448, 449, 450, 452, 453 Heinzel, P., 474 Hemker, M., 482, 486 Henderson, B. S., 19, 30, 34, 408, 409 Henderson, J., 409 Hendrickson, L., 50, 62 Heneghan, M., 63, 65, 66, 72 Heneghen, M. N., 72 Hennig, S., 448 Hensel, M., 262 Hensens, O. D., 98 Hertweck, C., 6, 9, 11, 33, 34, 50, 83, 93, 124, 172, 368, 389 Higgins, D. G., 174, 177 Hilde, W., 50 Hildebrand, M., 170 Hill, E. A., 36, 54 Hill, J. A., 21, 27, 28 Hill, R. L., 419 Hill, S., 145 Hiltunen, J. K., 262, 276 Hintz, M., 358 Hirata, A., 262, 279 Hiratsu, K., 171 Hirayama, T., 501 Hitchman, T. S., 22, 54, 61, 177, 403 Hitzeroth, G., 170, 171 Ho, S. N., 55 Hoang, T. T., 420, 423 Hochmuth, T., 11 Hoette, I., 217 Hoffman, D., 144 Hoffmeister, D., 50, 387 Hofle, G., 272 Hofmann, C., 350 Hofmann, J., 411, 412 Hogenauer, G., 405, 407, 411 Holden, D. W., 262 Holden, H. M., 541 Holland, D. R., 407, 411 Holmes, D. J., 410, 423 Holmes, D. S., 195, 198 Ho¨ltzel, A., 145 Holzbaur, I. E., 117 Holzenka¨mpfer, M., 145 Hong, H., 11, 22, 99, 191, 192, 193, 199, 201, 204, 205, 206, 209, 210, 382 Hong, J. S., 503, 505, 515 Hong, L., 530, 532, 533, 534, 536 Hong, S. T., 345 Hong, Y. S., 411, 438 Hoogerheide, J. C., 217 Hopke, J., 345, 350, 353

553 Hopwood, D. A., 21, 32, 34, 35, 36, 37, 52, 197, 202, 229, 230, 245, 246, 249, 266, 321, 323, 327, 330, 340, 344, 345, 346, 347, 348, 349, 351, 368, 373, 375, 381, 387, 413, 414, 442, 445, 447, 472, 474, 499 Hori, M., 503 Horinouchi, S., 226, 227, 262, 279 Horinuchi, S., 59 Horsman, G. P., 97 Horstmann, N., 167, 170, 183 Horton, R. M., 55, 302 Hosomi, Y., 495, 500 Hosted, T. J., Jr., 522 Hothersall, J., 171 Hotta, K., 191, 359, 510 Hou, X., 506 Houssin, C., 260, 276 Houston, S., 30, 407 Howells, A. J., 438 Hu, H., 229 Hu, Z., 36, 69, 118, 310, 344, 348, 349, 351, 353, 356, 364 Huang, F., 501, 502, 504, 505, 506, 510, 511 Huang, G., 170 Huang, J., 351, 353 Huang, K., 99 Huang, K. X., 99 Huang, X., 229, 233, 235, 237 Huang, Y.-S., 81 Huang, Z., 501 Hubbard, B. K., 533 Hubbard, B. R., 188, 195 Huber, D., 416 Huber, R., 473 Hudson, A. T., 275 Hughes-Thomas, Z. A., 11, 191, 193, 196, 197, 199, 201, 204, 205, 206, 209, 346 Hui, D., 171, 172 Hu¨lsmann, A., 485 Humm, A., 473 Hunt, H. D., 55 Hunziker, D., 206 Hur, G. H., 380, 414 Hurlbert, J., 25 Hurley, D., 65 Hurley, J. H., 34 Hurtubise, Y., 485 Hutchinson, C. R., 21, 23, 24, 30, 32, 34, 62, 63, 195, 322, 344, 348, 349, 350, 351, 353, 356, 364, 368, 372, 380, 381, 382, 386, 387, 388, 414 Hwang, S. J., 327 Hwang, Y. I., 227 Hwangbo, Y., 398, 425 Hyun, C. G., 381

554

Author Index I

Ichinose, K., 171, 345, 385 Iimura, Y., 58 Ikeda, D., 503 Ikeda, H., 119, 123, 145, 193, 225 Imanaka, T., 473 Inomata, K., 477, 478, 502 Inoue, S., 188 Ipsen, H., 34 Irschik, H., 167, 170, 183 Ishida, K., 11, 33 Ishida, N., 400 Ishii, T., 171 Ishikawa, J., 191, 193, 225 Ishiyama, A., 188 Isogai, A., 145 Itoh, S., 468 Itoh, T., 385 Itou, K., 345 Iwasaki, S., 99 Iwase, N., 501 Iyer, L. M., 34 Izumikawa, M., 194, 279, 375, 381, 382, 384, 388, 389 J Jackowski, S., 398, 403, 405, 416, 418, 424 Jackson, M., 260, 266, 268, 278, 279, 282, 355 Jackson, P. D., 418 Jacobs, W. R., Jr., 262, 268, 270, 272, 276, 277 Jacobsen, J. R., 23, 25, 167, 313, 348, 372 Jahn, D., 360, 414 Jain, M., 262, 263, 270, 271 Jakimowicz, D., 443, 444 Jakobsen, O. M., 247, 255 James, K. D., 193, 345, 346 Jansen, R., 167, 170, 183 Janson, C. A., 24, 25, 27 Janssen, D. B., 209 Jansson, A., 21, 204, 386, 387 Jaoua, S., 308 Jarlier, V., 260 Jarzabek, M. E., 414 Jaskolski, M., 34 Jaworski, J. G., 424, 425 Jay, M., 254 Jayaraj, S., 323 Jekel, P. A., 209 Jeminet, G., 188 Jencks, W. P., 420 Jenke-Kodama, H., 11, 360 Jenkin, G. A., 10 Jenni, S., 19, 37, 272 Jeong, K. L., 229, 233, 235, 237 Jeong, T. S., 438 Jeschek, C., 407

Jiang, H., 9, 79, 84, 95, 99 Jiang, J., 522, 523 Jiang, J. S., 18, 19 Jiang, Y., 418, 419 Johnson, P. D., 10 Jomaa, H., 358 Jones, M. A., 11, 191, 193, 197, 199, 201, 204, 205, 206, 209 Jones, M. D., 262 Jones, T. A., 199, 263, 276 Joost van Neerven, R. J., 34 Jordan, P. M., 50, 63 Jornvall, H., 28 Joshi, A. K., 18, 19, 21, 23, 24, 25, 29, 32, 39, 55, 64, 275 Joshi, K., 81 Ju, J., 3, 172, 183 Ju, J. H., 98, 100 Julian, E., 272, 276 Julien, B., 131, 170, 308, 314, 315, 345 Jung, M. E., 387, 388 Jung, W. S., 503, 505, 515 K Kadziola, A., 19, 24, 25, 27, 490 Kahne, D., 389, 448, 450, 451 Kaiser, O., 170, 412 Kakavas, S. J., 355 Kakinuma, K., 477, 478, 495, 500, 501, 502, 503, 511 Kalaitzis, J. A., 34, 345 Kalinowski, J., 412 Kalk, K. H., 482, 484 Kallberg, Y., 28 Kallenborn, H. G., 145 Kallender, H., 263, 275, 276 Kallio, P., 21, 377, 386, 388 Kaminishi, T., 464 Kammerer, B., 441, 442, 446, 447, 448, 453 Kampers, T., 485 Kana, B. D., 265 Kang, J. S., 438 Kang, S. H., 438, 500, 516 Kanjilal, S., 278 Kao, C. M., 123, 196, 251, 253, 320, 327, 341, 344, 349, 351, 353, 357 Kaplan, G., 268, 272, 279 Kaplan, L., 145 Kapur, V., 278 Karchin, R., 209 Kase, H., 468, 484 Kastaniotis, A. J., 263, 276 Katayama, K., 21, 386 Kato, Y., 145, 308, 314, 315, 356 Katsumata, S., 167 Katz, E., 474

Author Index

Katz, L., 10, 113, 114, 119, 144, 145, 166, 174, 177, 191, 196, 266, 296, 320, 341, 344, 349, 353, 355, 356, 357, 366 Kaulmann, U., 9, 83, 93 Kauppinen, S., 404 Kawamoto, I., 167 Kawazoe, M., 464 Kayser, O., 244 Kealey, J. T., 52, 69, 321, 324, 325, 331, 358 Kealey, T. J., 69 Keating, D. H., 398, 416 Keating, T. A., 161, 280, 281 Keatinge-Clay, A. T., 21, 22, 24, 25, 26, 27, 28, 30, 32, 34, 72, 117, 298, 306, 368, 376 Keithly, J. S., 172 Kelemen, G. H., 345 Kelleher, N. L., 9, 36, 57, 100, 145, 156, 160, 167, 381 Kelleher, N. P., 50 Kellenberger, L., 29, 327, 329 Keller, J. M., 265 Kelly, L., 209 Kelly, T. M., 406 Kemperman, R. F., 80 Kendrew, S. G., 11, 22, 62, 191, 193, 197, 386, 387 Kennedy, D. R., 98 Kennedy, E. P., 400 Kennedy, J., 12, 62, 63, 321, 324, 325, 326, 331, 358, 359 Kevany, B. M., 144, 145 Khalameyzer, V., 477 Khandekar, S. S., 24, 25, 27 Kharel, M. K., 489, 501, 502 Khaw, L., 23, 24 Khaw, L. E., 218, 234, 237, 251 Khosla, C., 10, 21, 22, 23, 24, 25, 26, 27, 32, 34, 36, 37, 39, 52, 60, 64, 69, 72, 145, 151, 152, 167, 175, 177, 179, 182, 196, 206, 214, 272, 306, 320, 323, 327, 329, 330, 340, 341, 344, 345, 346, 348, 349, 355, 356, 357, 358, 359, 368, 372, 373, 375, 376, 379, 380, 381, 382, 383, 384, 385, 399, 413 Kiderlen, A. F., 244 Kieser, H., 193, 201, 202, 245, 246, 345, 346, 347, 348, 349 Kieser, H. M., 245, 249, 373, 472 Kieser, T., 197, 202, 203, 229, 245, 249, 302, 303, 304, 309, 373, 442, 443, 444, 445, 447, 472, 499 Kikuchi, H., 193, 225 Kim, B.-G., 501 Kim, B. S., 222, 327 Kim, C. G., 236, 438 Kim, C. Y., 21, 22, 24, 25, 27, 37, 39, 60, 64, 69, 383 Kim, D. H., 438, 481 Kim, E. S., 345

555 Kim, H. J., 264 Kim, J. Y., 500, 516 Kim, K. R., 480 Kim, K. W., 438 Kim, S. O., 480 Kim, W., 314 Kim, W. G., 412 Kim, Y., 398, 425 Kim, Y. S., 356, 377 Kimber, M. S., 30, 407 Kimberley, M. R., 385 Kinashi, H., 171 Kinoshita, K., 206, 313 Kinoshita, S., 478, 507, 508 Kinsland, C. L., 501 Kirayama, K., 495, 500 Kirschning, A., 472, 527, 530, 531 Kitade, Y., 400 Kitamura, S., 484 Kitani, S., 172, 227 Kittendorf, J. D., 21, 27, 31, 37, 39, 136 Klages, A. L., 404, 417, 419, 421 Klages Ulrich, A., 405, 410 Klassen, J. B., 421 Klich, M., 438 Klimowicz, A. K., 145, 156, 160, 167 Klohr, S. E., 98 Kluepfel, D., 485 Knauf, V., 9, 81, 83, 99, 413 Kobayashi, H., 99 Kobayashi, K., 272 Kobayashi, S., 375, 376, 379, 383 Kodama, K., 171 Kodumal, S. J., 321, 348, 350, 358 Koglin, A., 37 Kogure, T., 501 Kohana, A., 188 Kohli, R. M., 37 Koike-Takeshita, A., 500 Koizumi, K., 145 Kojima, K., 501 Koketsu, K., 359 Kolattukudy, P. E., 262, 263, 264, 265, 268, 270, 271, 272, 278, 280 Kollas, A., 358 Kollenz, G., 407, 411 Kolodzi-Ejczyk, P., 229 Kolter, R., 170 Kolvek, S. J., 345, 350, 353 Kominek, L. A., 442 Kondo, K., 227 Kondo, S., 510 Ko¨nig, A., 23, 24, 218, 222, 234, 237 Konishi, M., 99 Konz, D., 360, 414 Koonin, E. V., 34 Koorbanally, N. A., 506, 510 Kopp, M., 170, 308

556

Author Index

Koppisch, A. T., 368, 382, 384 Kordulakova, J., 268 Korman, T. P., 21, 27, 28, 29, 30, 35, 384, 385 Korotkova, N., 355 Koskiniemi, H., 387 Koumoutsi, A., 170, 171 Kozubek, A., 278 Krastel, P., 145 Kreiswirth, B. N., 268 Kremer, L., 260, 260, 272, 273, 419, 421, 422 Krishnan, B., 98 Krithika, R., 281 Kroken, S., 50 Kroutil, W., 199, 201, 204, 205, 206, 209 Krucinski, J., 21, 37, 39 Krug, D., 171 Kruger, R. G., 389 Kruh, N. A., 415 Krumbach, K., 262, 263, 276 Kubo, Y., 58 Kudo, F., 206, 477, 478, 493, 495, 500, 501, 502, 503, 504, 505, 507, 508, 509 Kuhstoss, S., 119 Kula, M.-R., 530, 531 Kull, A., 29 Kumar, P., 262, 272, 296 Kumasaka, T., 501 Kuner, J., 83 Kunieda, K., 345 Kunimoto, S., 188 Kunst, F., 170 Kunze, B., 272 Kurepina, N., 268 Kuriki, H., 502 Kurth, D., 346, 355 Kurumbang, N. P., 503 Kusunose, M., 265 Kuszewski, J., 18, 19 Kuwahara, M., 477, 478, 502 Kwak, J. H., 411 Kwon, H.-J., 480, 500, 516, 531, 532, 533, 550 Kwon-Chung, K. J., 60 L Laatsch, H., 254 Labesse, G., 263, 276 Lacave, C., 273 LaCelle, M., 52, 151, 175, 179, 182, 373, 399 Lackner, P., 34 Lacroix, C., 278 LaGier, M. J., 172 Lai, C. Y., 403, 406 Laila, P., 172 Laiple, K. J., 167, 170, 183 Lam, K. S., 98 Lamb, S., 522 Lamba, D., 34

Lambalot, R. H., 52, 151, 152, 175, 179, 182, 343, 373, 398, 399, 425 Lamichhane, J., 438 Lancelin, J. M., 218, 254, 255, 256 Lane, J. M., 262 Laneelle, G., 261, 262, 265, 268, 274, 276, 277 Laneelle, M. A., 268, 270, 274, 276, 277 Langenhan, J. M., 524 Lantz, A. E., 441 Lardizabal, K., 9, 81, 83, 99, 413 Large, S. L., 357 Larrabee, A. R., 400 Larrouy-Maumus, G., 260 Larsen, J. N., 34 Larsen, S., 19, 24, 25, 27 Larson, T., 80, 93 Larson, T. J., 403, 404 Laskowski, R. A., 19 Lassaigne, P., 438 Lassner, M., 9, 81, 83, 99, 413 Latif, M., 350 Lau, J., 22, 23, 24, 117, 177 Laval, F., 266 Lawson, D. M., 438 Lazarus, C. M., 48, 50, 54, 63, 65, 66, 67, 72 Leach, B. I., 347, 356 Leadlay, P. F., 10, 11, 12, 22, 23, 24, 30, 99, 191, 192, 193, 196, 197, 198, 199, 200, 201, 204, 205, 206, 209, 210, 218, 222, 237, 251, 271, 278, 296, 320, 322, 327, 329, 340, 341, 345, 346, 505 Leaf, T., 344, 356 Leary, J. A., 262, 263, 271, 272 Lea-Smith, D. J., 262, 276 Leavell, M. D., 262, 268, 272 Lebrihi, A., 197, 475 Lederer, E., 260, 261, 266, 267, 270, 274 Lee, C. K., 227 Lee, C. M., 262 Lee, D. H., 263, 271 Lee, H. C., 481, 489, 501, 502, 530 Lee, H. Y., 383 Lee, I.-K., 523 Lee, K. J., 356 Lee, K. M., 227 Lee, R. E., 10, 274 Lee, S., 411, 466, 479, 480 Lee, S.-G., 530 Lee, S. Y., 229, 233, 235, 237 Lee, T. S., 32, 375, 376, 379, 381, 383 Lee, Y. S., 381 Leemhuis, H., 464, 482, 483, 484 Leesong, M., 19, 30, 34, 408, 409 Lei, M., 147, 152 Leibundgut, M., 19, 24, 25, 27, 37, 39 Leigh, C. D., 262 Leimkuhler, C., 389 Leistner, E., 144

557

Author Index

Leitinger, B., 407, 411 Lemassu, A., 260, 270, 277 Lemke, A., 244 Lengeler, J., 475 Lennarz, W. J., 422 Lenz, J., 482, 486 Leonard, A. E., 81, 83 Leroy, Y., 260 Lesjean, S., 262, 276 Lester, J. B., 11, 191, 192, 193, 197, 198, 199, 201, 218, 234 Lever, M., 251 Levering, C., 9, 81, 83, 99, 413 Levy, S. B., 407, 411 Lewis, T., 9 Li, J., 21, 30, 37, 202, 203, 205, 206, 209 Li, L., 278, 480, 523 Li, Q., 21, 22, 37 Li, S. M., 441, 446, 448, 449, 450, 452 Li, W., 156 Li, W. B., 276, 409 Li, W. L., 100 Li, X., 350 Li, Y., 34, 422, 502, 506, 510, 511 Liang, T.-C., 188, 195 Liang, X. Z., 101 Liao, J., 523 Licari, P., 344, 356 Lidstrom, M. E., 355 Light, R. J., 422 Ligon, J., 130, 145 Lill, R., 22 Lim, S.-K., 165, 503, 505, 515 Lima, C. D., 267, 268 Limpkin, C., 21, 27, 28 Lin, D. C., 421 Lin, E. C. C., 475 Lin, F. L., 262, 263, 271, 272 Lin, X., 203, 204, 205, 206, 209 Lindquist, Y., 19, 24, 25, 27 Lindsay, Y., 327 Linenberger, M., 98 Linz, J. E., 54 Liou, G. F., 22, 23, 177 Liou, K., 481, 489, 501, 502, 503, 530 Lipata, F., 388 Lipman, D. J., 192 Liras, P., 224, 226, 227, 235, 237 Liu, C. W., 21, 36, 37 Liu, H., 170, 355 Liu, H.-W., 521, 522, 523, 524, 530, 531, 532, 533, 534, 535, 536, 537, 538, 539, 540, 541, 550 Liu, J., 193, 201, 202 Liu, L., 52, 69 Liu, T., 187, 202, 203, 204, 205, 206, 209 Liu, W., 99, 100, 101 Liu, Y., 145, 326, 359

Liu, Y.-N., 524, 530, 531, 532, 533, 535, 536 Liu, Z., 377, 388 Llewellyn, N. M., 489, 495, 504, 505, 506, 510, 511, 514 Lobo, S., 404, 421 Locht, C., 260, 262, 276 Logan, R., 247, 353 Lombo´, F., 34, 246, 356 Lomovskaya, N., 99 Long, J., 403 Long, P. F., 11, 191, 193, 197, 278 Lonsdale, J. T., 24, 25, 27, 263, 276 Lopanik, N. B., 127, 170 Lopez, P., 410 Losey, H. C., 522, 533 Loughran, M. S., 23, 222 Low, L., 22 Lowary, T. L., 263 Lowden, P. A., 124 Lowenstein, H., 34 Lozano, M. J., 387 Lu, H., 21, 37, 39, 206, 214 Lu, P., 398, 423 Lu, W., 389 Lu, X., 262, 268 Lu, Y., 30, 407 Lu, Y. W., 414 Lucher, L. A., 472, 473, 501 Luft, T., 446, 452 Luiten, R., 223 Lum, A. M., 251, 253, 351, 353 Lundgren, B., 254 Luniak, N., 360 Luo, G., 116, 196, 341 Luo, R., 21, 27, 29, 30, 384, 385 Lupoli, T., 389, 451 Luquin, M., 272, 277 Lurz, R., 485 Luzhetskyy, A., 6, 350, 368, 389, 524 Lydiate, D. J., 245, 249, 373, 472 Lynch, S., 249, 250 Lynen, F., 50 Lynett, J., 272 M Ma, S. M., 56, 60, 63, 384 Ma, Y., 9 MacArthur, M. W., 19 MacNeil, D. J., 442 MacNeil, I. A., 345, 350, 353 MacNeil, T., 442 Madden, T. L., 192 Maddry, J. A., 263 Madduri, K., 386 Madhusudhan, M. S., 209 Maeda, H., 98 Maes, E., 260

558 Magnuson, K., 403, 404 Magnusson, M., 274 Maharjan, J., 438 Mahmud, T., 203, 466, 478, 479, 480, 495 Maier, T., 19, 24, 25, 27, 37, 39, 268, 306 Majerus, P. W., 421 Malaga, W., 262, 268 Malpartida, F., 216, 217, 225, 236, 245, 254, 368 Maltby, D. A., 21, 24, 25, 34, 72, 368, 376 Manabe, C., 188 Manca, C., 268, 271, 278 Maniatis, T., 499 Mansilla, M. C., 410 Mansoor, H., 278 Mansouri, K., 472, 473, 474, 490, 502 Mantsala, P., 21, 204, 377, 386, 387, 388 Mao, Y., 124 Marahiel, M. A., 37, 52, 151, 175, 179, 182, 340, 360, 373, 399, 414 Marathe, U. B., 276, 280, 281 Marchis-Mouren, G., 484 Marcotte, E. M., 398, 423 Marcus, M., 356, 357, 366 Margeat, E., 262, 276 Markovic-Housley, Z., 34 Marquez, S., 356, 357, 366 Marrakchi, H., 262, 274, 276, 277, 410, 411, 419, 421, 423 Marsden, A. F. A., 23, 24, 116, 123, 222 Marsh, E. N., 387 Marshall, B. J., 280, 281 Marshall, J. W., 63, 65, 66 Marti, T., 36 Martin, C., 270, 272 Martin, C. J., 6, 273, 346 Martin, F., 30, 407 Martıń, J. F., 215, 216, 217, 218, 220, 223, 224, 225, 226, 227, 228, 229, 230, 233, 234, 235, 236, 237 Martinez, A., 345, 350, 353 Martinez, R. A., 145 Marti-Renom, M., 209 Massengo-Tiasse, R. P., 411, 420, 421, 423 Matern, U., 469 Matharu, A. L., 278 Mathema, B., 268 Mathews, I., 383 Mathews, I. I., 21, 22, 24, 25, 27, 39, 60, 64, 69 Mathur, M., 266, 270 Matsunaga, I., 272 Matsunaga, S., 5, 171, 172 Matsushima, Y., 501 Matsuura, S., 145 Matter, A. M., 170 Mattes, R., 358 Maurin, D., 270 Maurus, R., 464 Mauvais, P., 438

Author Index

Maxwell, A., 438 May, B. J., 278 Mayama, M., 145 Mayasundari, A., 412 Mayer, A., 66, 350 Mayer, G., 487 Mazodier, P., 246 McAda, P. A., 50, 62 McAdam, R. A., 262 McAllister, K. A., 399, 400 McAlpine, J. B., 11, 191 McArthur, H. A., 11, 191, 193, 197, 404, 415 McConville, M. J., 265, 276 McDaniel, R., 23, 24, 30, 34, 36, 296, 300, 301, 305, 307, 310, 322, 327, 329, 340, 344, 345, 348, 349, 350, 351, 353, 356, 364, 368, 373, 375 McDonald, F. E., 191 McGuire, K. A., 403, 422 McKeown, D. S. J., 53 McKinney, J. D., 265 McLafferty, F. W., 501 McMahon, H., 245 McMeekin, T., 9 McMurry, L. M., 407, 411 McNeil, M. R., 261, 268, 277 McNicholas, C., 53 McPherson, A., 18 Mdluli, K., 274 Meadows, E. S., 21, 24, 26, 372, 383 Medina, N., 278 Medzihradszky, K. F., 21, 24, 25, 34, 72, 368, 376 Megessier, S., 170 Mehta, R., 464 Meier, J. L., 380, 414 Meiser, P., 171 Melancon, C. E. III, 522, 523, 524, 530, 531, 532, 533, 534, 536, 540, 541, 550 Meluzzi, D., 381, 388, 389 Menche, D., 132, 133, 167, 170, 183 Mendes, M. V., 216, 217, 218, 223, 224, 225, 227, 237 Meńdez, C., 11, 34, 246, 387, 524 Menzella, H. G., 116, 298, 319, 321, 323, 324, 325, 327, 329, 330, 331, 332, 335, 339, 340, 348, 350, 358, 359 Mercer, A. C., 400 Mersinias, V., 246, 247, 347, 348 Merski, M., 36, 54 Merson-Davies, L. A., 356, 357 Metcalfe, S., 251 Metsa-Ketela, M., 377, 388 Metz, J. G., 9, 81, 83, 84, 85, 95, 99, 405, 413, 421 Meyer, F., 412 Meyer, W. E., 217 Miao, V., 350 Michel, J.-M., 375, 447, 477, 486

559

Author Index

Middlebrook, G., 270 Miercke, L. J., 21, 22, 24, 26, 37, 39 Migita, A., 200, 359 Miller, W., 192 Millership, J. J., 172 Mills, J., 403 Mills, P. R., 72 Minagawa, K., 203, 480 Minnikin, D. E., 260, 262, 274, 276, 277, 279, 282 Minnikin, S. M., 274 Mironenko, T., 11, 191, 192, 193, 197, 198, 199, 201, 210, 346, 502 Misaki, A., 261 Mislin, R., 29 Mittag, M., 99 Miyara, I., 68 Miyoshisaitoh, M., 99 Mizrahi, V., 266 Mizugaki, M., 406 Mizuguchi, K., 199 Mochizuki, S., 171 Modak, R., 434 Mofid, M. R., 399 Mogensen, J. E., 34 Mohan, S., 406 Mohanty, D., 11, 23, 259, 261, 262, 263, 265, 268, 269, 270, 271, 272, 273, 276, 279, 280, 281, 282, 283, 411, 412, 413 Molle, V., 261, 276 Mller Jensen, I., 254 Molnar, I., 23, 24, 122, 125, 126, 145, 218, 222, 234, 237 Monaghan, R. L., 145 Money, S., 145 Monsempe, C., 278 Montrozier, H. L., 262, 276, 277 Moody, D. B., 272 Moore, B. S., 32, 34, 36, 124, 282, 345, 375, 380, 381, 382, 384, 388, 389, 414 Mootz, H. D., 414 Morehouse, C. B., 263 Moreira, A. L., 268 Morgan-Warren, R. J., 464 Mori, K., 227 Mori, Y., 58 Moriguchi, T., 53 Morimoto, M., 167 Morita, N., 81, 83, 95 Morosoli, R., 485 Mosi, R., 482, 484 Moss, D. S., 19 Moss, S. J., 6, 11, 144, 145, 298 Moszer, I., 170 Motamedi, H., 32 Mougous, J. D., 262, 263, 271, 272 Moura, R. S., 227, 237 Mowbray, S. L., 199

Muchel, S. E., 263, 271 Muhlecker, W., 272 Mukerji, P., 83 Mu¨ller, J., 145 Mu¨ller, R., 6, 12, 129, 145, 167, 170, 171, 183, 271, 296, 306, 341, 349, 360, 523 Munoz, M., 277 Murakami, K., 523, 534, 535, 538, 539 Murata, M., 5, 194 Murli, S., 23, 24, 321, 325, 358 Murphy, B., 223, 235, 250, 254 Murphy, C. K., 411, 420, 421, 422 Murray, M., 21, 34, 36, 37 Murry, J. P., 264 Murugan, E., 101 Musicki, B., 438 Muskiet, F. A., 80 Muth, G., 450 Mutka, S. C., 69, 310, 326, 359 Muttucumaru, D. G., 264 Muyrers, J. P., 12, 13 Mve-Obiang, A., 10 N Naganawa, H., 188 Naharro, G., 228, 229, 230 Nakaido, H., 260, 261, 273 Nakamura, A., 482, 484 Nakamura, M., 188 Nakanishi, M., 400 Nakano, H., 167 Nakano, M. M., 147, 152 Nakashima, T. T., 195 Nakayama, J., 145 Nakayama, K., 468, 484 Nampoothiri, K. M., 262, 276 Nango, E., 500, 501, 502 Naoki, H., 5 Napier, J. A., 9, 80, 93 Narasimhan, M., 272, 409 Narumi, K., 266 Natarajan, V. T. D., 267, 272 Nedal, A., 236, 258 Nelsen, J. S., 405, 421 Ness, T., 188 Neumann, B., 102 Newburn, S. A., 264 Newton, G. L., 460, 464 Neyrolles, O., 270, 278 Nguyen, D. K., 50, 62 Nguyen, N. T., 464 Nguyen, T., 11 Nguyen, V. D., 438 Ni, X., 506 Nichols, B. P., 233 Nichols, C. M., 9 Nichols, D., 9

560

Author Index

Nichols, P. D., 9 Nicholson, G., 171 Nicholson, T. P., 37, 50, 51, 58, 61, 65, 382, 414 Nic Lochlainn, L., 252 Nie, L. P., 99 Nielsen, J., 145, 441 Niemi, J., 21, 204, 377, 386, 387, 388 Nierhaus, K. H., 464 Nietlispach, D., 11 Nigou, J., 260, 262 Nihira, T., 172, 227 Nijkamp, H. J., 403 Nikaido, H., 260 Nikodinovic, J., 249 Nilges, M., 18, 19 Nishida, T., 81, 83, 95, 511 Noel, J. P., 8, 24, 279, 345 Noguchi, H., 279 Noll, H., 261, 272 Nonaka, K., 99 Numakura, M., 503 Nur-e-alam, M., 388 Nussbaumer, B., 450 O Oberthur, M., 389, 450 O’Brien, K., 247, 353 O’Connell, J. D. III, 21, 22, 24, 26 Odakura, Y., 468, 484 Oefner, C., 403 Oesch, F., 199 Oethinger, M., 407, 411 Ogasawara, N., 170 Oguri, H., 200, 359 Oguro, S., 279 Oh, J., 530 Oh, T. J., 99, 215, 262, 263, 481, 503 Oh, W., 403, 404 O’Hagan, D., 28, 46, 267, 344, 368 O’Hare, H. M., 30, 117, 263, 276 Ohlrogge, J. B., 424, 425 Ohmori, D., 503, 504 Ohnishi, Y., 59 Ohno, T., 188 Ohnuki, T., 473 Ohta, T., 476 Oikawa, H., 200, 359 Oikawa, T., 262, 276 Okada, H., 227 Oki, T., 99 Okuyama, H., 81, 83, 95 Olano, C., 11, 134, 246 Oliveira, M. A., 388 Olivera, N., 218, 223, 224, 225 Oliynyk, M., 11, 99, 122, 191, 192, 193, 197, 198, 199, 201, 218, 234, 249, 296, 322 Oliynyk, Z., 193

Olland, A., 18 Olsen, J. G., 19, 24, 25, 27, 403, 405, 422 O’Maille, P. E., 8 Omura, S., 119, 188, 193, 216, 225, 381 Ono, M., 145 Ono, Y., 52 Onwueme, K. C., 265, 268, 270 Oppermann, U., 28 Orelle, C., 224 Orikasa, Y., 81, 83, 95 Orr, E., 171 Ortmann, R., 469 Osburne, M. S., 345, 350, 353 Osmark, P., 34 Ostergaard, L. H., 29 Ota, Y., 502 Otoguro, K., 188 Otsuka, M., 171 Otzen, D. E., 34 Ou, S., 348, 349, 353, 356, 364 Overall, C. M., 464 Owen, P., 380, 403 Ozanick, S. G., 490 Ozawa, H., 262, 279 Ozawa, M., 345 P Pacey, M. S., 123 Paget, M. S., 198 Paitan, Y., 171 Palaniappan, N., 32, 128 Palzer, M., 34 Pan, H., 21, 24, 26 Pannu, N. S., 18, 19 Papa, F., 273 Pape, H., 482, 486 Parajuli, N., 503, 505, 515 Parekh, S., 351 Parish, T., 262, 263, 276 Park, C., 62 Park, H. R., 227 Park, J. W., 503, 505, 515 Park, S. H., 500, 516 Park, S.-H., 531 Park, S. K., 416 Park, S. R., 503, 505, 515 Parkhill, J., 267, 268, 272, 276, 279, 282 Parkinson, J. A., 21, 34, 36, 37 Parlett, J. H., 274 Partida-Martinez, L. P., 172 Pashley, C. A., 262 Pasternak, O., 34 Patel, A., 170 Patel, H. M., 161 Patel, K., 301, 304 Patel, K. G., 321, 323, 327, 330, 348, 350, 358 Pathak, A. K., 263

561

Author Index

Pathak, V., 264 Patrick, J. B., 217 Paul, T. R., 260 Paulsen, H., 469 Pawlak, J., 229 Payan, F., 484 Payne, D. J., 411, 423 Payne, G. A., 53 Payre, G. A., 260 Pease, L. R., 55, 302 Peery, R. B., 399, 400 Peiru´, S., 310, 319, 321, 332, 335, 346, 355, 359 Pel, H. J., 50 Pelzer, S., 167, 170, 183, 341 Penn, J., 350 Pereira, S. L., 81, 83 Perez, E., 262, 268, 270, 282 Perez-Zuniga, F. J., 225 Perham, R. N., 222 Perlova, O. K., 167, 170, 183, 360 Pernodet, J.-L., 197, 475 Persson, B., 28 Petit, J. F., 260 Petkovic, H., 22, 122, 301 Petrakovsky, O. V., 353, 356, 357, 366 Petter, R., 246 Petzold, C. J., 262, 263, 271, 272 Pfeifer, B. A., 12, 345, 356, 358, 359, 372 Pflugrath, J. W., 18 Phan, T. H., 500, 516 Phatale, P. A., 262 Phillips, G. J., 416 Piagentini, M., 23, 24, 30 Piel, J., 11, 32, 125, 126, 127, 129, 170, 171, 172, 372, 388, 389, 453 Piendl, W., 486 Pieper, U., 209 Piepersberg, W., 246, 375, 447, 459, 460, 464, 465, 466, 467, 468, 472, 473, 474, 476, 477, 478, 479, 480, 481, 482, 485, 486, 487, 490, 495, 502, 530 Pieretti, I., 170 Pinkerton, M., 188 Piraee, M., 10 Pissowotzki, K., 473, 490 Pitarque, S., 260 Platzer, M., 11, 171, 172 Podesta, F., 346, 355 Podevels, A. M., 144, 145, 156, 160, 167 Pohl, N. L., 36 Polacco, M. L., 405, 408, 409 Polderman-Tijmes, J. J., 209 Polgar, N., 272 Polizzi, B. J., 411, 423 Popovic, B., 30, 502 Porcelli, S. A., 272, 276 Portevin, D., 276 Pospiech, A., 102

Post-Beittenmiller, D., 424, 425 Poulsen, F. M., 34 Power, P., 223, 235, 249, 254 Powles, H., 65, 67 Pradella, S., 170 Praseuth, A. P., 359 Pratt, M. R., 263, 271 Prescott, D. J., 398 Prestegard, J. H., 398, 425 Price, A. C., 19, 24, 25, 27, 29, 405 Priestley, N. D., 386 Prome, J. C., 277 Pross, E. K., 167, 170, 183 Prusky, D., 68 Pryor, M. J., 10, 278 Puehler, A., 450 Pugh, E. L., 419 Puglisi, J. D., 21, 36, 37 Pullen, J. K., 55 Pulsawat, N., 127, 129, 172 Puzo, G., 260 Pyke, J. S., 262, 276 Q Qiu, X., 24, 25, 27 Quadri, L. E., 147, 152, 161, 262, 267, 268, 270, 280, 281, 414 Quail, M. A., 193, 345, 346 Que, N. L., 531, 532, 535, 536 Quemard, A., 262, 263, 276 Quinlivan, E. P., 233 Quinn, C., 411, 423 Quiocho, F. A., 19, 37 Quiros, L. M., 387 R Rachid, S., 171 Radauer, C., 34 Radax, R., 13 Raetz, C. R., 397, 406 Rafanan, E. R., Jr., 388 Rafferty, J. B., 398, 419, 425 Rafidinarivo, E., 277 Rai, D., 223, 235, 254 Rainwater, D. L., 263, 266, 267 Rajkarnikar, A., 480 Rajski, S. R., 79 Ramakrishnan, V., 464 Rangan, V. S., 19, 22, 56, 64 Ranganathan, A., 327, 329 Rao, R. N., 247, 353 Rapp, H., 449 Rarick, D., 54 Rascher, A., 124 Ratledge, C., 280, 281 Ratnatilleke, A., 346 Rauzier, J., 270

562 Ravanel, S., 233 Rawlings, B., 223, 235, 236, 252, 254 Rawlings, B. J., 265 Rawlings, M., 404, 406, 416 Ray, T. K., 418 Rayment, I., 21, 386 Raynal, M.-C., 375, 447, 477, 486 Read, R. J., 18, 19 Re´beille´, F., 233 Rebets, Y., 6, 368, 389 Recio, E., 216, 217, 223, 225, 226, 227, 237 Redfield, C., 18, 23, 24, 32 Reed, M., 414 Reed, M. A., 37, 268 Reed, M. B., 268, 272, 279 Reed, S. L., 411, 423 Rees, D. O., 65 Reeves, A. R., 347, 356, 357, 366 Reeves, C. D., 23, 24, 69, 144, 145, 295, 301, 304, 311, 321, 340, 353, 356 Reeves, P., 469 Reeves, P. C., 50, 62 Reid, R., 30, 52, 69, 117, 151, 170, 175, 179, 182, 307, 321, 323, 324, 325, 327, 330, 331, 348, 350, 353, 356, 358, 373, 399 Reinscheid, U. M., 453 Reisinger, S. J., 321, 323, 324, 325, 327, 330, 331 Remakrishnan, V., 464 Remsing, L. L., 6, 387, 388 Revill, W. P., 144, 145, 301, 353, 356, 357, 366, 413, 414 Reynolds, K. A., 32, 222, 327, 355, 403, 415, 421, 422 Reynolds, R. C., 263 Rheinheimer, J., 145 Rice, D. W., 398, 419, 425 Rice, L. M., 18, 19 Richardson, C. C., 490 Richter, L., 9, 84, 95, 99 Rickards, R. W., 217 Ridell, M., 277 Riezman, H., 217 Rigden, D. J., 400 Riley, L. W., 262, 264, 271 Ripka, S., 50, 51 Rittmann, D., 264 Ritzenthaler, J. D., 264 Rivera-Marrero, 264 Rix, U., 8, 387, 388 Roach, C., 50, 62 Roberts, G. A., 10, 191, 196, 320, 341 Roberts, M., 368 Roberts, W. C., 3 Robinson, J. A., 188, 195, 198, 346 Rock, C. O., 19, 24, 25, 27, 30, 32, 37, 276, 398, 400, 403, 404, 405, 406, 407, 408, 409, 411, 412, 416, 417, 418, 419, 420, 421, 422, 423, 424

Author Index

Rock, Y. M., 19, 25, 405, 406, 407 Rockser, Y., 479, 480, 481, 483, 486 Rodriguez, E., 30, 295, 311, 321, 332, 335, 339, 340, 346, 348, 349, 353, 355, 356, 358, 359, 364 Rodriguez, G. M., 280 Rodrı´guez-Garcıá, A., 227, 237 Roepenack-Lahaye, E., 34 Roessler, P., 9, 81, 83, 99, 413 Rogers, L. M., 267, 268 Rohr, J., 8, 387, 388 Roig-Zamboni, V., 270 Rokem, J. S., 441 Rolon, M. S., 225 Roman, J., 265 Romer, U., 472, 530 Ron, E. Z., 171 Ronson, C. W., 412 Rosat, J. P., 272 Rosenberg, A. H., 490 Rosenberg, E., 171 Rosenfeld, I. S., 404, 405 Rosenzweig, B., 83 Rossi, A., 209 Roujeinikova, A., 398, 419, 425 Rousseau, C., 278, 279, 282 Rowe, C. J., 134, 269, 345 Roy, R. P., 11, 262, 271, 272, 273, 281 Royer, M., 170 Ruan, X., 122, 296, 301 Rubin, E. J., 262 Rubin, J. R., 407, 411 Rubin-Pitel, S. B., 280 Ruby, C. L., 442 Ruch, F. E., 22 Rudd, B. A., 22, 50, 65, 272 Rude, M. A., 145 Rudner, D. Z., 170 Rukmini, R., 279 Rumbero, A., 226, 227 Rupp, B., 18, 24 Rupprath, C., 472, 524, 530 Rupprath, G., 527, 531 Russell, D. W., 441 Russell, S., 195, 198 Rutter, K., 277 Ruzsicka, B. P., 415 Rydberg, E. H., 464 Ryu, J. S., 262, 266 Ryu, Y. G., 356 S Sacchettini, J. C., 276 Sacco, E., 261, 276 Sahm, H., 262, 263, 276 Sakaki, Y., 193, 225 Sakaue, M., 188

Author Index

Sakazaki, R., 145 Sakuda, S., 145, 227 Salah-Bey, K., 375, 447, 477, 486 Salas, J. A., 11, 34, 246, 387, 524 Samborskyy, M., 192, 193, 197, 198, 199, 201 Sambrook, J., 441, 499 Samel, S. A., 37 Sampson, A. E., 274 Samrat, S. K., 262 Sanchez, C., 55, 101, 107, 175, 179, 182, 387 Sanchez, E. L., 92 Sanchez, J. F., 267 Sanderbrand, S., 358 Sanderson, K., 9 Sankaranarayanan, R., 262, 268, 276, 279, 282 Sankawa, U., 52, 387 Sano, Y., 280 San Roman, A. K., 414 Santi, D. V., 30, 52, 69, 160, 321, 323, 324, 325, 327, 329, 330, 331, 344, 348, 350, 358 Santos-Aberturas, J., 218, 225, 237 Santos-Beneit, F., 227 Saran, S., 262 Sarma, S. P., 434 Sasaki, S., 501 Sassetti, C. M., 262 Satake, M., 5 Sath, S., 468 Sato, H., 274 Sato, K., 227 Sauer, F., 419 Saunton, J., 11 Saurel, O., 262, 264 Savage, D. F., 21, 22 Savagnac, A. M., 277 Savin, A., 170 Savvi, S., 266 Saxena, P., 11, 262, 263, 265, 276, 279, 281, 282, 411, 412, 413 Scapin, G., 488 Schaefer, W. B., 270 Schaeffer, M. L., 263, 276 Schafer, A., 360, 414 Schaffer, A. A., 192 Scha¨gger, H., 475 Scharfe, M., 272 Schatz, A., 460 Scheerer, J. R., 9, 36, 57 Scheffel, F., 485 Scheible, H., 446 Scheiner, O., 34 Schelle, M. W., 262, 272 Schenk, P. M., 358 Scherlach, K., 172 Schiltz, E., 50, 51 Schimana, J., 145 Schirmer, A., 353, 356 Schlo¨rke, O., 479, 480

563 Schlo¨sser, A., 485 Schluenzen, F., 464 Schmid, D. G., 145 Schmid, M., 30, 407 Schmidt, E. W., 54, 65 Schmidt, J., 446 Schmidt-Beibner, H., 460, 464, 466, 468, 473, 476, 495 Schmitz, R. A., 341 Schnarr, N. A., 10, 21, 37, 272 Schneider, E., 485 Schneider, G., 21, 204, 386, 387 Schneider, K., 170, 171 Schobert, M., 360, 414 Scholz, R., 170, 171 Schoner, B. E., 247, 353 Schou, C., 34 Schrempf, H., 245, 249, 373, 472, 485 Schro¨der, W., 482, 486 Schroeder, B. G., 274, 281 Schroeder, F. C., 170 Schuhmann, T., 145 Schulz, H., 403 Schumacher, T., 524, 530 Schu¨mann, J., 50 Schupp, T., 145 Schwab, J. M., 19, 30, 34, 272, 408, 409, 421 Schwarzer, D., 37 Schwecke, T., 23, 24, 124, 218, 222, 234, 237 Schweizer, E., 50, 51, 52, 99, 344, 411, 412 Schweizer, H. P., 410, 420, 423 Schweizer, M., 37 Scott, A. I., 50 Scott, N., 192, 193, 197, 198, 199, 201 Scrutton, N. S., 222 Sebek, O. K., 442 Seco, E. M., 225, 254 Seidel, M., 264 Seki, M., 233 Sekurova, O. N., 224, 236, 246, 247, 248, 255, 258 Sello, J., 280, 281 Senaratne, R. H., 262, 263, 271 Seno, E. T., 247, 353 Seo, M. J., 480 Seo, W. M., 481 Serre, L., 19, 22, 55, 56, 64, 403 Seto, N., 261 Shaflee, A., 145 Shafran, H., 68 Shah, A. N., 36, 379 Shah, S., 345 Shanklin, J., 418, 419 Shanmugam, V. M., 279 Shareck, F., 485 Sharma, J., 263 Sharma, K. K., 222 Sharma, S., 434

564 Sharma, S. K., 434 Shea, J. E., 262 Shelat, A. A., 21, 22 Sheldon, P. J., 381, 382 Shen, B., 3, 6, 9, 11, 12, 21, 34, 55, 79, 84, 92, 95, 97, 98, 99, 100, 101, 107, 156, 165, 166, 167, 168, 171, 172, 175, 179, 182, 183, 296, 368, 372, 380, 387, 388, 414 Shen, M. Y., 209 Shen, Y., 36 Shen, Z., 419, 425 Shepard, E., 99 Shepherd, J., 3 Sherman, A., 68 Sherman, D. H., 21, 27, 31, 37, 39, 136, 170, 222, 272, 321, 327, 341, 345, 381, 413, 540, 541 Sherman, M. M., 195 Sherringham, J. A., 195 Shi, J., 199 Shiba, T., 193, 225 Shibuya, M., 59 Shichijo, Y., 200 Shinose, M., 193, 225 Shinozaki, K., 233 Shiomi, K., 188 Shioyama, S., 264, 276 Shirahata, K., 468 Shirouzu, M., 464 Shoji, J., 145 Shoji, S., 464 Shreve, A. L., 345 Sidebottom, P. J., 272 Sidhu, G., 464 Siegner, A., 50, 51 Sievers, E. L., 98 Siggaard-Andersen, M., 19, 24, 25, 27, 403, 404, 422 Sikorski, M. M., 34 Silakowski, B., 126, 130, 131, 272 Silbert, D. F., 406, 407, 422 Silva, C. J., 350 Simeone, R., 272 Simon, S., 274 Simon, W. J., 398, 419, 425 Simpson, K. E., 277 Simpson, T. J., 9, 21, 22, 27, 28, 34, 36, 37, 45, 47, 48, 50, 51, 53, 54, 56, 60, 61, 63, 65, 66, 67, 72, 171, 177, 274, 380, 379, 380, 382, 403, 414 Sims, J. W., 65 Simunovic, V., 125, 127, 129, 171 Singh, A., 263, 276 Singh, D., 480 Sinnwell, V., 469 Sirakova, T. D., 262, 263, 264, 268, 271, 272, 278, 279, 282 Siskos, A. P., 22, 30, 99 Skellam, E., 54, 66

Author Index

Slabas, A. R., 398, 419, 425 Slayden, R. A., 262, 274, 276 Sletta, H., 220, 224, 236, 244, 246, 247, 248, 254, 255, 256, 258 Small, P. L., 10 Smirnova, N., 415, 422 Smith, C. P., 245, 246, 247, 249, 347, 348, 373, 472 Smith, I., 278 Smith, I. P., 357 Smith, J. L., 19, 21, 27, 30, 31, 34, 37, 39, 272, 407, 409, 413 Smith, L. H., 9, 11, 56, 414 Smith, M. C., 253, 441 Smith, P., 21, 35 Smith, S., 18, 19, 21, 22, 23, 24, 25, 29, 30, 32, 39, 55, 56, 64, 118, 276, 296, 301, 306 Smith, W. C., 172 Smogowicz, A. A., 403 Snow, G. A., 280, 281 Snow, M. E., 405, 411 So, C. Y., 411, 423 Socci, N. D., 173 Sogaard, M., 490 Sohn, M. J., 411 Sohng, J. K., 481, 489, 501, 502, 503, 505, 515, 530 Sola-Landa, A., 227, 237 Soler, L., 245 Soll, C. E., 263, 268, 270 Solsbacher, J., 360, 414 Somers, W., 18 Snder Hansen, B., 254 Song, D., 172, 183 Song, K. I., 438 Song, L., 345 Song, Z., 63, 65, 66, 67, 72 Sood, G. R., 188 Sosio, M., 341 Soto, C. Y., 272 Soulas, F., 414 Souza, C. M., 217 Spangfort, M. D., 34 Spencer, J. B., 11, 30, 50, 63, 191, 197, 198, 199, 200, 201, 204, 205, 206, 209, 381, 382, 489, 495, 502, 504, 505, 506, 510, 511, 514 Spiteller, D., 30, 381, 382, 502, 506, 510 Sprecher, H., 270 Sridharan, V., 263, 268, 269, 270, 276, 282 Stadler, P., 469 Stadthagen, G., 267, 268, 269 Staffa, A., 99, 170, 171 Standage, S., 99 Stapon, A., 448 Stark, C. B., 193, 196, 197, 198, 199, 200 Stark, H., 18, 19, 23, 24, 32, 39 Stasiuk, M., 279 Stassi, D. L., 296, 301, 355

565

Author Index

Staunton, J., 10, 11, 22, 23, 24, 48, 166, 191, 193, 196, 197, 198, 199, 200, 201, 204, 205, 206, 209, 218, 222, 234, 237, 251, 269, 274, 320, 322, 327, 329, 340, 341, 345, 346 Staver, M. J., 10, 191 Steger, D., 13 Stein, A., 530, 531 Steinbiss, H. H., 358 Steinrauf, L. K., 188 Stejskal, F., 172 Stephens, E., 171 Stephenson, G. R., 385 Stewart, A. F., 349, 360 Stewart, F. A., 13 Stinear, T. P., 10 Stirrett, K. L., 263, 268 Stocker, N. G., 260 Stockmann, M., 473 Stokes, R. W., 272 Straight, P. D., 170 Strain, M., 274 Stratigopoulos, G., 237 Stratmann, A., 466, 479, 480, 482, 486 Strecker, G., 260 Streit, W. R., 341 Strittmatter, A. W., 170, 171 Strobel, R. J., 351 Strohl, W. R., 386 Strom, A. R., 246, 247, 254, 255 Stroshane, R., 188 Stroud, R. M., 21, 22, 24, 25, 26, 27, 28, 30, 34, 37, 39, 72, 214, 306, 368, 376 Struyk, A. P., 217 Stubbs, M. T., 37 Studier, W. F., 490 Stuitje, A. R., 19, 22, 55, 398, 403, 419, 425 Su, H., 268 Su, N., 411, 420, 421, 422 Subba, B., 489, 501, 502 Subbarayan, C. R., 50 Sudek, S., 126, 127, 129, 170 Sugamura, K., 227 Sugihara, S., 83 Sugino, F., 171 Suh, J. W., 480, 500, 516 Suling, W. J., 264 Sullivan, J. T., 412 Sullivan, S. A., 420, 423 Sultana, A., 21, 386 Sulzenbacher, G., 270 Summers, R. G., 368, 380, 414 Sun, Y., 11, 191, 192, 193, 201, 202, 203, 205, 206, 209, 210, 230 Sunaga, R., 99 Surolia, A., 11, 260, 267, 268, 269, 279, 434 Surolia, N., 434 Susani, M., 34 Susskind, M. M., 416

Su¨ssmuth, R. D., 12, 170, 171 Sutherland, A., 62, 63 Suwa, M., 171 Suzukake, K., 503 Suzuki, A., 145 Suzuki, H., 356, 357, 366 Svendsen, I., 405 Svenson, K., 18 Svensson, B., 490 Swanson, S., 355 Swanson, S. J., 10, 191 Swindell, A. C., 406 Szaflarski, W., 464 Szafranska, A. E., 22, 37, 177, 380, 400, 414 Szu, P.-H., 538 T Ta, P., 460, 464 Tabor, S., 490 Tachibana, K., 194 Tada, H., 52 Tagami, U., 495, 500 Taguchi, T., 385 Takagi, M., 501 Takahashi, H., 524, 530, 531, 532, 533, 535, 536 Takahashi, K., 167, 468 Takahashi, Y., 188 Takaku, H., 501 Takano, E., 346, 375, 472, 479 Takasawa, S., 468 Takayama, K., 261, 274, 276, 277 Takazawa, Y., 484 Takemoto, C., 464 Takeuchi, T., 188 Takiguchi, T., 167 Tamegai, H., 477, 478, 495, 500, 501, 502 Tammer, M. E., 66 Tan, D. S., 263, 268 Tan, Y. H., 21, 27, 29, 30, 384, 385 Tanaka, H., 216 Tanaka, N., 501 Tang, G. L., 11, 125, 126, 127, 131, 166, 167, 168, 171, 182, 183 Tang, L., 32, 124, 126, 135, 296, 298, 344, 350, 351 Tang, X., 502 Tang, Y., 10, 21, 22, 24, 25, 27, 32, 34, 35, 39, 60, 63, 64, 69, 70, 272, 306, 367, 371, 374, 375, 376, 379, 381, 382, 383, 384, 385, 387, 388 Tanner, J. A., 61 Tatsuno, S., 132, 134 Tauch, A., 412 Taudien, S., 11 Taylor, B. L., 224 Taylor, J. W., 50 Taylor, M. W., 13

566 Taylor, N. L., 271 Teartasin, W., 21, 27, 28 Tekaia, F., 267, 268, 272, 278, 279, 282 ten Napel, H. H., 217 Terwilliger, T., 18 Textor, A., 167, 170, 183 Thamchaipenet, A., 322, 329, 344 Theisen, M., 129 Thibodeaux, C. J., 521, 522, 523, 524, 540 Thirumala, A. K., 271 Thoden, J. B., 541 Thomas, C. M., 171 Thomas, I., 134, 272 Thomas, I. P., 327, 329 Thomas, J., 395, 400 Thomas, M. G., 143, 144, 145, 156, 160, 167, 490, 533 Thomas, P. M., 9, 36, 57 Thomas, R., 36 Thompson, C., 246 Thompson, C. J., 375, 472, 474, 479 Thompson, J. D., 174, 177 Thompson, T. B., 21, 386 Thomson, N. R., 193, 345, 346 Thornton, J. M., 19 Thorpe, H. M., 441 Thorson, J. S., 3, 98, 99, 101, 381, 451, 522, 523, 524 Tickoo, R., 263, 268, 269, 270, 276, 282 Timmerman, P., 260 Timoney, M., 327, 329 Tobert, J. A., 62 Toelzer, S., 144, 145 Tokiwa, Y., 99 Tokiwano, T., 200 Tokunaga, K., 503 Toney, M. D., 55, 92, 101, 107, 175, 179, 182 Tonge, P. J., 415 Torrelles, J., 277 Toubiana, R., 274 Towne, T. B., 191 Townsend, C. A., 9, 36, 54, 56, 57, 60, 61, 67, 188, 191, 195, 197 Trail, F., 54, 72 Tran, C. Q., 324, 325, 335 Tremblay, S., 11 Trinh, K. Q., 144 Trivedi, O. A., 263, 268, 269, 270, 276, 282 Truman, A. W., 504, 505 Tsai, H. F., 60 Tsai, S. C., 17, 19, 21, 22, 24, 25, 26, 27, 28, 29, 30, 35, 37, 39, 118, 206, 214, 296, 301, 306, 374, 379, 381, 384, 385 Tsan, P., 254, 255, 256 Tsay, J. T., 401, 403 Tsenova, L., 268 Tsui, G., 58 Tsuji, S. Y., 39, 152, 323, 327, 329

Author Index

Tsujishita, Y., 34 Tu, G., 11, 191, 192, 193, 201, 202 Tull, D., 264, 276 Tunca, S., 224, 227, 237 Turgeon, B. G., 50 Turnowsky, F., 406, 407, 411 Tyman, J. H., 275 U Udwary, D. W., 36, 54, 282, 345 Uehara, Y., 503 Ui, H., 188 Uitdehaag, J. C. M., 482, 484 Ulrich, A. K., 405 Ulrichs, T., 272 Umezawa, H., 503 Unkefer, C. J., 145 Utsumi, Y., 280 Uttaro, A. D., 81 V Vagelos, P. R., 22, 398, 400, 403, 404, 405, 407, 421, 422, 423 Vagstad, A. L., 9, 36, 56, 57, 60, 61 Valentine, R., 9, 81, 83, 99, 413 Valenzano, C., 32, 202, 203, 205, 206, 209 Valero-Guillen, P., 273 Valla, S., 246, 247, 248, 254, 255 Van der Heever, J. P., 63 van der Linden, K. H., 403 van der Veen, B. A., 482, 484 van der Werf, M. J., 199 van Eek, T., 217 van Halbeek, H., 277 Van Lanen, S. G., 3, 9, 84, 95, 97, 98, 99, 100, 156, 296 van Wettstein-Knowles, P., 404 Vater, J., 145, 170, 171 Vats, A., 264, 268 Vedadi, M., 30, 406 Vederas, J. C., 62, 63, 195 Veitch, J. A., 98 Vente, A., 341 Verbree, E. C., 19, 22, 55, 403 Verseck, S., 472, 530 Verwoert, I. I., 403 Vetcher, L., 307, 308 Vial, H. J., 188 Viard, M., 170 Viell, P., 486 Vilcheze, C., 272 Villanueva, J. R., 228, 229, 230 Vinci, V. A., 351 Vining, L. C., 230, 387 Viswanathan, N., 30, 170 Vivien, E., 170 Vogel, C., 398, 423

567

Author Index

Volchegursky, Y., 344, 348, 349, 353, 356, 364 Volff, J. N., 486 Volk, K. J., 98 Volker, C., 276 Vollmar, D., 145 von Jagow, G., 475 von Mulert, U., 350 von Wettstein-Knowles, P., 19, 24, 25, 27, 403, 404, 422 Vos, C. J., 268, 270 Vosburg, D. A., 37 Vrijbloed, J. W., 346 Vroom, J. A., 251, 253 Vu, T. N., 21, 27, 28, 35 W Waddell, S. J., 262, 279, 282 Waggoner, L. E., 170 Wagner, B., 37 Wagner, G., 37 Wagner, M., 13 Wahl, H. P., 472 Waisvisz, J. M., 217 Wakil, S. J., 19, 37, 406, 419 Wakisaka, N., 501 Wakisaka, Y., 145 Waksman, S., 460 Walczak, R. J., 386 Waldrop, G. L., 399 Walker, J. B., 466, 472, 473, 501, 502 Walker, M. S., 501, 502 Walker, S., 389 Wallace, K. K., 403, 415 Wallis, J. G., 9, 80, 81, 93, 413 Wallis, N. G., 411, 423 Wallner, P., 407, 411 Walsh, C. T., 10, 37, 52, 147, 151, 152, 161, 170, 175, 179, 182, 279, 281, 296, 317, 340, 343, 359, 373, 389, 398, 399, 414, 425, 448, 450, 451, 522, 533 Walton, L. J., 145 Wang, B., 11, 191, 192, 201 Wang, C., 261, 274, 276, 277 Wang, C. C., 60, 263, 327, 330, 359, 371, 375, 385, 387 Wang, H., 406, 410 Wang, J., 18, 24 Wang, J. S., 21, 386 Wang, M., 11, 191, 192, 201 Wang, R., 398, 423 Wang, S. C., 503 Wang, X. W., 98 Wang, Y., 12, 386, 464 Wang, Z., 172, 183 Ward, J. M., 245, 249, 373, 472 Ward, S., 356 Ward, S. L., 119, 128, 313, 353, 356, 357, 366

Warner, D. D., 65 Warner, D. F., 266 Warude, D., 81 Washington, J. A., 3 Watanabe, A., 58, 59 Watanabe, C. M. H., 54, 67 Watanabe, K., 21, 60, 70, 81, 83, 95, 200, 327, 330, 359, 371, 375, 384, 385, 386, 387, 388 Watanabe, M., 200 Watanabe, T., 280 Wateman, C. L., 381 Watts, J. L., 9, 80, 81, 93, 413 Weaver, C. A., 85 Webb, B. M., 209 Webb, J. S., 217 Weber, J. M., 347, 356 Weber, T., 37, 167, 170, 183, 341 Webster, R. W., Jr., 160 Weeks, G., 406 Weeks, R. E., 188 Wehmeier, U. F., 375, 447, 459, 460, 464, 466, 468, 473, 476, 477, 478, 479, 480, 481, 482, 483, 484, 485, 486, 495 Weingarten, P., 375, 447, 472, 477, 486, 530 Weinreb, P. H., 147, 152, 280, 281, 414 Weinstein, D., 418 Weiser, J., 197, 475 Weisshaar, B., 412 Weissman, K. J., 3, 10, 11, 12, 21, 48, 99, 116, 117, 118, 166, 191, 192, 193, 201, 210, 296, 306, 314, 320, 340 Weist, S., 12 Welby, M., 277 Welch, M., 321, 322, 324, 325, 331, 348, 350, 358 Wellein, C., 99 Welscher, Y. M., 217 Welzel, K., 167, 170, 183, 341 Wemakor, E., 449 Wen, G., 171, 172 Wendt-Pienkowski, E., 99, 101, 172, 183 Wenzel, S. C., 6, 12, 145, 167, 170, 183, 272, 349, 360 Werbitzky, O., 474 Wessel, W. A., 368, 380, 414 Wessjohann, L., 448 Wessjohann, L. A., 446 West, J., 411, 423 Westcott, J., 51, 61, 278, 382, 414 Westley, J. W., 188, 195, 214 Westrich, L., 453 Weymouth-Wilson, A. C., 522 Wheatcroft, M. P., 11, 346 Wheeler, M. H., 57 White, J., 375, 472, 479 White, J. A., 530, 532, 534 White, R., 233

568

Author Index

White, S. W., 19, 24, 25, 27, 29, 403, 405, 406, 407, 410 White-Phillip, J. A., 521, 531, 532, 533, 550 Whitfield, C., 397 Whiting, A., 350 Whitwam, R. E., 99 Whitworth, K. P., 60, 61 Wiesmann, K. E., 196, 320, 341 Wiesner, J., 358 Wietzerbin, J., 260 Wilkinson, B., 6, 11, 22, 65, 272, 327, 329 Wilkinson, C., 193 Wilkinson, C. J., 130, 269, 346 Willett, N. J., 53 Williams, C., 21, 209 Williams, G. J., 451 Williams, J. S., 381 Williams, K., 72 Williams, M. G., 30 Williams, R. P., 217 Williams, S. J., 263 Williamson, R. M., 145 Willis, C. L., 65 Wilson, D. N., 464 Wilson, S. E., 253, 441 Wilson, W. R., 3 Wimberly, B. T., 464 Wimmer, R., 34 Winfield, C., 51, 61, 382 Winsor, C., 21, 36, 37 Winston, R. L., 57 Wirtz, G., 346 Wissenbach, M., 405 Withers, S. G., 464, 482, 484 Witkowska, H. E., 19, 21, 25, 32, 56, 64 Witkowski, A., 18, 19, 21, 23, 24, 25, 32, 39, 55, 64, 123, 276 Witter, D. J., 63 Wohlleben, W., 341, 450 Wolpert, M., 441, 442 Wong, J., 21, 27, 29, 30, 384, 385 Woo, E., 344 Woo, J. S., 501 Worthen, D. R., 254 Worthington, A. S., 380, 414 Wright, G., 522 Wrigley, S. K., 350 Wu, B., 37 Wu, H., 229, 234, 235 Wu, J., 32, 130, 132, 206 Wu, K., 124, 144, 145, 356 Wu, N., 330 Wu, X., 478, 479, 480 Wybenga, G., 209 X Xia, H., 506 Xia, Z. J., 308

Xiang, L., 34, 345, 381, 388, 389 Xiang, Y., 501 Xie, H., 98 Xie, X., 60, 70, 384 Xie, Z., 260 Xu, H., 203, 415, 450, 480 Xu, J., 144, 145 Xue, Q., 145, 296, 309, 311, 348, 350, 356 Xue, Y., 135 Xx, X., 18, 19 Y Yadav, G., 11, 23, 262, 263, 271, 272, 273, 277, 280, 281, 282 Yakasai, A. A., 63, 65, 66, 72 Yamada, A., 9, 81, 83, 99, 413 Yamada, H., 188 Yamada, K., 171 Yamada, Y., 227 Yamamoto, Y., 477, 478, 501, 502, 503, 504, 505, 509 Yamane, K., 482, 484 Yamase, H., 523, 534, 535, 538, 539 Yamashita, J. T., 273 Yamauchi, N., 500, 501 Yanagimoto, M., 227 Yang, C. F., 99 Yang, K. Q., 387 Yang, M., 170 Yang, X., 170, 171 Yang, Y., 383 Yano, I., 266 Yao, X., 398, 423 Yasumoto, T., 5 Yasuoka, Y., 60 Yatome, C., 400 Yazawa, K., 9, 81, 83, 99, 413 Yeung, S. M., 538 Yip, C. L., 345, 350, 353 Ylihonko, K., 386 Yocum, R. R., 412 Yoder, O. C., 50 Yokoyama, K., 477, 478, 501, 502, 503, 504, 505, 509 Yokoyama, S., 464 Yoo, J. C., 438 Yoon, Y. J., 32, 503, 505, 515 Yoshida, M., 167 You, D., 202, 203, 205, 206, 209 Young, D., 263 Young, D. C., 272 Yu, J., 53, 502 Yu, Q., 202, 203, 205, 206, 209 Yu, T. W., 36, 124, 144, 145, 236, 345, 346, 356, 413, 488 Yu, W.-L., 531, 532, 533, 540, 541, 550 Yu, Y., 203, 480, 506 Yu, Y. J., 410

569

Author Index

Yuan, Y., 274 Yukita, T., 511 Yuqing, T., 443, 444 Z Zaleski, T. J., 32 Zapp, J., 171 Zawada, B. I., 32, 34 Zawada, R. J., 372, 385 Zawodny, J., 145 Zazopoulos, E., 11, 99, 170, 171 Zeeb, M., 18, 23, 24, 32 Zeeck, A., 145, 453, 479, 480 Zeidner, D., 355 Zerbe-Burkhardt, K., 346 Zha, W., 276 Zhan, J., 60, 70, 384, 387, 388 Zhang, C., 451, 523 Zhang, E., 407, 411 Zhang, G., 193, 201, 202 Zhang, H., 531, 532, 533, 536, 550 Zhang, H. R., 12 Zhang, J., 99, 100, 192 Zhang, L., 170 Zhang, Q., 278 Zhang, W., 21, 34, 35, 60, 367, 371, 374, 375, 381, 384, 385, 387, 388 Zhang, Y., 13, 65, 203, 349, 360, 405, 412, 480 Zhang, Y. M., 19, 25, 29, 37, 404, 405, 406, 407, 410 Zhang, Z., 192

Zhao, B., 403, 415 Zhao, C., 156, 172, 183 Zhao, G., 399, 400 Zhao, H., 280 Zhao, L., 523, 534, 535, 538, 539, 540, 541 Zhao, Z., 523, 533, 534, 535, 536, 538, 539 Zheng, C. J., 411 Zheng, J., 19, 25, 37, 402, 404, 405 Zhou, H., 70 Zhou, X., 11, 172, 183, 191, 192, 193, 201, 202, 203, 204, 205, 206, 209, 229, 233, 234, 235, 237, 480 Zhou, Y., 229, 234, 235 Zhu, D., 172, 183 Zhu, G., 172 Zhu, J., 229, 234, 235 Zhu, K., 410, 412 Zhulin, I. B., 224 Zielinski, J., 229 Ziermann, R., 310, 345, 347, 348, 375 Ziminski, T., 229 Zirkle, R., 9, 84, 85, 95, 99, 145 Zlatopolskiy, B. D., 453 Zotchev, S. B., 216, 217, 220, 223, 224, 235, 236, 243, 244, 245, 246, 247, 248, 254, 255, 256, 258 Zou, J., 199 Zuber, B., 260 Zuber, P., 52, 147, 151, 152, 175, 179, 182, 373, 399 Zurita, J., 268, 270

Subject Index

A AA, see Arachidonic acid Acarbose biosynthetic gene cluster, 463 structure, 461 synthesis, 465, 478–486 transporters, 485–486 Acarviosyl transferase, cyclitol-containing aminoglycoside synthesis, 482–484 N-Acetylcysteamine thioesters substrate preparation, 205–206, 208 thioesterase assays, 206–208 2000 -N-Acetyl-6000 -deamino6000 hydroxyneomycin C deacetylase, neomycin synthesis, 509 20 -N-Acetylparomamine deacetylase, cyclitolcontaining aminoglycoside synthesis, 505 20 -N-Acetylparomamine synthase, cyclitolcontaining aminoglycoside synthesis, 504 ACP, see Acyl carrier protein Actinorhodin, structure, 369 Acyl carrier protein acyltransferase-less type I polyketide synthase acyltransferase-catalyzed loading assay of acyl CoA extender substrate to holoprotein, 179–182 holoprotein preparation, 175–177 mass spectrometry characterization, 177–178 overexpression of recombinant apoprotein, 174–175 extender unit linkage, see Polyketide synthase extender units fatty acid synthesis in bacteria functions, 397–400 gel electrophoresis of species, 423–425 gel filtration of species, 425 preparation from Escherichia coli acyl substrate synthesis, 418–420 apoprotein, 418 holoprotein, 416–418 overview, 415–416 strains and plasmids, 416 iterative type I polyketide synthase apoprotein overexpression and purification, 107–108 holoprotein preparation in vitro, 108–109 multiple domains, 130–131

phosphopantetheinyl transferase activation, 118 polyketide synthase/fatty acid synthase structure, 36–37 tandem acyl carrier protein domains in polyunsaturated fatty acid synthases acyl carrier protein overexpression, 89–91 holoprotein preparation, 92–93 number of active acyl carrier proteins and fatty acid production, 93–94 phosphopantetheinyl transferase overexpression, 91–92 plasmid construction, 88–89 recombinant Shewanella japonica enzyme expression in Escherichia coli, 85–86 site-directed mutagenesis and active site mapping, 86–88 type II polyketide synthase assays ketosynthase–acyl carrier protein interaction assay, 380–381 malonyl-CoA:acyl carrier protein transacylation assay, 380 expression and modification for studies, 372–373 Acyltransferase domain, polyketide synthase discrete acyltransferase loading assay of acyl CoA extender substrate to holo-acyl carrier protein, 179–182 overexpression of recombinant enzyme, 177 substrate specificity assay, 177, 179 ketide unit structure determination by domains, 298–299 multiple domains, 130 phylogenetic analysis of cognate and discrete acyltransferases, 167, 173 replacement engineering a-carbon substitution alterations, 300–301 chromosomal insertion, 301–305 specificity extension modules, 122 loading modules AMP-ligase loading domain, 124–125 GNAT domain, 125 KSQ domain, 123–124 leinamycin loading module, 125–126 overview, 123 structure, 22–24

571

572 Acyltransferase-less type I polyketide synthases acyl carrier proteins acyltransferase-catalyzed loading assay of acyl CoA extender substrate to holoprotein, 179–180 holoprotein preparation, 175–177 mass spectrometry characterization, 177–178 overexpression of recombinant apoprotein, 174–175 discovery, 166–167 discrete acyltransferase overexpression of recombinant enzyme, 177 substrate specificity assay, 177, 179 modules, 166 phylogenetic analysis of cognate and discrete acyltransferases, 167, 173 products, 167, 169–172 prospects for study, 182–183 trans-polyketide synthase activities, 127 Albicidin structure, 169 synthesis, 170 Ambruticin VS3, structure, 122 Aminocoumarin antibiotics, see also specific drugs biosynthetic gene clusters deletions in integrative cosmids, 442–444 heterologous expression, 441–442 chemoenzymatic synthesis clorobiocin analogs, 449–451 substrate generation, 447–449 clorbiocin biosynthesis, 440 metabolic engineering, 451–453 mutasynthesis amide synthetase assays for suitable ring A analog identification, 446–447 genes for expanding product range, 447 generation of new antibiotics, 444–446 structures, 438–439 Aminoglycosides, see Cyclitol-containing aminoglycosides p-Aminobenzoic acid, biosynthesis, 230, 233, 238 AMP-ligase loading domain, polyketide synthase, 124–125 Amphotericin B antifungal activity, 244 biosynthesis gene clusters, 244–246 overview, 10 structure, 4, 244 Apramycin structure, 461 synthesis, 465 Arachidonic acid, synthesis, 80 L-Arginine:scyllo-inosamine-phosphate amidinotransferase, cyclitol-containing aminoglycoside synthesis, 473

Subject Index

Aromatase/cyclases, polyketide synthases classes, 32–34 polyketide synthase structure, 34–36 type II polyketide synthase assay, 384–386 Avermectins, structures, 121 B Bacillaene structure, 121, 169 synthesis, 170 Bacterial fatty acids, see also Cell wall, Mycobacteria tuberculosis acyl carrier protein functions, 397–400 gel electrophoresis of species, 423–425 gel filtration of species, 425 preparation from Escherichia coli acyl substrate synthesis, 418–420 apoprotein, 418 holoprotein, 416–418 overview, 415–416 strains and plasmids, 416 biosynthesis assays, 420–422 branched-chain fatty acids, 410 enoyl-acyl carrier protein reductase, 407, 410–411 enzyme purification, 420 3-hydroxyacyl-acyl carrier protein dehydratase, 407 3-ketoacyl-acyl carrier protein reductase, 406 3-ketoacyl-acyl carrier protein synthase, 404–406 overview, 400–402 type I megasynthase fatty acid synthesis, 411–412 overview, 396–397 polyketide synthesis relationship, 412–415 polyunsaturated fatty acid synthesis anaerobic synthesis, 410 gene clusters, 83–84 3-hydroxydecanoyl-acyl carrier protein dehydratase, 407–409 industrial synthesis, 93, 95 overview, 80–81, 400 pathways, 81–83 tandem acyl carrier protein domains in polyunsaturated fatty acid synthases acyl carrier protein overexpression, 89–91 holoprotein preparation, 92–93 number of active acyl carrier proteins and fatty acid production, 93–94 phosphopantetheinyl transferase overexpression, 91–92 plasmid construction, 88–89

573

Subject Index

recombinant Shewanella japonica enzyme expression in Escherichia coli, 85–86 site-directed mutagenesis and active site mapping, 86–88 reconstitution of synthesis systems, 422–423 Bikaverin nonaketide synthase, see Fungal type I polyketide synthases Borrelidin, structure, 122 Broken modules, polyketide synthase, 131–133 Bryostatin structure, 121, 169 synthesis, 170 Butirosin biosynthetic gene cluster, 462, 495–497 structure, 461 synthesis 20 -N-acetylparomamine deacetylase, 505 20 -N-acetylparomamine synthase, 504 60 -dehydro-60 -oxoparomamine aminotransferase, 506–507 2-deoxy-scyllo-inosose synthase, 500–501 g-L-glutamyl-acyl carrier protein decarboxylase, 512–513 g-L-glutamyl-acyl carrier protein ligase, 511–512 g-L-glutamyl-4-aminobutyryl acyl carrier protein monooxygenase, 513 g-L-glutamyl-4-amino-2-hydroxybutyryl acyl carrier protein:ribostamycin g-Lglutamyl-4-amino-2hydroxybutyryltransferase, 514 g-L-glutamyl-butirosin B g-L-glutamyl cyclotransferase, 514–515 L-glutamine:deoxy-scyllo-inosose aminotransferase ‘radical S-adenosylmethionine dehydrogenase, 503–504 NAD-dependent dehydrogenase, 503 overview, 501–502 overview, 465, 495.498 paromamine 60 -dehydrogenase, 505–506 phosphoribostamycin phosphatase, 508 phosphoribostamycin synthase, 507–508 C C-1027 anticancer activity, 98 structure, 98 CAGs, see Cyclitol-containing aminoglycosides Calcicheamicin anticancer activity, 98 structure, 98 Candidicin applications, 228–229 biosynthesis p-aminobenzoic acid biosynthesis, 230, 233 gene cluster, 229–232

monooxygenase genes, 234–235 mycosamine biosynthesis, 236 phosphate repression, 237 polyketide synthases, 233–234 prospects for study, 237–238 regulatory genes, 236–237 transporter genes, 235 structure, 216–217, 228 Carboxymycobactin siderophore synthesis, 280–281 structure, 280 Cell wall, Mycobacteria tuberculosis composition, 260–259 functions, 260 high-performance liquid chromatography of polyketide synthase-derived saturated fatty acids, 282–284 iron-chelating siderophore synthesis, 280–281 lipid biosynthesis acetate/propionate feeding studies, 265–266 assay of PKS5 and PKS6, 281–282 dimycocerosate ester synthesis, 267–270 enzyme characterization techniques, 263–264 pathway evaluation techniques, 262 PKS2 and sulfolipid synthesis, 270–272 PKS3/4 and phthenoic acid synthesis, 277–278 PKS10-PKS11 cluster, 278–279 PKS12 and mannosyl-b–1phosphomycoketide synthesis, 272–273 PKS13 and mycolic acid synthesis, 273–276 PKS18 and long-chain pyrone synthesis, 279 polyketide synthases identification with genome sequencing, 266 types and domain organizations, 267 Chalcomycin, synthesis, 120, 127 Chemobiosynthesis polyketide synthase engineering, 311, 313 thymidine diphosphate deoxysugars, see Thymidine diphosphate deoxysugars Chivoavol A structure, 169 synthesis, 170 Cholesterol oxidase, pimaricin synthesis regulation, 225 Clorbiocin, see also Aminocoumarin antibiotics biosynthetic gene clusters deletions in integrative cosmids, 442–444 heterologous expression, 441–442 chemoenzymatic synthesis of analogs, 449–451 structure, 439 synthesis, 440 Coumermycin A1, see also Aminocoumarin antibiotics biosynthetic gene clusters deletions in integrative cosmids, 442–444

574

Subject Index

Coumermycin A1, see also Aminocoumarin antibiotics (cont.) heterologous expression, 441–442 structure, 439 Curacin A structure, 121 synthesis, 130 Cyclitol-containing aminoglycosides, see also specific drugs biosynthesis butirosin, see Butirosin acarbose, 478–486 acarviosyl transferase, 482–484 L-arginine:scyllo-inosamine-phosphate amidinotransferase, 473 classes of products, 467–469 dTDP-D-glucose 4,6-dehydratase, 471 dTDP-4–6-glucose 3,5-epimerase, 471 dTDP-D-glucose synthase, 470 dTDP-L-rhamnose synthase, 471–472 expression of recombinant enzymes, 499–500 gene clusters, 462–464 L-glutamine:scyllo-inositol transaminase, 472–473 glycosyltransferases, 476–477 inositol-2-dehydrogenase, 475–476 KanM1, KanM2, and KanN, 477–478 neomycin, see Neomycin pathways, 464–467 prospects for study, 515–516 starch-degrading enzymes, 484–485 streptomycin-phosphate phosphatase, 473–474 structures, 461 therapeutic targets, 460, 464, 494–495 Cyclohexanecarboxylate, synthesis, 124–125 D Daunorubicin, structure, 369 6000 -Deamino-6000 -dehydro-6000 -oxoneomycin, neomycin synthesis, 510 6000 -Deamino-6000 -hydroxyneomycin C dehydrogenase, neomycin synthesis, 509–510 DEBS, see Deoxyerythronolide B synthase Dehydratase domain, polyketide synthase engineering for b-carbon processing alteration, 305–307 ketide unit structure determination by domains, 298–299 structure, 30–32 trans-polyketide synthase activities, 128 60 -Dehydro-60 -oxoparomamine aminotransferase, cyclitol-containing aminoglycoside synthesis, 506–507 Deoxyerythronolide B synthase

docking domains, 117 engineering, 296 modules, 114, 116–118 structure acyl carrier protein, 36–37 acyltransferase domain, 22–24 dehydratase domain, 30–32 examples, 21 fatty acid synthase homology, 18 ketoreductase domains, 27–30 ketosynthase domain, 24–27 nuclear magnetic resonance overview, 18, 21 paradigm generalization, 118–119, 136 prospects for study, 39 thioesterase, 37–39 X-ray crystallography overview, 18–19, 21 2-Deoxy-scyllo-inosose synthase, cyclitol-containing aminoglycoside synthesis, 500–501 Deoxythymidine diphosphate, see d TDP enzymes; Thymidine diphosphate sugars DHA, see Docosahexaenoic acid Difficidin structure, 169 synthesis, 170 Dimycocerosate esters, synthesis, 267–270 Disorazole structure, 122, 169 synthesis, 132, 170 Docosahexaenoic acid, synthesis, 80–83 Docosapentaenoic acid, synthesis, 80, 82–83 Doxorubicin, structure, 4–5 DPA, see Docosapentaenoic acid dTDP-D-glucose 4,6-dehydratase, cyclitolcontaining aminoglycoside synthesis, 471 dTDP-4–6-glucose 3,5-epimerase, cyclitolcontaining aminoglycoside synthesis, 471 dTDP-D-glucose synthase, cyclitol-containing aminoglycoside synthesis, 470 dTDP-L-rhamnose synthase, cyclitol-containing aminoglycoside synthesis, 471–472 E Eicosapentaenoic acid, synthesis, 80, 82–83 Enediyne synthesis, see Iterative type I polyketide synthase Enoyl-acyl carrier protein reductase diversity in bacteria, 410–411 fatty acid production in bacteria, 406–407 Enoyl reductase domain, polyketide synthase engineering for b-carbon processing alteration, 305–307 ketide unit structure determination by domains, 298–299 EPA, see Eicosapentaenoic acid Epo polyketide synthase, intermodular interactions, 135

575

Subject Index

Epothilone polyketide synthase assembly line, 296–297 engineering, 298 epothilone structures, 121 intermodular interactions, 135 stuttering modules, 132 Epoxidase, polyether biosynthesis, 198–199 Epoxide hydrolase, polyether biosynthesis, 199–201 Erythromycin A biosynthesis, 10, 115 structure, 4, 115 Etnangien structure, 122, 169 synthesis, 170 Extender units, see Polyketide synthase extender units F FAS, see Fatty acid synthase Fatty acids, see Bacterial fatty acids; Cell wall, Mycobacteria tuberculosis; Fatty acid synthase Fatty acid synthase classification, 19 domain organization, 19–21 electron microscopy, 18 fatty acid biosynthesis, 6 polyunsaturated fatty acid synthesis gene clusters, 83–84 industrial synthesis, 93, 95 overview, 80–81 pathways, 81–83 tandem acyl carrier protein domains in polyunsaturated fatty acid synthases acyl carrier protein overexpression, 89–91 holoprotein preparation, 92–93 number of active acyl carrier proteins and fatty acid production, 93–94 phosphopantetheinyl transferase overexpression, 91–92 plasmid construction, 88–89 recombinant Shewanella japonica enzyme expression in Escherichia coli, 85–86 site-directed mutagenesis and active site mapping, 86–88 structure acyl carrier protein, 36–37 acyltransferase domain, 22–24 dehydratase domain, 30–32 examples, 21 ketoreductase domains, 27–30 ketosynthase domain, 24–27 nuclear magnetic resonance overview, 18, 21 polyketide synthase homology, 18

prospects for study, 39 X-ray crystallography overview, 18–19, 21 FK228 structure, 169 synthesis, 170 Fortimicin A biosynthetic gene cluster, 463 structure, 461 synthesis, 465 FR-008, see Candidicin Frenolicin, structure, 369 Fungal type I polyketide synthases bikaverin nonaketide synthase expression and purification, 62–63 substrate specificity, 63 domains, 54 fusarin synthetase, 67–68 genome sequencing, 52 lovastatin synthesis Diels-Alder cyclization, 65 expression vectors, 65–66 polyketide synthases, 64 structure of synthases, 66 6-methyl salicylic acid domain architecture, 53–54 purification, 52–53 recombinant protein expression, 54–55 norsolorinic acid synthase aflatoxin synthesis role, 55 domain studies, 57–59 purification, 56 polymerase chain reaction primers, 51–52 products classes, 50 features, 49 structures, 48 structural overview, 49–50 tenellin synthetase, 68–70 tetrahydroxynaphthalene synthase cell-free extract, 60–61 domains, 61–62 expression, 60 functions, 59–60 zearalenone synthesis, 70–72 Fusarin synthetase, see Fungal type I polyketide synthases G Gel filtration, see Size-exclusion chromatography Geldanamycin, structure, 121 Gentamicin C1 biosynthetic gene cluster, 462 structure, 461 L-Glutamine:deoxy-scyllo-inosose aminotransferase, cyclitol-containing aminoglycoside synthesis NAD-dependent dehydrogenase, 503

576

Subject Index

L-Glutamine:deoxy-scyllo-inosose

aminotransferase, cyclitol-containing aminoglycoside synthesis (cont.) overview, 501–502 radical S-adenosylmethionine dehydrogenase, 503–504 L-Glutamine:scyllo-inositol transaminase, cyclitol-containing aminoglycoside synthesis, 472–473 g-L-Glutamyl-acyl carrier protein decarboxylase, butirosin synthesis, 512–513 g-L-Glutamyl-acyl carrier protein ligase, butirosin synthesis, 511–512 g-L-Glutamyl-4-aminobutyryl acyl carrier protein monooxygenase, butirosin synthesis, 513 g-L-Glutamyl-4-amino-2-hydroxybutyryl acyl carrier protein:ribostamycin g-L-glutamyl4-amino-2-hydroxybutyryltransferase, butirosin synthesis, 514 g-L-Glutamyl-butirosin B g-L-glutamyl cyclotransferase, butirosin synthesis, 514–515 Glycosyltransferase cyclitol-containing aminoglycoside synthesis, 476–477 type II polyketide synthase assay, 389 GNAT domain, polyketide synthase, 125 H HCS cassette, b-carbon methylation, 128–130 High-performance liquid chromatography modified acyl carrier protein characterization, 152–154 mycobacterial saturated fatty acids, 282–284 nystatin analogs, 254–256 type II polyketide synthase assay, 379 HMG-CoA synthase, see HCS cassette HPLC, see High-performance liquid chromatography 3-Hydroxyacyl-acyl carrier protein dehydratase assay, 421 fatty acid production in bacteria, 407 3-Hydroxydecanoyl-acyl carrier protein dehydratase, fatty acid production in bacteria, 407–409 Hygromycin B biosynthetic gene cluster, 463 structure, 461 synthesis, 465 I Inositol-2-dehydrogenase, cyclitol-containing aminoglycoside synthesis, 475–476 Iterative type I polyketide synthase acyl carrier protein preparation

apoprotein overexpression and purification, 107–108 holoprotein preparation in vitro, 108–109 domain organization, 99–101 heterologous expression in Escherichia coli expression construct generation, 103–104 histidine-tagged protein overproduction and purification, 104–105 overview, 98–99 polyene intermediate production and isolation coexpression of pksE and pksE10, 106 isolation of polyene product, 106–107 pksE10 expression construct, 105–106 polymerase chain reaction of PKSE cassettes for predictive classification of new enediynes amplification product generation, 102 primers, 101–103 sequencing and phylogenetic analysis, 103 J Jadomycin B, structure, 369 Jamaicamide structure, 121 synthesis, 130 K Kanamycin B biosynthetic gene cluster, 462 structure, 461 synthesis, 465, 477–478 3-Ketoacyl-acyl carrier protein reductase, fatty acid production in bacteria, 406 3-Ketoacyl-acyl carrier protein synthase assay, 421 fatty acid production in bacteria, 404–406 Ketoreductase domain, polyketide synthase engineering for b-carbon processing alteration, 305–307 ketide unit structure determination by domains, 298–299 structure, 27–30 trans-polyketide synthase b-ketoreductase activities, 127–128 type II polyketide synthase assay, 384–386 Ketosynthase acyl carrier protein interaction assay, 380–381 polyketide synthase structure, 24–27 Kirromycin structure, 169 synthesis, 170 KSQ domain, polyketide synthase, 123–124 L Lactimidomycin structure, 169 synthesis, 171

577

Subject Index

Lankacidin C structure, 169 synthesis, 171 Lankacidin polyketide synthase, stuttering modules, 132 Lasalocid A, structure, 189 Leinamycin loading module in polyketide synthase, 125–126 structure, 121 synthesis, 166–168 Lovastatin structure, 4 synthesis Diels-Alder cyclization, 65 expression vectors, 65–66 polyketide synthases, 64 structure of synthases, 66 Lsd19, polyether biosynthesis, 199–201 M Macrolactin structure, 169 synthesis, 171 Maitotoxin, structure, 4–5 Malonyl-Co:acyl carrier protein transacylase, assay, 380, 421 Mannosyl-b–1-phosphomycoketide, synthesis in mycobacteria, 272–273 MAS, dimycocerosate ester synthesis, 267–270 Mass spectrometry acyl carrier proteins characterization, 147–157 polyketide synthase extender units, 158–160 nystatin analogs, 253–254 type II polyketide synthase assay, 379, 381 MAT, see Malonyl-Co:acyl carrier protein transacylase MbtC, siderophore synthesis, 280–281 MbtD, siderophore synthesis, 280–281 Methylation domains, polyketide synthase C-methylation, 126–127 O-methylation, 126 6-Methyl salicylic acid structure, 4 synthase, see Fungal type I polyketide synthases Methyltransferase, type II polyketide synthase assay, 386–387 Mithramycin, structure, 369 MonBI/BII, polyether biosynthesis, 199–201 MonCI, polyether biosynthesis, 198–199 Monensin A biosynthetic gene clusters, 192–194 premonensin intermediate, 195–198 structure, 4, 189 synthesis, 190 Monooxygenase, candidicin synthesis, 234–235

MS, see Mass spectrometry Mupirocin structure, 122, 169 synthesis, 171 Mutasynthesis aminocoumarin antibiotics amide synthetase assays for suitable ring A analog identification, 446–447 genes for expanding product range, 447 generation of new antibiotics, 444–446 polyketide synthase engineering, 313–314 Mycobactin siderophore synthesis, 280–281 structure, 280 Mycolic acid, synthesis in mycobacteria, 273–276 Mycosamine, biosynthesis, 236 Mycothiol, structure, 461 Myxalamid B, structure, 122 Myxothiazole A, structure, 121 Myxovirescen structure, 121, 169 synthesis, 171 N Nanchangmycin biosynthetic genes clusters, 192–193 transcriptional analysis, 209 structure, 189 NanE N-acetylcysteamine thioester substrate preparation, 205–206, 208 assay, 206–208 expression in Escherichia coli, 204 histidine tag removal, 205 polyether biosynthesis overview, 201–204 purification of histidine-tagged protein, 204–205 site-directed mutagenesis, 209 NanI, polyether biosynthesis, 199–201 NanO, polyether biosynthesis, 198–199 Narbomycin, synthesis, 120 Neocarzinostatin, structure, 4 Neomycin biosynthetic gene cluster, 495–497 structure, 461 synthesis 2000 -N-acetyl-6000 -deamino6000 hydroxyneomycin C deacetylase, 509 20 -N-acetylparomamine deacetylase, 505 20 -N-acetylparomamine synthase, 504 6000 -deamino-6000 -dehydro-6000 oxoneomycin, 510 6000 -deamino-6000 -hydroxyneomycin C dehydrogenase, 509–510

578 Neomycin (cont.) 60 -dehydro-60 -oxoparomamine aminotransferase, 506–507 2-deoxy-scyllo-inosose synthase, 500–501 L-glutamine:deoxy-scyllo-inosose aminotransferase NAD-dependent dehydrogenase, 503 overview, 501–502 radical S-adenosylmethionine dehydrogenase, 503–504 overview, 465, 495.498 paromamine 60 -dehydrogenase, 505–506 phosphoribostamycin phosphatase, 508 phosphoribostamycin synthase, 507–508 UDP-GlcNAc:ribostamycin N-acetylglucosaminyltransferase, 508–509 NigBI/BII, polyether biosynthesis, 199–201 NigCI, polyether biosynthesis, 198–199 Nigericin biosynthetic gene clusters, 192–193 structure, 189 NMR, see Nuclear magnetic resonance Norsolorinic acid synthase, see Fungal type I polyketide synthases Novobiocin, see also Aminocoumarin antibiotics biosynthetic gene clusters deletions in integrative cosmids, 442–444 heterologous expression, 441–442 structure, 439 Nuclear magnetic resonance nystatin analogs, 254 polyketide synthase/fatty acid synthase structure acyl carrier protein, 36–37 acyltransferase domain, 22–24 aromatase/cyclases, 32–36 dehydratase domain, 30–32 examples, 21 ketoreductase domains, 27–30 ketosynthase domain, 24–27 overview, 18, 21 prospects for study, 39 thioesterase, 37–39 protein size limitations in structure elucidation, 18 Nystatin A1 analogs liquid chromatography–mass spectrometry purification, 255–256 production and identification, 254–255 scaled-up production, 255 antifungal activity, 244 biosynthesis gene clusters, 244–246 gene modification in Streptomyces nodosus overview, 249–252

Subject Index

phage-mediated gene replacement, 252–254 gene modification in Streptomyces noursei conjugative transfer of plasmid, 247–248 inactivation, 248 overview, 246–247 replacement, 248–249 structure, 244 O Oleandomycin, synthesis, 120 Onnamide A structure, 169 synthesis, 171 Orsellinic acid, structure, 4 Oxazolomycin structure, 169 synthesis, 172 Oxygenase, type II polyketide synthase assay, 387–389 Oxytetracycline, structure, 369 P PABA, see p-Aminobenzoic acid biosynthesis, 230, 233 Paromamine 60 -dehydrogenase, cyclitolcontaining aminoglycoside synthesis, 505–506 PCR, see Polymerase chain reaction Pederin structure, 4, 121, 169 synthesis, 172 Phoslactomycin B structure, 121 synthesis, 128 Phosphopantetheinyl transferase acyl carrier protein activation, 118 holoprotein preparation, 92–93 overexpression, 91–92 Phosphoribostamycin phosphatase, cyclitolcontaining aminoglycoside synthesis, 508 Phosphoribostamycin synthase, cyclitolcontaining aminoglycoside synthesis, 507–508 Pik polyketide synthase, intermodular interactions, 135–136 Pikromycin, synthesis, 120 Pimaricin applications, 218 biosynthesis in Streptomyces natalensis cholesterol oxidase regulation, 225 gene cluster, 218, 220 global regulation, 226–227 inducers, 226 pimarocinolide synthase complex, 218, 220, 222

Subject Index

pimarocinolide tailoring and export, 223–224 scheme, 221 transcriptional regulation, 224–225 discovery, 217 structure, 216–219 PKS, see Polyketide synthase PKSE, see Iterative type I polyketide synthase Platenolide, synthesis, 120 Polyketide production, heterologous bacteria Escherichia coli as heterologous host, 357–359 general considerations, 340–344 overview, 340–341 polyketide synthase engineering, see Polyketide synthase Pseudomonas putida as heterologous host, 359–360 Streptomyces coelicor conjugation protocol optimization Escherichia coli–Streptomyces cross-streak technique, 354–355 Saccharopolyspora erythraea, 354 large polyketide synthase genes assembly, 349–350 bacterial artificial chromosome plasmids, 350 multiplasmid approach, 350–351 model system features, 344–347 overview of steps, 347–349 polyketide titer improvement high copy number plasmids, 351 superhost vector systems, 351–353 precursor supply optimization enhancement of supply, 355–356 precursor pathway incorporation, 356–357 Polyketide synthase, see also Acyltransferase-less type I polyketide synthases; Deoxyerythronolide B synthase; Fungal type I polyketide synthases; Iterative type I polyketide synthase; specific products engineering acyltransferase replacements, 300–305 a-carbon alterations, 300–301 b-carbon processing, 305–307 chemobiosynthesis, 311, 313 gene knockouts, 314–315 heterologous expression of proteins, 309–312 ketide unit structure determination by domain organization, 298–299 mutasynthesis, 313–314 overview, 296–298 pathway gene design and synthesis assembly of large synthases, 330–332 module combination optimization assay, 327, 329–330

579 operon design and construction for sugar pathway gene expression, 332–335 overview, 320–322 restriction site incorporation for expression in Escherichia coli, 322–324 validation of synthetic gene design, 324–327 single crossover events, 308–309 strategies, 300 extender units, see Polyketide synthase extender units modular enzymes, 10–12 structure acyl carrier protein, 36–37 acyltransferase domain, 22–24 aromatase/cyclases, 32–36 dehydratase domain, 30–32 examples, 21 fatty acid synthase homology, 18 ketoreductase domains, 27–30 ketosynthase domain, 24–27 nuclear magnetic resonance overview, 18, 21 prospects for study, 39 thioesterase, 37–39 X-ray crystallography overview, 18–19, 21 type II enzymes, see Type II polyketide synthase types and features, 6–12 Polyketide synthase extender units, acyl carrier protein-linked biosynthesis, 144–146 zwittermicin A biosynthesis system reconstitution and characterization acylation studies, 161 (2R)-aminomalonyl-acyl carrier protein formation, 157–158 ATP/pyrophosphate exchange assays, 160–161 (2R)-hydroxymalonyl-acyl carrier protein formation, 154–157 mass spectrometry analysis of acyl carrier proteins, 158–160 phosphopantetheinylation of acyl carrier proteins, 151–152 recombinant protein overproduction and purification materials, 147–148 nickel affinity chromatography of histidine-tagged proteins, 149–151 overproduction in Escherichia coli, 148–149 principles, 147 reverse-phase high-performance liquid chromatography characterization of modified acyl carrier proteins, 152–154

580

Subject Index

Polyketide synthase extender units, acyl carrier protein-linked (cont.) serine modification studies, 161 Polymerase chain reaction fungal type I polyketide synthase primers, 51–52 PKSE cassettes for predictive classification of new enediynes amplification product generation, 102 primers, 101–103 sequencing and phylogenetic analysis, 103 Pradimicin, structure, 369 Premonensins structures, 121 synthesis, 195–198 Pthenoic acid, synthesis in mycobacteria, 277–278 R Rapamycin structure, 121 synthesis, 124 Rhizoxin structure, 121, 169 synthesis, 172 Rifamycin, structure, 121 Rosaramidin, synthesis, 120 S Salinomycin biosynthetic gene clusters, 193–194 structure, 189 Siderophore, synthesis in mycobacteria, 280–281 Simocyclinone D8, structure, 439 Single crossover event, polyketide synthase engineering Site-directed mutagenesis acyl carrier protein active site mapping, 86–88 NanE, 209 Size-exclusion chromatography acyl carrier proteins, 425 thymidine diphosphate-a-D-glucose, 527–528 Skipping modules, polyketide synthase, 133–134 Soraphen A, structure, 122 Spectinomycin biosynthetic gene cluster, 462 structure, 461 synthesis, 465 Squalestatin S1, structure, 4 Stigmatellin A, structure, 121 Stigmatellin polyketide synthase, stuttering modules, 132 Streptomycin biosynthetic gene cluster, 462 structure, 461 synthesis, 465, 469–477

Streptomycin-phosphate phosphatase, cyclitolcontaining aminoglycoside synthesis, 473–474 Stuttering modules, polyketide synthase, 134–135 Sulfolipids, synthesis in mycobacteria, 270–262 T Tenellin synthetase, see Fungal type I polyketide synthases Tetracenomycin, structure, 369 Tetrahydroxynaphthalene synthase, see Fungal type I polyketide synthases Tetronomycin, structure, 189 Thioesterase NanE and polyether biosynthesis N-acetylcysteamine thioester substrate preparation, 205–206, 208 assay, 206–208 expression in Escherichia coli, 204 histidine tag removal, 205 overview, 201–204 purification of histidine-tagged protein, 204–205 site-directed mutagenesis, 209 polyketide synthase structure, 37–39 Thymidine diphosphate deoxysugars biosynthetic pathway reconstitution overview, 533–534 thymidine diphosphate-a-D-desosamine synthesis, 538–539 thymidine diphosphate-a-D-forosamine synthesis, 536–537 thymidine diphosphate-a-D-mycaminose one-pot synthesis, 534–535 thymidine diphosphate-b-L-mycarose synthesis, 535–536 glycosylation, 522–523 metabolic pathway engineering for in vivo synthesis, 539–541 thymidine diphosphate-a-D-glucose enzymatic synthesis incubation conditions, 526–527 overview, 524–525 recombinant enzyme preparation, 525–526 purification liquid chromatography and desalting, 528–529 size-exclusion chromatography, 527–528 thymidine diphosphate-4-keto-6-deoxy-a-Dglucose 2,6-dideoxysugar generation, 531–533 enzymatic synthesis overview, 529–531 Tylactone, synthesis, 120 Tylosin, synthesis, 120 Type II polyketide synthase assays of individual enzyme activities

581

Subject Index

aromatase/cyclases, 384–386 glycosyltransferases, 389 ketoreductase, 384–386 methyltransferase, 386–387 oxygenases, 387–389 components, 368, 370 enzymatic total synthesis, 389 expression and purification acyl carrier protein expression and modification, 372–373 Escherichia coli as heterologous host, 371–372 Streptomyces as heterologous host, 373–376 gene clusters, 368 minimal activity assays ketosynthase–acyl carrier protein interaction assay, 380–381 malonyl-Co:acyl carrier protein transacylation assay, 380 mass spectrometry, 381 product assays, 376–380 products, 368–369 starter unit synthesis and incorporation assays direct priming of minimal polyketide synthase, 382 overview, 381 starter unit biosynthesis, 382–384 U UDP-GlcNAc:ribostamycin N-acetylglucosaminyltransferase, neomycin synthesis, 508–509 V Validamycin A biosynthetic gene cluster, 463 structure, 461 synthesis, 465 Virginiamycin M structure, 122, 169 synthesis, 172

X X-ray crystallography, polyketide synthase/fatty acid synthase structures acyl carrier protein, 36–37 acyltransferase domain, 22–24 aromatase/cyclases, 32–36 dehydratase domain, 30–32 examples, 21 ketoreductase domains, 27–30 ketosynthase domain, 24–27 overview, 18–19, 21 prospects for study, 39 thioesterase, 37–39 Z Zearalenone, synthesis, 70–72 Zwittermicin A, biosynthesis system reconstitution and characterization acylation studies, 161 (2R)-aminomalonyl-acyl carrier protein formation, 157–158 ATP/pyrophosphate exchange assays, 160–161 (2R)-hydroxymalonyl-acyl carrier protein formation, 154–157 mass spectrometry analysis of acyl carrier proteins, 158–160 phosphopantetheinylation of acyl carrier proteins, 151–152 recombinant protein overproduction and purification materials, 147–148 nickel affinity chromatography of histidinetagged proteins, 149–151 overproduction in Escherichia coli, 148–149 principles, 147 reverse-phase high-performance liquid chromatography characterization of modified acyl carrier proteins, 152–154 serine modification studies, 161

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