Methods in Enzymology 468 - Biophysical, Chemical, and Functional Probes of RNA Structure, Interactions and Folding:, Part 1

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

Author: Daniel Herschlag

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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: 978-0-12-374399-2 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

Tadepalli Adilakshmi Weis Center for Research, Geisinger Medical Center, Danville, Pennsylvania, USA Philip C. Bevilacqua Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania, USA Durga M. Chadalavada Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania, USA Eric L. Christian Center for RNA Molecular Biology, Department of Biochemistry, Case Western Reserve University School of Medicine, Cleveland, Ohio, USA Gloria M. Culver Department of Biology, University of Rochester, Rochester, New York, USA Victoria J. DeRose Department of Chemistry, University of Oregon, Eugene, Oregon, USA Caia D. S. Duncan Department of Chemistry, University of North Carolina, Chapel Hill, North Carolina, USA Martha J. Fedor Departments of Chemical Physiology and Molecular Biology, The Skaggs Institute for Chemical Biology, The Scripps Research Institute, La Jolla, California, USA Andrew L. Feig Department of Chemistry, Wayne State University, Detroit, Michigan, USA Marcello Forconi Department of Biochemistry, Stanford University, Stanford, California, USA John K. Frederiksen The Pritzker School of Medicine, and Department of Biochemistry and Molecular Biology, University of Chicago, Chicago, Illinois, USA, and Department of Pathology and Laboratory Medicine, University of Rochester Medical Center, Rochester, New York, USA

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Contributors

Michael E. Harris Center for RNA Molecular Biology, Department of Biochemistry, Case Western Reserve University School of Medicine, Cleveland, Ohio, USA Daniel Herschlag Departments of Biochemistry and Chemistry, Stanford University, Stanford, California, USA James L. Hougland Department of Chemistry, University of Michigan, Ann Arbor, Michigan, USA Laura Hunsicker-Wang Department of Chemistry, Trinity University, San Antonio, Texas, USA Alain Laederach Developmental Genetics and Bioinformatics, Wadsworth Center, Albany, New York, USA Andreas Liebeg Max F. Perutz Laboratories, Department of Biochemistry and Cell Biology, University of Vienna, Vienna, Austria Joshua S. Martin Developmental Genetics and Bioinformatics, Wadsworth Center, Albany, New York, USA Jennifer L. McGinnis Department of Chemistry, University of North Carolina, Chapel Hill, North Carolina, USA David Mitchell III Department of Chemistry and Biochemistry, Institute for Cellular and Molecular Biology, University of Texas at Austin, Austin, Texas, USA Somdeb Mitra Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York, USA Tao Pan Department of Biochemistry & Molecular Biology, University of Chicago, Chicago, Illinois, USA Joseph A. Piccirilli Departments of Chemistry and Biochemistry & Molecular Biology, University of Chicago, Chicago, Illinois, USA Rick Russell Department of Chemistry and Biochemistry, Institute for Cellular and Molecular Biology, University of Texas at Austin, Austin, Texas, USA

Contributors

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Susan J. Schroeder Department of Chemistry and Biochemistry, University of Oklahoma, Norman, Oklahoma, USA Inna Shcherbakova Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York, USA, and Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts, USA Katrina Simmons Developmental Genetics and Bioinformatics, Wadsworth Center, Albany, New York, USA Sarah F. C. Soper Program in Cell, Molecular and Developmental Biology and Biophysics, Johns Hopkins University, Baltimore, Maryland, USA Scott A. Strobel Department of Molecular Biophysics and Biochemistry and Department of Chemistry, Yale University, New Haven, Connecticut, USA Ian T. Suydam Department of Molecular Biophysics and Biochemistry and Department of Chemistry, Yale University, New Haven, Connecticut, USA Douglas H. Turner Department of Chemistry, University of Rochester, Rochester, New York, USA Matthew Vogt Laboratory of Pathology, National Cancer Institute, Bethesda, Maryland, USA Christina Waldsich Max F. Perutz Laboratories, Department of Biochemistry and Cell Biology, University of Vienna, Vienna, Austria Yaqi Wan Department of Chemistry and Biochemistry, Institute for Cellular and Molecular Biology, University of Texas at Austin, Austin, Texas, USA Peter Y. Watson Departments of Chemical Physiology and Molecular Biology, The Skaggs Institute for Chemical Biology, The Scripps Research Institute, La Jolla, California, USA Kevin M. Weeks Department of Chemistry, University of North Carolina, Chapel Hill, North Carolina, USA Terrence N. Wong Department of Biochemistry & Molecular Biology, University of Chicago, Chicago, Illinois, USA

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Contributors

Sarah A. Woodson T.C. Jenkins Department of Biophysics, Johns Hopkins University, Baltimore, Maryland, USA Zhili Xu Department of Biology, University of Rochester, Rochester, New York, USA

PREFACE

After the discovery of catalytic RNA nearly 30 years ago, and after the initial excitement wore off, RNA was viewed predominantly as an ancient biological macromolecule with vestigial, albeit critical, functions in modern-day biology. Thus, while important and informative, studies of RNA behavior and function took a back seat to the interrogation of transcription factors, protein kinases, and other molecules directly involved in the regulation of gene expression. But more recent discoveries have clearly illuminated the central importance of RNA to modern-day biology—the discovery of fewer genes but vastly more alternative spliced gene products than anticipated in the human genome, the discovery of RNA regulatory elements in the form of riboswitches, the finding of many noncoded RNAs, the finding of functional groupings of RNAs by RNA binding proteins, and, of course, the discovery of RNA interference (RNAi) and its likely role in regulation of about half of all human genes. Thus, it is now clear that we need to understand these molecules in order to understand how biology works, and applications to understanding and curing diseases, while still largely remote, are ultimately likely to become common. Fortunately, in the years since the discovery of catalytic RNA, many incisive and powerful chemical, biochemical, and biophysical tools have been developed to study the folding and conformational behavior of RNA. Here, we have assembled many of these together, in this two volume series. Descriptions of two of the most common and powerful techniques are not covered, NMR and X-ray crystallography (although there is a chapter on preparation of RNA for crystallography), as these approaches can warrant volumes on their own and have been treated separately (see, e.g., Volume 394 on Biological NMR). I believe that this compilation is particularly important, not just because of the importance of the subject matter, but because the tools and contributors span previously distinct fields from molecular biology, biochemistry, chemistry, and physics. Now students and researchers in each of these areas can get a sense of, as well as detailed protocols for, the entire gambit of approaches. I hope that the efforts of the many contributors will inspire new students and investigators to join the search for understanding of these most fascinating macromolecules. And finally I want to thank a phenomenal group of contributors for agreeing to contribute and then coming through, even sometimes on time, with chapters of uniform high quality. DAN HERSCHLAG xv

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

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


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

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


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

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


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

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


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

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


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VOLUME 381. Oxygen Sensing Edited by CHANDAN K. SEN AND GREGG L. SEMENZA VOLUME 382. Quinones and Quinone Enzymes (Part B) Edited by HELMUT SIES AND LESTER PACKER VOLUME 383. Numerical Computer Methods (Part D) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 384. Numerical Computer Methods (Part E) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 385. Imaging in Biological Research (Part A) Edited by P. MICHAEL CONN VOLUME 386. Imaging in Biological Research (Part B) Edited by P. MICHAEL CONN VOLUME 387. Liposomes (Part D) Edited by NEJAT DU¨ZGU¨NES, VOLUME 388. Protein Engineering Edited by DAN E. ROBERTSON AND JOSEPH P. NOEL VOLUME 389. Regulators of G-Protein Signaling (Part A) Edited by DAVID P. SIDEROVSKI VOLUME 390. Regulators of G-Protein Signaling (Part B) Edited by DAVID P. SIDEROVSKI VOLUME 391. Liposomes (Part E) Edited by NEJAT DU¨ZGU¨NES, VOLUME 392. RNA Interference Edited by ENGELKE ROSSI VOLUME 393. Circadian Rhythms Edited by MICHAEL W. YOUNG VOLUME 394. Nuclear Magnetic Resonance of Biological Macromolecules (Part C) Edited by THOMAS L. JAMES VOLUME 395. Producing the Biochemical Data (Part B) Edited by ELIZABETH A. ZIMMER AND ERIC H. ROALSON VOLUME 396. Nitric Oxide (Part E) Edited by LESTER PACKER AND ENRIQUE CADENAS VOLUME 397. Environmental Microbiology Edited by JARED R. LEADBETTER VOLUME 398. Ubiquitin and Protein Degradation (Part A) Edited by RAYMOND J. DESHAIES VOLUME 399. Ubiquitin and Protein Degradation (Part B) Edited by RAYMOND J. DESHAIES

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VOLUME 400. Phase II Conjugation Enzymes and Transport Systems Edited by HELMUT SIES AND LESTER PACKER VOLUME 401. Glutathione Transferases and Gamma Glutamyl Transpeptidases Edited by HELMUT SIES AND LESTER PACKER VOLUME 402. Biological Mass Spectrometry Edited by A. L. BURLINGAME VOLUME 403. GTPases Regulating Membrane Targeting and Fusion Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 404. GTPases Regulating Membrane Dynamics Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 405. Mass Spectrometry: Modified Proteins and Glycoconjugates Edited by A. L. BURLINGAME VOLUME 406. Regulators and Effectors of Small GTPases: Rho Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 407. Regulators and Effectors of Small GTPases: Ras Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 408. DNA Repair (Part A) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 409. DNA Repair (Part B) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 410. DNA Microarrays (Part A: Array Platforms and Web-Bench Protocols) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 411. DNA Microarrays (Part B: Databases and Statistics) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 412. Amyloid, Prions, and Other Protein Aggregates (Part B) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 413. Amyloid, Prions, and Other Protein Aggregates (Part C) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 414. Measuring Biological Responses with Automated Microscopy Edited by JAMES INGLESE VOLUME 415. Glycobiology Edited by MINORU FUKUDA VOLUME 416. Glycomics Edited by MINORU FUKUDA VOLUME 417. Functional Glycomics Edited by MINORU FUKUDA


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VOLUME 418. Embryonic Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA 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

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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 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


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VOLUME 449. RNA Turnover in Eukaryotes: Analysis of Specialized and Quality Control RNA Decay Pathways Edited by LYNNE E. MAQUAT AND MEGERDITCH KILEDJIAN VOLUME 450. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 451. Autophagy: Lower Eukaryotes and Non-Mammalian Systems (Part A) Edited by DANIEL J. KLIONSKY VOLUME 452. Autophagy in Mammalian Systems (Part B) Edited by DANIEL J. KLIONSKY VOLUME 453. Autophagy in Disease and Clinical Applications (Part C) Edited by DANIEL J. KLIONSKY VOLUME 454. Computer Methods (Part A) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 455. Biothermodynamics (Part A) Edited by MICHAEL L. JOHNSON, JO M. HOLT, AND GARY K. ACKERS (RETIRED) VOLUME 456. Mitochondrial Function, Part A: Mitochondrial Electron Transport Complexes and Reactive Oxygen Species Edited by WILLIAM S. ALLISON AND IMMO E. SCHEFFLER VOLUME 457. Mitochondrial Function, Part B: Mitochondrial Protein Kinases, Protein Phosphatases and Mitochondrial Diseases Edited by WILLIAM S. ALLISON AND ANNE N. MURPHY VOLUME 458. Complex Enzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles and Peptides Edited by DAVID A. HOPWOOD VOLUME 459. Complex Enzymes in Microbial Natural Product Biosynthesis, Part B: Polyketides, Aminocoumarins and Carbohydrates Edited by DAVID A. HOPWOOD VOLUME 460. Chemokines, Part A Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 461. Chemokines, Part B Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 462. Non-Natural Amino Acids Edited by TOM W. MUIR AND JOHN N. ABELSON VOLUME 463. Guide to Protein Purification, 2nd Edition Edited by RICHARD R. BURGESS AND MURRAY P. DEUTSCHER VOLUME 464. Liposomes, Part F Edited by NEJAT DU¨ZGU¨NES,

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VOLUME 465. Liposomes, Part G Edited by NEJAT DU¨ZGU¨NES, VOLUME 466. Biothermodynamics, Part B Edited by MICHAEL L. JOHNSON, GARY K. ACKERS, AND JO M. HOLT VOLUME 467. Computer Methods, Part B Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 468. Biophysical, Chemical, and Functional Probes of RNA Structure, Interactions and Folding, Part A Edited by DANIEL HERSCHLAG

C H A P T E R

O N E

Nucleotide Analog Interference Mapping Ian T. Suydam and Scott A. Strobel Contents 4 7 7 8 8 8 9

1. Introduction 2. Materials and Reagents 2.1. Chemicals 2.2. Equipment 2.3. Enzymes 2.4. Buffers 3. Methods 3.1. Synthesis of phosphorothioate-tagged nucleoside triphosphates 3.2. Incorporation of phosphorothioate analogs by in vitro transcription 3.3. Incorporation efficiency controls 3.4. Activity selections 3.5. Quantitating sites of interference 4. Properties of Analogs 4.1. Ribose modifications 4.2. Base modifications 5. Nucleotide Analog Interference Suppression 6. Conclusions References

9 12 15 16 18 19 21 23 25 26 27

Abstract Nucleotide analog interference mapping (NAIM) is a powerful chemogenetic technique that rapidly identifies chemical groups essential for RNA function. Using a series of phosphorothioate-tagged nucleotide analogs, each carrying different modifications of nucleobase or backbone functionalities, it is possible to simultaneously, yet individually, assess the contribution of particular functional groups to an RNA’s activity at every position within the molecule. In contrast to traditional mutagenesis, which modifies RNA on the nucleobase Department of Molecular Biophysics and Biochemistry and Department of Chemistry, Yale University, New Haven, Connecticut, USA Methods in Enzymology, Volume 468 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)68001-0

#

2009 Elsevier Inc. All rights reserved.

3

4

Ian T. Suydam and Scott A. Strobel

level, the smallest mutable unit in a NAIM analysis is a single atom, providing a detailed description of interactions at critical nucleotides. Because the method introduces modified nucleotides by in vitro transcription, NAIM offers a straightforward and efficient approach to study any RNA that has a selectable function, and it can be applied to RNAs of nearly any length.

1. Introduction Structured RNAs rely on a large and diverse collection of functional group interactions to form folded states in active conformations (Noller, 2005). One approach to studying these interactions is the site-specific modification of functional groups, followed by a direct assay of activity. These experiments are most often performed using chimeric RNAs, typically generated by reassembling the full-length sequence from a substituted synthetic oligonucleotide and a truncated RNA transcript (Das and Piccirilli, 2005; Lafontaine et al., 2002). Although powerful, these techniques are limited by the difficulty of assembling and measuring the effect of each singly substituted RNA. For this reason, site-specific substitution experiments are often restricted to a small number of modifications at a minimal number of sites. In contrast, the method described in this section provides a semiquantitative approach to rapidly assay the effect of a nucleotide modification at all sites simultaneously. Nucleotide analog interference mapping (NAIM) utilizes a series of 50 -O-(1-thio)nucleoside triphosphate analogs, which are randomly incorporated at low frequency into an RNA with a selectable function. The method relies on two advantageous properties of nucleoside phosphorothioates. First is the observation that 50 -O-(1-thio)nucleoside triphosphates can be incorporated by T7 RNA polymerase (Eckstein, 1985). In these transcriptions, the polymerase accepts only the Sp diastereomer, a result explained by the Mg2þ coordination pattern observed in recent crystal structures (Fig. 1.1A) (Yin and Steitz, 2004). Because incorporation proceeds with inversion of configuration, the resulting RNA transcripts contain Rp phosphorothioate linkages (Griffiths et al., 1987). When parental phosphorothioate-tagged nucleotides (NTPaS) are used, they are incorporated as readily as the corresponding nucleoside triphosphates (NTPs) (Christian and Yarus, 1992). For these phosphorothioate-tagged nucleotides, the level of incorporation is directly related to the molar ratios of NTPaS and NTP in the transcription reaction (Fig. 1.1B). The second property of phosphorothioates exploited by NAIM is the selective cleavage of phosphorothioate linkages by treatment with iodine (Fig. 1.1C). Although specific, the extent of cleavage at substituted sites is only 15%, which may indicate the formation of a phosphotriester intermediate, the

5

Nucleotide Analog Interference Mapping

CTP

GTP

ATP ATPaS

Y639

UTP

B

A

*

...AGCACAGCA... T7 RNAP

...AGCACAGCA... ...AGCACAGCA...

NTP concentration

ab mATP 3⬘-NT Mg

C O O

O

P O− O O

Rp

O O

Mg

− P S O

O

Base

O

O

I2

OH Analog

O

P O− O

O

P

O

Base

P O− O O

H2O

O

O O O

Analog

~15%

O

Base

O P − O O HO

O

OH

O

O

OH O

Analog

OH

Figure 1.1 (A) Crystal structure of a-, b-methylene ATP bound to the active site of T7 RNA polymerase (PDB #1s76). The location of the sulfur substitution in Sp phosphorothioate-tagged triphosphates is indicated with an asterisk. (B) Schematic of ATPaS incorporation by T7 RNA polymerase leading to transcripts with a low level of substitution at random sites (underlined). (C) Cleavage of resulting Rp phosphorothioate linkage with iodine.

hydrolysis of which produces cleavage products or phosphodiester linkages (Gish and Eckstein, 1988). This selective cleavage has been used in RNA sequencing strategies (Gish and Eckstein, 1988), and provides a method of marking the site of functional group modification by the inclusion of a phosphorothioate tag. The NAIM method expands on the technique of interference mapping by phosphorothioate substitution, and the initial work of Krupp and coworkers (Conrad et al., 1995; Gaur and Krupp, 1993; Ruffner and Uhlenbeck, 1990). The method uses a series of 50 -O-(1-thio)nucleoside triphosphate analogs (dTPaS), each containing an incremental chemical modification in the nucleobase or ribose sugar, and consists of four basic steps. First, the parental and modified phosphorothioate-tagged triphosphates are randomly incorporated into the RNA by in vitro transcription (Fig. 1.2, step 1). In this step, the level of analog incorporation is adjusted such that any given transcript has approximately one site of substitution. Next, the pool of transcribed RNAs is subjected to a selection in which the active and inactive members are either physically separated or selectively

6


1. Incorporation of phosphorothioate-tagged parent nucleotide (N) or modified analog (d ) 1 N 1 d 2 N 2 d 3 N 3 d 4 N d 4 5 N 5 d 6 N 6 d 2. Selection of functional RNAs Unselected controls

1 2 3 4 5 6

* d 1* N * d * 3* N d * 4* N d * 5* N d * 6* N d 3. Cleavage of phosphorothioate linkage with iodine

* * * * * *

1 2 3 4 5 6

3*

d

d

I2

1*

1*

3 4 5 6

3*

* * * *

d

5* 6*

I2

* * * * * *

N-unselected

1 2 3 4 5 6

I2

1* d

5* 6*

d -selected

I2

N-selected

* N * N * N * N * N * N

d -unselected

1 2 3 4 5 6

Selected RNAs

Uncleaved RNA 6 5 4. Electrophoresis and autoradiography

4

Analog effect

3 2

Phosphorothioate effect

1

Figure 1.2 Schematic of the NAIM experiment. In this example, parental and analogmodified nucleotides are incorporated with equal efficiency at all sites. In the activity selected pool the Rp phosphorothioate interferes at position 2, while the modified analog interferes at position 4.


7

labeled (Fig. 1.2, step 2). Unselected RNAs from the original pool are also labeled to serve as controls. Each sample is then treated with iodine to selectively cleave the RNAs at the site of analog substitution (Fig. 1.2, step 3). Finally, the analog distribution at each site in the selected and unselected samples is revealed by resolving the cleavage products with polyacrylamide gel electrophoresis (PAGE) (Fig. 1.2, step 4). Sites of analog substitution that interfere with the selected function are identified as gaps in the sequencing ladder from the active pool of RNAs. Selections carried out with phosphorothioate-tagged parental nucleotides control for sites of interference that arise from the backbone sulfur substitution. Because every position in the sequence is resolved as a unique band on the sequencing gel, a single screen defines the effect of a particular modification at every site in the RNA. NAIM can be expanded to use any analog that is incorporated by an RNA polymerase and can be applied to any RNA with a selectable function. Examples of RNA functions previously investigated with NAIM include RNA folding (Schwans et al., 2003; Waldsich and Pyle, 2007), catalytic activity (Boudvillain and Pyle, 1998; Kazantsev and Pace, 1998; Oyelere et al., 2002; Ryder and Strobel, 1999; Sood et al., 2002; Strobel and Shetty, 1997), and the binding of proteins (Batey et al., 2000), metal ions (Basu et al., 1998), and small molecule metabolites ( Jansen et al., 2006; Kwon and Strobel, 2008). Many of these studies have identified specific interactions or RNA structural motifs that have been confirmed with subsequent atomic resolution structures (Adams et al., 2004; Klein and Ferre-D’Amare, 2006; Ryder and Strobel, 2002; Toor et al., 2008). More recently, the technique has been extended to study RNA interactions in vivo (Kolev and Steitz, 2006; Szewczak et al., 2002). Several descriptions of the NAIM method are currently available (Ryder et al., 2000; Strobel, 1999), including detailed protocols for specific applications (Cochrane and Strobel, 2004; Waldsich, 2008). Here, we focus on the methods required to synthesize and employ new analogs, including guidance in applying NAIM to new systems. Whenever multiple protocols have been used, we present those that have most efficiently produced high-quality results in a number of systems.

2. Materials and Reagents 2.1. Chemicals

50 -O-(1-thio)nucleoside triphosphates. There are currently 20 commercially available phosphorothioate-tagged nucleoside triphosphates, including each parental NTPaS (Glen Research). A number of unprotected nucleosides are also available from a variety of sources that can be used

8


to synthesize phosphorothioate-tagged nucleotide triphosphates (SigmaAldrich, RI Chemical, Toronto Research Chemical, TCI America, Berry & Associates). [g-32P]-adenosine 50 -triphosphate (6000 Ci/mmol, PerkinElmer #NEG035C). All other chemicals and reagents were purchased from Sigma-Aldrich.

2.2. Equipment

Phosphor autoradiography. All sequencing gels were visualized and quantitated with a Storm 820 Phosphorimager using ImageQuant 5.2 software (GE Healthcare). AKTA prime FPLC purification system (GE Healthcare). HiTrap Q HP anion-exchange columns (GE Healthcare #17-1153-01). QIAquick Nucleotide Removal Kit (Qiagen #28304).

2.3. Enzymes

Appropriate Restriction Endonuclease (New England Biolabs). T7 RNA polymerase. Overexpressed from plasmid pT7-911Q (T.E. Shrader, personal communication) or purchased (New England Biolabs #M0251). Y637F mutant form of T7 RNA polymerase. Overexpressed from plasmid pDPT7-Y639F as described (Sousa and Padilla, 1995) or purchased (Epicentre Biotechnologies #D7P9201K). Escherichia coli Inorganic Pyrophosphatase (Sigma #I5907). Calf Intestinal Alkaline Phosphatase (New England Biolabs #M0290). T4 Polynucleotide Kinase (New England Biolabs #M0201).

2.4. Buffers

TEAB. Triethylammonium bicarbonate, pH 8.0. Prepared by bubbling CO2 through a suspension of triethylamine in water. Transcription buffer. 40 mM Tris–HCl (pH 8.0), 2.0 mM spermidine, 5.0 mM dithiothreitol (DTT), 0.005% Triton X-100, MgCl2 (dependent on transcript). Elution buffer. 10 mM Tris–HCl (pH 7.5), 250 mM NaCl, 2% sodium dodecyl sulfate (SDS). PCA. Mixture of phenol/chloroform/isoamyl alcohol in ratio of 25:24:1, saturated with 10 mM Tris–HCl (pH 8.0), 1 mM EDTA. TE. 10 mM Tris–HCl (pH 7.5), 0.1 mM EDTA. VS ribozyme reaction buffer. 50 mM Tris–HCl (pH 7.5), 5 mM MgCl2, 25 mM KCl.


9

2 loading buffer. Prepared in deionized formamide, 20 mM EDTA, 0.05% (w/v) bromophenol blue, 0.05% (w/v) xylene cyanol FF. Iodine/ethanol. 100 mM iodine in 100% ethanol.

3. Methods 3.1. Synthesis of phosphorothioate-tagged nucleoside triphosphates Several methods are available for the synthesis of 50 -O-(1-thio)nucleoside triphosphates, including enzymatic synthesis of pure Sp isomers (Eckstein, 1985). Chemical methods generally produce equimolar mixtures of Rp and Sp diastereomers, which can be separated by reverse-phase HPLC (Fischer et al., 1999). However, these separations are not required for NAIM, since T7 polymerase is selective for the Sp isomer, and transcriptions for NAIM are often carried out using mixtures of both isomers. Although several high yielding chemical syntheses are now available starting from protected nucleosides, the simple one-pot method developed by Arabshahi and Frey (1994) is still preferred for most preparations. The modified procedure outlined below has been used to synthesize a wide range of purine and pyrimidine analogs in 5–40% yield starting from unprotected nucleosides. The synthesis is accomplished using limited equipment with commercially available reagents in less than 2 days. A typical reaction employs 100 mg of starting material, but we have routinely scaled the reaction down to as little as 25 mg of nucleoside. It is worth noting that even on this scale an overall yield of 5% produces sufficient analog triphosphate for 50–500 typical NAIM transcriptions (see below). The conversion of unprotected nucleosides to 50 -O-(1-thio)nucleoside triphosphates is carried out in two steps using a single reaction flask (Fig. 1.3A). In the first step, the nucleoside is converted to the 50 -O-(1,1dichloro-1-thio)phosphorylnucleoside by the addition of thiophosphoryl chloride (PSCl3). This reaction has been shown to be regioselective for the 50 -hydroxyl when carried out in triethyl phosphate (TEP) (Arabshahi and Frey, 1994). In the second step, the initial product is converted directly to the triphosphate by the addition of tributylammonium pyrophosphate (TBAP). This reaction proceeds via a cyclic triphosphate intermediate which is subsequently hydrolyzed with water (Fig. 1.3A). Two factors greatly affect the overall yield of the reaction. The first is the concentration of reactants in the first step, so every effort should be made to dissolve the nucleoside in a minimal volume of TEP, aided by slight heating if necessary. The second and more important consideration is the strict exclusion of water in the initial steps to prevent the hydrolysis of PSCl3 or the initially

10


−

OO P O O O P P O O − S O

A HO

R1 O OH R2

B

PSCl3

2FA

Cl Cl P O S

R1 O

OH R2

TBAP

g

a R1

−

O

OH R2

b

O O O O P O P O P O − − O O S−

R1

H2O

O OH R2

2FATPaS (m/e = 540.2)

Diphosphate (m/e = 460.4) Monophosphate (m/e = 380.5)

Tetraphosphate (m/e = 620.2) Pentaphosphate (m/e = 700.1)

1.0 M TEAB

50 mM TEAB 0

Figure 1.3 (A) Scheme for the synthesis of 5 -O-(1-thio)nucleoside triphosphates, where R1 represents nucleobase analogs and R2 represents 20 -ribose modifications. The unstable cyclic triphosphate intermediate is outlined in brackets. (B) Purification of the phosphorothioate-tagged 2-fluoroadenosine triphosphate by anion-exchange chromatography. The observed mass obtained with negative ion mode mass spectra of each elution peak indicates the formation of mono-, di-, tri-, tetra-, and pentaphosphate products.

formed 50 -O-(1,1-dichloro-1-thio)phosphorylnucleoside. For this reason, all glassware and syringes should be thoroughly dried, anhydrous solvents should be purchased in sealed containers, and reactions should be kept under an atmosphere of dry argon. Because TBAP is hygroscopic the introduction of water with this reagent is of particular concern. We have had good success purchasing a small quantity of the reagent and using it in its entirety after opening. The general procedure that follows has no strict timing requirements and is easily run in parallel, allowing the rapid synthesis of multiple analogs from a single stock of reagents. The nucleoside (100 mg) is transferred to a 20-ml round-bottom flask and dried by coevaporating 3 10 ml of anhydrous pyridine and put under vacuum overnight. Once the nucleoside is dry the reaction flask is capped with a rubber septum and purged with dry argon. The nucleoside is then dissolved in a minimal volume (0.5–2.0 ml) of TEP. For less soluble nucleosides, the reaction flask can be heated gently with an air gun to obtain a clear solution. As was reported in the original description of the synthesis, we have observed reactivity differences between purine and pyrimidine


11

nucleosides and have employed a slightly different procedure for the synthesis of their 50 -O-(1,1-dichloro-1-thio)phosphorylnucleosides. For purine nucleoside analogs, 1.1 equivalents of trioctylamine (MW ¼ 353.7, d ¼ 0.809 g/ml) is added to the stirred nucleoside solution, followed by 1.1 equivalents of PSCl3 (MW ¼ 169.4, d ¼ 1.668 g/ml). This solution is stirred at room temperature for 20–60 min until a significant amount of 50 -O-(1,1-dichloro-1-thio)phosphorylnucleoside has formed. The reaction progress can be monitored qualitatively by quenching a small aliquot (100–200 ml) in water and resolving the products with cellulose thin-layer chromatography using a solvent system of 0.5 M LiCl in water. The reaction products are visualized by shadowing with a hand held UV light, with the monophosphate migrating slower than the starting material. Alternatively, small aliquots (10–20 ml) of reaction can be quenched in 50 mM triethylammonium bicarbonate (TEAB) and loaded directly onto a prepacked anion-exchange column (see below). Elution with a steep gradient of TEAB easily separates the nucleoside from the monophosphate and provides a rapid quantitation of reaction progress. For pyrimidine analogs the dry nucleoside is dissolved in a minimal volume (0.5–2.0 ml) of TEP as above, then cooled to 0 C. To the cooled solution 1.2 equivalents of trioctylamine and 1.2 equivalents of collidine (MW ¼ 121.2, d ¼ 0.931 g/ml) are added, followed by the dropwise addition of 1.3–1.5 equivalents of PSCl3. After stirring at 0 C for 30 min, the reaction is allowed to warm slowly to room temperature and stirred for an additional 45 min with the reaction progress monitored as above. For all subsequent chemical steps and purifications the procedure is the same for purine and pyrimidine analogs. TBAP (MW ¼ 548.7) is dissolved in TEP to a final concentration of 0.1 g/ml by gently heating the solution with an air gun. Once a substantial quantity of 50 -O-(1,1-dichloro-1-thio)phosphorylnucleoside from the preceding step has formed the TBAP/TEP solution is added to the reaction until the final molar ratio of TBAP to PSCl3 has reached 4:1. The addition of 4 equivalents of TBAP increases the overall yield of the triphosphate product but can also increase the quantity of branched higher order phosphate products (see below). The reaction is stirred at room temperature for 30 min and the reaction progress is monitored by rapid ion-exchange chromatography as above. Once a significant amount of triphosphate products are observed the reaction is exposed to air and transferred to a centrifuge tube. The nucleoside triphosphates are precipitated via amine exchange by the dropwise addition of 60 equivalents of triethylamine (TEA, MW ¼ 101.2, d ¼ 0.726 g/ml) to the stirred reaction solution. The precipitate is collected by centrifugation, the supernatant is discarded and the white solid is dissolved in 5 ml of 50 mM TEAB. Incubation of this solution overnight at room temperature hydrolyzes the initially formed cyclic triphosphates to the linear form.

12


The 50 -O-(1-thio)nucleoside triphosphates are purified on prepacked anion-exchange columns using a linear gradient from 50 mM to 1.0 M TEAB while monitoring UV absorption at 260 nm. Poor binding to the matrix is often attributed to residual salt from the reaction and can be improved by further diluting the crude product in 50 mM TEAB before loading. The expected elution fraction can be identified by running a standard sample of the parental nucleoside triphosphate and is typically between 0.5 and 0.6 M TEAB. For some reactions several additional products are observed and these have been identified as the mono-, di-, tetra-, and pentaphosphates (Fig. 1.3B). The fractions containing the nucleoside triphosphate are pooled and lyophilized to a thin film, then redissolved in less than 1.0 ml TE. The pH of this solution should be checked and adjusted to near neutral if necessary to improve the stability of frozen stocks. The 50 -O-(1-thio)nucleoside triphosphates are characterized by UV absorption, 31P NMR, and mass spectroscopy. Because nucleobase modifications can produce significant changes in the UV spectrum it is important to measure extinction coefficients for the free nucleosides and to use these values when calculating nucleotide concentrations. NMR samples are prepared with 10% D2O in buffered solutions (pH 8.0) and should contain approximately 1 mM nucleotide. The correct nucleotide has three resonances in the 31P NMR spectrum at 42 to 43 (a), 5 to 10 (g), and 20 to 25 (b) ppm relative to an external 85% phosphoric acid standard (Arabshahi and Frey, 1994). The distinctive downfield phosphorothioate resonance exists as a pair of doublets in a proton-decoupled spectrum, corresponding to the mixture of Rp and Sp diastereomers. Spectra taken without proton decoupling show additional splitting from the 50 -CH protons, which further confirms the 50 -O-(1-thio) linkage. An impurity that occasionally coelutes with the correct nucleotide and produces a 31P NMR resonance between 33 and 38 ppm results from the reaction of unconsumed PSCl3 with TBAP. Electrospray ionization mass spectra are obtained in negative ion mode from diluted nucleotide samples containing 95% methanol. A strong signal for the singly ionized parent ion is typically observed. Analogs can be stored as dry pellets or TE solutions at 20 C for at least 2 years without substantial degradation.

3.2. Incorporation of phosphorothioate analogs by in vitro transcription Nucleotide analogs are randomly incorporated into RNA by in vitro transcription using T7 RNA polymerase (Eckstein, 1985). For some analogs, particularly those that make modifications in the minor groove, the Y639F point mutant of T7 RNA polymerase greatly improves the level of incorporation (Sousa and Padilla, 1995) (Table 1.1). Transcription conditions are optimized to yield an analog incorporation level of approximately 5%. This level of incorporation provides iodine cleavage bands that are

13


Table 1.1 Conditions to incorporate nucleotide analogs for NAIM experiments Analog dTPaS (Sp isomer only)

[dTPaS] (mM)

[Parent Polymerase NTP] (WT/ (mM) Y639F)

AaS DAPaS 2AP

0.05 0.025 0.50

1.0 1.0 1.0

WT WT WT

m6AaS

0.10

1.0

WT

PuraS

2.0

1.0

WT

c3AaS 7dAaS

1.0 0.05

0.5 1.0

Y639F WT

7F7dAaS 2F7dAaS 2FAaS FormAaS n8AaS dAaS

0.05 0.2 0.2 0.05 0.25 0.75

1.0 1.0 1.0 1.0 1.0 1.0

WT WT WT WT WT Y639F

OMe

2.0

0.2

Y639F

0.25

1.0

Y639F

SH

AaS GaS IaS m2GaS

0.2 0.05 0.1 0.75

1.0 1.0 1.0 0.5

Y639F WT WT Y639F

S6GaS (þ Mn2þ) 7dGaS dGaS OMe GaS SH GaS CaS n6CaS f5CaS CiCaS ZaS dCaS OMe CaS SH CaS UaS

0.25

1.0

WT

Christian and Yarus (1992) Strobel and Shetty (1997) Ortoleva-Donnelly et al. (1998b) Ortoleva-Donnelly et al. (1998b) Ortoleva-Donnelly et al. (1998b) Ryder et al. (2001) Ortoleva-Donnelly et al. (1998b) Suydam and Strobel (2008) Suydam and Strobel (2008) Suydam and Strobel (2008) Ryder et al. (2001) Ryder et al. (2001) Ortoleva-Donnelly et al. (1998b) Ortoleva-Donnelly et al. (1998b) Ortoleva-Donnelly et al. (1998b) Schwans et al. (2003) Christian and Yarus (1992) Strobel and Shetty (1997) Ortoleva-Donnelly et al. (1998a) Basu et al. (1998)

0.05 0.25 2.0 0.25 0.05 0.5 0.5 0.05 0.50 0.75 2.0 0.2 0.05

1.0 1.0 0.1 1.0 1.0 0.5 1.0 1.0 0.5 1.0 0.05 1.0 1.0

WT Y639F Y639F Y639F WT WT WT WT Y639F Y639F Y639F Y639F WT

Kazantsev and Pace (1998) Szewczak et al. (1998) Conrad et al. (1995) Schwans et al. (2003) Christian and Yarus (1992) Oyelere and Strobel (2000) Oyelere and Strobel (2000) Oyelere and Strobel (2000) Szewczak et al. (1999) Ryder and Strobel (1999) Conrad et al. (1995) Schwans et al. (2003) Christian and Yarus (1992)

F

AaS

AaS

References

(continued)

14


Table 1.1 (continued) Analog dTPaS (Sp isomer only) 5

m UaS CaS dUaS OMe UaS F UaS SH UaS

[dTPaS] (mM)

[Parent Polymerase NTP] (WT/ (mM) Y639F)

References

0.50 0.05 0.25 2.0 0.25 0.2

1.0 1.0 1.0 0.1 1.0 1.0

Unpublished observation Unpublished observation Szewczak et al. (1998) Conrad et al. (1995) Szewczak et al. (1998) Schwans et al. (2003)

WT WT Y639F Y639F Y639F Y639F

intense enough to easily quantitate, but minimizes the possibility of cooperative interferences due to multiple substitutions within the same RNA. Incorporation above this level also increases the number of RNAs cleaved at more than one location by iodine, leading to an overrepresentation of short cleavage products in sequencing gels. The approximate conditions used to produce this level of incorporation for several analogs is provided in Table 1.1, but incorporation levels can be sequence dependent and may need to be reoptimized for a particular transcript. When incorporating new analogs the proper conditions are determined by carrying out multiple transcriptions with a range of analog concentrations, 50 -end labeling the transcripts, and comparing the intensity of the iodine cleavage pattern to the one obtained with the parent analog (see below). Run-off transcriptions from DNA templates are carried out following standard procedures (Milligan and Uhlenbeck, 1989; Paschal et al., 2008). If plasmid DNA is used the template is first linearized with the appropriate restriction enzyme. When designing these plasmids 30 -overhangs at the cut site should be avoided as these are known to lead to poor transcription (Milligan and Uhlenbeck, 1989). In many cases, the restriction enzyme can be heat-denatured and the cut plasmid used without further purification. Alternatively the reaction is PCA extracted, ethanol precipitated, and resuspended. It is essential that the DNA template is completely digested as even a small amount of undigested template drastically reduces the desired transcript. For short transcripts the DNA template can also be quickly assembled by PCR amplification of synthesized oligonucleotides. A typical transcription contains 1 transcription buffer, 10–20 mM MgCl2 (optimized for transcript), 0.05 mg/ml linearized DNA template, 0.001 U/ml inorganic pyrophosphatase, 5 U/ml T7 RNA polymerase, 1.0 mM NTPs, and analog concentrations according to Table 1.1. The reactions are incubated for 2 h at 37 C. If a self-cleaving ribozyme has been incorporated into the transcript for uniform end formation the transcription reaction is heated to 65 C for 2 min and slow cooled to room temperature. The transcripts are purified by denaturing PAGE, visualized by UV shadowing, excised


15

from the gel, and eluted by rocking in elution buffer for 1 h at room temperature. The filtered elution is PCA extracted, ethanol precipitated and redissolved in a minimal volume of TE. RNA concentrations are determined by UV absorption at 260 nm.

3.3. Incorporation efficiency controls Before performing NAIM selections the level of analog incorporation at each site of the transcript must be determined. For new analogs these controls identify the concentration of analog required for an incorporation level of approximately 5% and ensures that the analog is incorporated specifically at coded positions. Incorporation controls also quantitate the level of incorporation at each site, which has been shown to be sequence specific, but reproducible for a given sequence. This sequence-dependent incorporation must be accounted for when analyzing sites of interference (see below). NAIM selections should be carried out using the same RNAs as the incorporation controls, so a sufficient quantity of RNA for the controls and activity selections should be transcribed once the incorporation efficiency is determined. Incorporation levels are determined by 50 -end labeling analogincorporated RNAs, cleaving with iodine and resolving the cleavage products by gel electrophoresis. The efficiency and distribution of analog incorporation is determined by comparing band intensities from the parental nucleotide and nucleotide analog cleavage patterns. For this reason incorporation controls for all analogs and the parent nucleotide should be included on a single gel. Transcripts retaining a 50 -terminal phosphate are first treated with calf intestinal alkaline phosphatase (CIP). Ten picomoles of each RNA are treated 50 units of CIP for 1 h at 37 C. CIP is removed either by PCA extraction or more conveniently by the use of commercially available silica membrane spin columns (i.e., QIAquickÒ Nucleotide Removal Kit). The phosphatase step can be eliminated if a self-cleaving ribozyme is inserted in the leading sequence or if the transcription is initiated with guanosine. RNAs are 50 -end labeled with 20 units of T4 polynucleotide kinase (PNK) and 150 mCi of [g-32P]ATP by incubating at 37 C for 1 h. Labeled RNAs are purified by denaturing PAGE, visualized by autoradiography, excised from the gel, and eluted by rocking in elution buffer for 1 h at room temperature. Short elution times in buffers containing SDS minimize the level of RNA degradation. RNAs are PCA extracted, ethanol precipitated and redissolved in a minimal volume of TE. To normalize the loading of sequencing gels a small aliquot of each RNA solution is scintillation counted. Each labeled RNA is then combined with TE and 2 loading buffer to provide a 10 ml solution that contains 200,000 cpm, which is split into two 5 ml aliquots. One aliquot is treated with 0.5 ml of 100 mM I2/ethanol, while the other is treated with 0.5 ml of ethanol alone to control for nonspecific degradation. Because the iodine

16


cleavage reaction has an endpoint of approximately 15% at least 100,000 cpm should be loaded in each lane to accurately quantitate band intensities. To improve resolution in the sequencing gel samples are heated to 90 C for 1 min and cooled on ice before loading. Gels are prerun for at least 2 h, loaded with iodine treated and untreated samples for the parent nucleotide and all analogs, and run for various lengths of time to maximally resolve different portions of the sequence. A phosphor storage screen is exposed to the dried gel overnight and imaged. Individual band intensities are quantitated by area integration of line profiles drawn through each lane (see below). The quality of the cleavage ladder is typically poorest for very short fragments and those with lengths closest to the uncleaved RNA. For difficult regions, the RNA can alternatively be labeled at its 30 -end with [32P]pCp and T4 RNA ligase or additional noninteracting sequence can be added to the transcript at the 50 - or 30 -end (Kwon and Strobel, 2008).

3.4. Activity selections NAIM studies require an assay to separate active and inactive RNAs from a pool of transcripts containing analog substitutions at random locations. A wide variety of selection strategies appropriate for NAIM studies have been developed, and detailed protocols for several of these methods have been published elsewhere (Cochrane and Strobel, 2004; Waldsich, 2008). In general, two broad categories of selections have been used. The first is to physically separate the active and inactive fractions, where the assay selects for a change in the RNAs’ length, fold, or interaction with a binding partner. Separations are carried out with denaturing PAGE, native gel mobility shift assay, column chromatography, or filter binding. This approach has been used to investigate functional groups required for ribozyme activity, RNA folding, metabolite binding to riboswitches, and protein–RNA or RNA–RNA interactions (Cochrane et al., 2003; Kwon and Strobel, 2008; Oyelere et al., 2002; Waldsich and Pyle, 2007). In these studies, the analog-substituted RNAs are radiolabeled at their 30 - or 50 -end either before or after the selection, and band intensities can often be quantitated for both the active and inactive fractions. The second class of selections relies on the specific radiolabeling of the active fraction. This approach has been used to study ribozyme-mediated ligation reactions where a radiolabeled substrate is transferred to active members of the RNA pool ( Jones and Strobel, 2003; Ryder and Strobel, 1999; Strobel et al., 1998). A major advantage of these types of selections is that no separation step is required since only the active members become radiolabeled and contribute to the iodine cleavage pattern. Regardless of the type of selection employed the assay should be as stringent as possible. For equilibrium assays involving RNA binding or folding, the reaction conditions are typically set such that the active fraction is the minor

17


component. For kinetic assays, stringent selections are produced by quenching the reactions as early in the time course as possible while still obtaining enough radiolabeled product for high-quality sequencing gels. An example of a ribozyme-mediated ligation selection is provided below (Fig. 1.4). In this study, fluorine-substituted adenosine analogs were used to identify sites of functional ionization in a self-ligating form of the Varkud Satellite (VS) ribozyme (Suydam and Strobel, 2008). These analogs have N1 pKa values spanning more than 4 pKa units as measured for the free nucleosides; 7-deazaadenosine (7dA, pKa 5.2), 7-fluoro-7-deazaadenosine 630

A

C U G C I G G G A A G 620 C G U C 610 U UCGCAA

600 VII 5⬘ * UAGUAGAGUG 3⬘ UCAUUUCAU AGCGUU U G A 780 790

IV

V

G 680 690 U C U C A UGAAAUUGU CGUAGCAGU G A GC U UAUUGUCA UC C G ACUUUAAC G U C G 710 670 C 700 A C U G U U A III C G C G G 640 A UA C G C 660 G 730 U A 720 G AGU 650 VI 740 A A U A A II AC G AA G GC GGGA GCU GUG CGGUAUUGGC U CG CCCGAACACGA CAC GUUAUGACUG AA GC 770 760 A 750

756 B

C Ligation reaction selected RNAs

2FA

2F7dA

7F7dA

7dA

A

2FA

+I2 2F7dA

7F7dA

A

7dA

–I2

2FA

2F7dA

7F7dA

A

7dA

2FA

2F7dA

7F7dA

A

7dA

5⬘-end-labeled unselected RNAs –I2 +I2

779

756 743

779

*

756 743

Figure 1.4 (A) Secondary structure of the Varkud Satellite ribozyme construct used for NAIM studies of ribozyme-catalyzed ligation. The site of ligation is marked with an arrow. (B) Sequencing gel of unselected RNAs labeled at their 50 -end and treated with or without iodine. (C) Sequencing gel of ligation selected RNAs treated with or without iodine. The site of pKa-dependent interference is denoted with an asterisk. Additional gels resolved all sites of incorporation from A645 to A779.

18


(7F7dA, pKa 4.5), adenosine (A, pKa 3.5), 2-fluoro-7-deazaadenosine (2F7dA, pKa < 1), and 2-fluoroadenosine (2FA, pKa < 1). The activity selection in this system is the ligation of a 50 -end labeled substrate (NT 599–620) onto the ribozyme sequence (NT 621–791) ( Jones and Strobel, 2003). An RNA sequence including a Hammerhead ribozyme at the 50 -end of the desired VS sequence is transcribed in the presence of analog nucleoside triphosphates according to Table 1.1. Hammerhead processing during transcription produces analog-substituted ribozyme RNA with a free 50 -hydroxyl at A621. A portion of these transcripts are labeled directly with PNK, purified by denaturing PAGE, iodine cleaved and loaded onto 8% denaturing sequencing gels for unselected incorporation efficiency controls (Fig. 1.4B). Prior to the reaction, unlabeled ribozyme RNAs are incubated in reaction buffer at 50 C for 15 min and cooled slowly to room temperature. Radiolabeled substrate RNA is produced with the appropriate 20 ,30 cyclic phosphate as described previously ( Jones et al., 2001). Ligation reactions containing 0.4 mM ribozyme and 0.05 mM substrate are initiated by the addition of 50 -end labeled substrate (>1,000,000 cpm). Reactions are allowed to proceed at room temperature until the fraction ligated reaches 15%, then quenched with 1 volume of 2 loading buffer. The quenched reactions are iodine cleaved and loaded onto 8% denaturing sequencing gels as described for the unselected incorporation efficiency controls (Fig. 1.4C). Quantitating individual band intensities for these gels as described below identified a single adenosine, A756, whose interference pattern is consistent with function ionization (Suydam and Strobel, 2008).

3.5. Quantitating sites of interference The analysis of NAIM data requires the inclusion of all relevant samples on a single gel. For every analog tested in a NAIM experiment, the relevant gel must also contain samples in which the parent nucleotide has been tested in a parallel experiment. Unselected controls can be run separately from the activity selected RNAs, but these gels must also contain lanes for all analogs and the parent nucleotide. Both types of gels also include lanes for parent and analog-incorporated RNAs that have not been iodine treated (Fig. 1.4B and C). These lanes serve as a control for nonspecific degradation of the RNA. Sites of nonspecific degradation often include unpaired sections of sequence as well as degradation products with lengths close to that of the uncleaved RNA. Positions of strong degradation are uninformative in NAIM experiments so every effort should be made to keep solutions RNAse free, to purify RNAs quickly, and to run sequencing gels soon after labeling reactions. Sites of interference correspond to individual bands that are missing or underrepresented in the activity selected sequencing gels relative to the


19

unselected controls. In some systems enhancements are also observed where bands are overrepresented in the activity selected lanes. The strongest interferences are easily identified as gaps in the activity selected sequencing ladder (Fig. 1.4C). To quantitate the level of interference at each site the peak intensities at every position of incorporation for both the parental nucleotide (NaS) and the nucleotide analog (daS) is calculated in both the activity selected and unselected sequencing gels. Line profiles are drawn through the appropriate lanes and peak areas are calculated by area integration. It is important to draw equivalent lines through each lane and to carefully define a consistent baseline for each before calculating the areas. Automated software has recently become available that may facilitate this process (Laederach et al., 2008). The quality of the line profiles and subsequent integration depends on the overall intensity of the cleavage ladder, the even migration of the lanes and the resolution of the individual bands, so gels should be run to optimize these features. The extent of interference at each position is calculated by substituting the individual band intensities into Eq. (1.1): k ¼ NF

ðNaS selected=daS selectedÞ : ðNaS unselected=daS unselectedÞ

ð1:1Þ

The normalization factor NF is introduced to account for uneven loading in the four lanes used to calculate the interference values. It is obtained by calculating the average of all interference values within two standard deviations of the mean prior to normalization, and is typically between 0.8 and 1.2. Interference values calculated with Eq. (1.1) normalize both for the extent of analog incorporation at a particular site and for the effect of the phosphorothioate substitution at that site. Sites of very weak incorporation or strong phosphorothioate interference cannot be quantitated in this way. Quantitation of interference values at over 100 sites in the group I intron provided an early experimental estimate for the noise in NAIM experiments (Ortoleva-Donnelly et al., 1998b). A conservative interpretation of NAIM data defines sites of interference as those with k greater than 2.0, sites of enhancements as those with k less than 0.5, and sites with intervening values as showing no effect. Due to the difficulty in quantitating very weak bands in the background of minor degradation k-values greater than 6.0 should be considered equivalent.

4. Properties of Analogs Almost 40 nucleotide analogs have been utilized in NAIM to date (Figs. 1.5–1.8). While these analogs represent a great diversity of functional group modifications, there remains a wide variety of nucleoside analogs

20


7

−O

g

b

a

O

O

O

N

8 N

P O P O P O O−

O−

NH2 6 N1 N

2

3

O

S−

OH

2⬘

OH

AaS

H3C

NH2 N N

TPaS −O

N

N NH2

N

TPaS

N NH2

N

TPa S

OH

OH

DAPa S

N

−O

O

O OH

N

−O

OH

OH

m6AaS NH2

TPa S −O

N

N

N N

OH

N

OH

OH

OH

OH

TPa S −O

NH2

N

TPaS −O

OH

OH

F

OH

OH

N N

TPa S −O

NH2 N

N N

OH

OH

O OH O OMeAaS

CH3

OH

OH

NH2

N

dAaS NH2

NH2

N

N N

N

TPaS −O

N N

O

n8AaS

FormAaS

N

TPa S −O

O

N

OH

NH2

O

F

2FAaS

N

N

N N

O

NH2

N

N

TPa S −O

2F7dAaS

N

TPaS −O

N

O

H N

OH

NH2 N

N

O

7F7dAaS

OH

7dAa S

N N

N

O

NH2

N

N

TPa S −O

c3AaS

F

TPa S −O

N

O

Pura S

OH

NH2

N

TPa S −O

O

N N

O OH

2APa S

N

NH

N

N

N N

N

TPa S −O

O

O

OH F FAaS

SHAaS

OH

N N

SH

Figure 1.5 Adenosine analogs. The chemical modification of each analog is highlighted with a box.

currently untested in NAIM studies. Many of these nucleosides are commercially available or have been synthesized in sufficient quantities for the preparation of 50 -O-(1-thio)nucleoside triphosphates. The application of new analogs will continue to expand the types of RNA interactions

21


7

-

g

b

a

N

O

O

O

N

O P O P O P O O-

O-

S-

O

6 NH N 2 NH2

O

2⬘ OH

OH

GaS O N N TPa S

O

NH

N

N

TPaS O

O OH

OH

OH

NH CH3

N TPa S O

OH

OH

N O

N

OH

O

O

NH2

N TPa S O

O

O OH

OH

7dGa S

dGa S

NH2

S6GaS

N

NH

NH N

O

m2Ga S O

OH

N

NH N

O

IaS

TPaS

S

O N

N

NH N

NH2

N TPaS O

NH N

NH2

O OH

O

OMeGaS

CH3

O N N TPaS

O

NH N

NH2

O OH

SH

SHGaS

Figure 1.6 Guanosine analogs. The chemical modification of each analog is highlighted with a box.

revealed by the NAIM method, many of which are described below. Nucleotide analogs can be approximately classified into three groups: those deleting a functional group, those adding steric bulk to the functional group, and those altering the chemical nature of the functional group. Many analogs clearly modify more than one property of the nucleotide, so whenever possible a series of closely related analogs should be employed to dissect these effects. Nucleotide analog interference suppression (NAIS) has also been used to refine the role of functional groups at specific sites of interference (described below).

4.1. Ribose modifications Four types of ribose modifications have been utilized in NAIM, each altering the 20 -position. These include 20 -deoxy modifications (dAaS, dGaS, dCaS, dUaS), 20 -O-methyl modifications (OMeAaS, OMeGaS,

22


NH2

4

g

b

a

O

O

O

-O

5 6

N N

O

P O P O P O O-

O-

S-

O OH

2⬘

OH

CaS NH2

NH2 F

N N TPaS

N

NH2 N

N

O

N TPa S O

O

OH

OH

O OH

N

TPaS O

N

N

O

TPa S O

N

O

TPaS O

O

O OH

OH

O O

OMeCaS

dCa S

OH

ZaS

NH2

NH2

O

OH

yiCaS

N

O

TPa S O

OH

OH

f 5CaS

NH2

N

N

O

O

n6Ca S

N

O TPaS O

O OH

NH

O

OH

CH3

SH

SHCaS

Figure 1.7 Cytosine analogs. The chemical modification of each analog is highlighted with a box.

O

-O

g

b

a

O

O

O

5

NH N

O

P O P O P O O-

O-

S-

O

2⬘ OH OH

Ua S O H3C

O HN

NH N

O

TPaS O

O NH O

TPa S O O OH

OH

OH

OH

OH

dUa S

NH TPaS O

N

O

O OH O

OMeUaS

CH3

O

O

N

NH

O TPa S O

O

yaS

m5UaS

N TPa S O

O

O NH

NH

O

N

O

TPaS O O OH

O F

FUaS

OH

SH

SHUaS

Figure 1.8 Uridine analogs. The chemical modification of each analog is highlighted with a box.


23

20 -deoxy-20 -fluoro modifications (FAaS, FUaS), and 2 -deoxy-2 -thio modifications (SHAaS, SHGaS, SHCaS, SHUaS). Each of these analogs is more readily incorporated by the Y639F point mutant of T7 RNA polymerase and may require increased analog concentration, decreased NTP concentration, or both for even incorporation (Table 1.1). These analogs probe the contribution of individual 20 -hydroxyls within an RNA (Conrad et al., 1995; Gaur and Krupp, 1993; Ortoleva-Donnelly et al., 1998b; Schwans et al., 2003; Szewczak et al., 1998). Interferences observed with dNaS substitutions identify those 20 -OH groups required for function, while the interference patterns observed with OMeNaS and FNaS analogs help define the functional role of these 20 -OH groups. Although 20 -O-methyl and 20 -deoxy-20 -fluoro analogs cannot act as hydrogen bond donors they have been shown to reproduce the hydrogen bond acceptor properties of 20 -hydroxyls (Ortoleva-Donnelly et al., 1998b). For this reason a site that shows interference with dNaS, OMeNaS, and FNaS suggests a 20 -OH that acts as a hydrogen bond donor, while a site that shows interference with dNaS alone suggests the 20 -OH acts as a hydrogen bond acceptor. Occasionally, FNaS interference is observed at sites that do not exhibit dNaS interference (Ortoleva-Donnelly et al., 1998b). This interference pattern has been attributed to sites that require a 20 -C-endo ribose conformation, a sugar pucker that is destabilized by the electron withdrawing properties of the 20 -fluoro substituent in favor of the 30 -C-endo conformation (Uesugi et al., 1979). Sites that exhibited this interference pattern in the group I intron map to ribose sugars refined as having 20 -C-endo conformations in subsequent crystal structures (Adams et al., 2004; Cate et al., 1996). Sites of OMeNaS interference that do not interfere with dNaS suggest tight packing in the minor groove that cannot accommodate the 20 -O-methyl substitution. These sites can be further investigated with SHNaS analogs which have been shown to cause interference at densely packed regions of RNA (Schwans et al., 2003). In principle, SHNaS analogs should also produce interference at sites of Mg2þ coordination that could be rescued by the addition of thiophilic metals. OMeCaS, OMeUaS),

0

0

4.2. Base modifications 4.2.1. Adenosine analogs The largest collection of nucleobase modifications developed for NAIM studies are those of adenosine (Fig. 1.5). The focus on adenosine analogs stems in part from the observation that adenosines are present at a disproportionately high frequency in several RNA structural motifs (Gutell et al., 1985). Nucleobase-modified adenosine analogs include diaminopurine riboside (DAPaS), 2-aminopurine riboside (2APaS), N-methyladenosine (m6AaS), purine riboside (PuraS), 3-deazaadenosine (c3AaS),

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7-deazaadenosine (7dAaS), 7-fluoro-7-deazaadenosine (7F7dAaS), 2-fluoro-7-deazaadenosine (2F7dAaS), 2-fluoroadenosine (2FAaS), formicin (FormaS), and 8-azaadenosine (n8AaS). Even incorporation at low analog concentrations is observed for each analog except PuraS, 2APaS, and c3AaS, which are incorporated with lower efficiency even at elevated concentrations (Table 1.1). Each of the nucleobase analogs provides specific information on the role of adenosine functional groups in RNA activity. PuraS, 2APaS, and m6AaS measure the effect of modifications at the N-6 exocyclic amine (OrtolevaDonnelly et al., 1998b). PuraS and 2APaS remove the amine, leading to interference at any site where this functional group is required for activity. m6AaS replaces one proton of the amine with a methyl group. Interference with this analog indicates either that both N6 protons are required for function or that there is insufficient space in the local structure to accommodate the additional methyl group. Interference caused by the additional C2 amine of 2APaS can also arise from close packing in the minor groove. These sites can be identified by a similar interference with DAPaS or in very close contacts with 2FAaS (Ortoleva-Donnelly et al., 1998b; Suydam and Strobel, 2008). 7dAaS replaces the N7 nitrogen with a C—H group. Interference with this analog indicates an important major groove contact with the N7 nitrogen (Kaye et al., 2002; Ortoleva-Donnelly et al., 1998b; Waldsich and Pyle, 2007). These sites will provide a similar result with 7F7dAaS and 2F7dAaS (Suydam and Strobel, 2008). The N7 nitrogen of adenosine also participates in metal binding in some systems and NAIM analysis has been used to identify these sites (Wrzesinski and Jozwiakowski, 2008). Interference with PuraS, m6AaS, and 7dAaS is a strong indicator of Hoogsteen hydrogen bonding (Ortoleva-Donnelly et al., 1998b; Waldsich and Pyle, 2007). c3AaS replaces the N3 nitrogen with a C—H group and provides a similar test for interactions with this nitrogen. Interference with c3AaS has identified a number of A-minor tertiary interactions involving the N3 nitrogen (Soukup et al., 2002). In addition to structural interactions, NAIM has been used to identify sites of functional adenosine ionization. Analogs appropriate for these studies exhibit large N1 pKa shifts with minimal structural perturbation and include FormAaS, 7dAaS, 7F7dAaS, n8AaS, 2F7dAaS, and 2FAaS ( Jones and Strobel, 2003; Ryder et al., 2001; Suydam and Strobel, 2008). 4.2.2. Guanosine analogs Four nucleobase-modified analogs of guanosine have been used in NAIM studies, including inosine (IaS), N2-methylguanosine (m2GaS), 6-thioguanosine (S6GaS), and 7-deazaguanosine (7dGaS) (Fig. 1.6). IaS and 7dGaS are both incorporated efficiently and evenly by wild-type polymerase, whereas m2GaS requires the Y639F point mutant of T7 RNA polymerase and decreased GTP concentrations (Table 1.1). S6GaS is incorporated by


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the wild-type polymerase but requires the addition of 4 mM Mn2þ in the transcription buffer. RNAs containing S6GaS are unusually unstable and should be used within 48 h of transcription (Basu et al., 1998). Interferences arising from IaS and m2GaS have been used extensively to distinguish between tertiary contacts in RNA (Heide et al., 1999; Ortoleva-Donnelly et al., 1998a; Strobel and Shetty, 1997; Szewczak et al., 1998). IaS lacks the N2 exocyclic amine of guanosine and interference with this analog results from reduced duplex stability or loss of a tertiary contact to the N2 amine. In contrast, m2GaS introduces a methyl group to the exocyclic amine and is isoenergetic with G in the context of G–C, GU, and GA base pairs (Rife et al., 1998). Interference with m2GaS has only been observed at sites of tertiary contact (Ortoleva-Donnelly et al., 1998a). 7dGaS like 7dAaS replaces the N7 nitrogen with a C—H group and produces interference at sites with important major groove contacts to this nitrogen as well as at sites of metal binding (Heide et al., 2001; Kazantsev and Pace, 1998; Kwon and Strobel, 2008; Wrzesinski and Jozwiakowski, 2008). S6GaS replaces the O6 keto group of guanosine with sulfur and has been used to identify sites of monovalent metal binding (Basu et al., 1998). 4.2.3. Pyridine analogs The development of pyrimidine base-modified analogs has been much more limited than for the purines. Four cytidine analogs have been synthesized for use in NAIM studies, including 6-azacytidine (n6CaS), 5-fluorocytidine (f 5CaS), pseudoisocytidine (CiCaS), and zebularine (ZaS) (Fig. 1.7). Each of these analogs has altered N3 pKa values and have been used to identify sites of functional cytidine ionization in catalytic RNAs (Oyelere and Strobel, 2000; Oyelere et al., 2002). Two nucleobasemodified uridine analogs have also been used NAIM studies, including 5-methyluridine (m5UaS) and pseudouridine (CaS), both of which modify the 5-position in the major groove (Fig. 1.8). Nucleobase analogs of pyrimidines are generally incorporated evenly with wild-type polymerase (Table 1.1).

5. Nucleotide Analog Interference Suppression Some of the interferences observed in traditional NAIM experiments arise from the deletion of a critical hydrogen bond and the subsequent destabilization of RNA folds, decrease in population of catalytically active conformations, or loss of affinity for small molecule ligands or proteins. Although NAIM identifies the RNA functional groups involved in these hydrogen bonds it does not identify the hydrogen bonding partners. NAIS is an extension of NAIM experiments that maps tertiary contacts by

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identifying pairs of interacting functional groups required for function (Boudvillain and Pyle, 1998; Fedorova and Pyle, 2005; Jansen et al., 2006; Strobel and Ortoleva-Donnelly, 1999; Strobel et al., 1998; Szewczak et al., 1998, 1999). The approach combines a single mutation or functional group substitution at a specific site with random incorporation of phosphorothioate-tagged nucleotides throughout the remainder of the RNA. These chimeric RNA molecules are typically constructed by joining or annealing synthetic oligonucleotides carrying the site-specific deletion with the pool of transcribed RNAs carrying the randomly incorporated analogs. The resulting RNAs are then subjected to the same type of selection as the original NAIM study. NAIS experiments rely on the expectation that the deletion of one functional group in a hydrogen bonding pair will eliminate the positive contribution made by the second, such that it can be deleted without an additional energetic penalty (Strobel and Ortoleva-Donnelly, 1999). In this way, the site-specific deletion of one functional group should lead to a suppression in interference caused by analog substitution at the interacting site, while leaving the remaining sites of interference unchanged. This pattern of interference suppression has been observed in several systems and the resulting predictions of tertiary contacts have agreed extremely well with subsequent crystal structures (Adams et al., 2004; Toor et al., 2008). The key to NAIS experiments is the ability to reconstruct the selection assay such that there is activity in the context of the site-specific substitution. This is often accomplished by modifying buffer conditions, lengthening reaction times, or increasing the concentration of substrates. Several additional controls are required to demonstrate that the suppression of interference is specific both to the nature of the site-specific substitution and to the site of suppression. First, site-specific substitutions with other functionalities or at other sites should not produce the same suppression pattern. Second, the observed suppression should be specific to a single residue among additional sites of interference. Finally, the observed suppression should be specific to a single or closely related set of analogs.

6. Conclusions NAIM is a generalizable chemogenetic method that rapidly identifies the chemical groups important for RNA function. In principle, NAIM can be extended to any nucleotide analog that can be incorporated by in vitro transcription to study any RNA with a selectable function. The robustness of the method has been demonstrated by the excellent agreement between structural contacts predicted by NAIM and atomic resolution structures. Because a single NAIM experiment can assay several functional group


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modifications at every site in parallel the method provides an efficient route to define key features of RNA structure and function. As the number of functional RNAs identified continues to expand, NAIM should provide a straightforward method to determine which individual nucleotides or regions of sequence warrant additional study.

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Ruffner, D. E., and Uhlenbeck, O. C. (1990). Thiophosphate interference experiments locate phosphates important for the hammerhead RNA self-cleavage reaction. Nucleic Acids Res. 18, 6025–6029. Ryder, S. P., and Strobel, S. A. (1999). Nucleotide analog interference mapping of the hairpin ribozyme: Implications for secondary and tertiary structure formation. J. Mol. Biol. 291, 295–311. Ryder, S. P., and Strobel, S. A. (2002). Comparative analysis of hairpin ribozyme structures and interference data. Nucleic Acids Res. 30, 1287–1291. Ryder, S. P., Ortoleva-Donnelly, L., Kosek, A. B., and Strobel, S. A. (2000). Chemical probing of RNA by nucleotide analog interference mapping. Methods Enzymol. 317, 92–109. Ryder, S. P., Oyelere, A. K., Padilla, J. L., Klostermeier, D., Millar, D. P., and Strobel, S. A. (2001). Investigation of adenosine base ionization in the hairpin ribozyme by nucleotide analog interference mapping. RNA 7, 1454–1463. Schwans, J. P., Cortez, C. N., Olvera, J. M., and Piccirilli, J. A. (2003). 20 -mercaptonucleotide interference reveals regions of close packing within folded RNA molecules. J. Am. Chem. Soc. 125, 10012–10018. Sood, V. D., Yekta, S., and Collins, R. A. (2002). The contribution of 20 -hydroxyls to the cleavage activity of the Neurospora VS ribozyme. Nucleic Acids Res. 30, 1132–1138. Soukup, J. K., Minakawa, N., Matsuda, A., and Strobel, S. A. (2002). Identification of A-minor tertiary interactions within a bacterial group I intron active site by 3-deazaadenosine interference mapping. Biochemistry 41, 10426–10438. Sousa, R., and Padilla, R. (1995). Mutant T7 RNA-polymerase as a DNA-polymerase. EMBO J. 14, 4609–4621. Strobel, S. A. (1999). A chemogenetic approach to RNA function/structure analysis. Curr. Opin. Struct. Biol. 9, 346–352. Strobel, S. A., and Ortoleva-Donnelly, L. (1999). A hydrogen-bonding triad stabilizes the chemical transition state of a group I ribozyme. Chem. Biol. 6, 153–165. Strobel, S. A., and Shetty, K. (1997). Defining the chemical groups essential for Tetrahymena group I intron function by nucleotide analog interference mapping. Proc. Natl. Acad. Sci. USA 94, 2903–2908. Strobel, S. A., Ortoleva-Donnelly, L., Ryder, S. P., Cate, J. H., and Moncoeur, E. (1998). Complementary sets of noncanonical base pairs mediate RNA helix packing in the group intron active site. Nat. Struct. Biol. 5, 60–66. Suydam, I. T., and Strobel, S. A. (2008). Fluorine substituted adenosines as probes of nucleobase protonation in functional RNAs. J. Am. Chem. Soc. 130, 13639–13648. Szewczak, A. A., Ortoleva-Donnelly, L., Ryder, S. P., Moncoeur, E., and Strobel, S. A. (1998). A minor groove RNA triple helix within the catalytic core of a group I intron. Nat. Struct. Biol. 5, 1037–1042. Szewczak, A. A., Ortoleva-Donnelly, L., Zivarts, M. V., Oyelere, A. K., Kazantsev, A. V., and Strobel, S. A. (1999). An important base triple anchors the substrate helix recognition surface within the Tetrahymena ribozyme active site. Proc. Natl. Acad. Sci. USA 96, 11183–11188. Szewczak, L. B. W., DeGregorio, S. J., Strobel, S. A., and Steitz, J. A. (2002). Exclusive interaction of the 15.5 kD protein with the terminal box C/D motif of a methylation guide snoRNP. Chem. Biol. 9, 1095–1107. Toor, N., Keating, K. S., Taylor, S. D., and Pyle, A. M. (2008). Crystal structure of a self-spliced group II intron. Science 320, 77–82. Uesugi, S., Miki, H., Ikehara, M., Iwahashi, H., and Kyogoku, Y. (1979). Linear relationship between electronegativity of 20 -substituents and conformation of adenine nucleosides. Tetrahedron Lett. 4073–4076.

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C H A P T E R

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Hydroxyl-Radical Footprinting to Probe Equilibrium Changes in RNA Tertiary Structure Inna Shcherbakova*,† and Somdeb Mitra* Contents 1. Introduction 2. Sample Preparation 3. Equilibrium OH Footprinting Based on Peroxidative Fenton Chemistry 4. Equilibrium OH Footprinting Based on Oxidative Fenton Chemistry 5. Cleavage Product Separation 6. Quantitation of the Changes in the Reactivity and Data Analysis 7. Conclusions Acknowledgments References

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Abstract Hydroxyl-radical footprinting utilizes the ability of a highly reactive species to nonspecifically cleave the solvent accessible regions of a nucleic acid backbone. Thus, changes in a nucleic acids structure can be probed either as a function of time or of a reagent’s concentration. When combined with techniques that allow single nucleotide resolution of the resulting fragments, footprinting experiments provide richly detailed information about local changes in tertiary structure of a nucleic acid accompanying its folding or ligand binding. In this chapter, we present two protocols of equilibrium hydroxyl-radical footprinting based on peroxidative and oxidative Fenton chemistry and discuss how to adjust the Fenton reagent concentrations for a specific experimental condition. We also discuss the choice of the techniques to separate the reaction products and specifics of the data analysis for equilibrium footprinting experiments. Protocols addressing the use of peroxidative Fenton chemistry for time-resolved

* Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York, USA Current address: Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts, USA

{

Methods in Enzymology, Volume 468 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)68002-2

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32

Inna Shcherbakova and Somdeb Mitra

studies have been published [Schlatterer and Brenowitz, 2009. Methods; Shcherbakova and Brenowitz, 2008. Nat. Protoc. 3(2), 288–302; Shcherbakova et al., 2006. Nucleic Acids Res. 34(6), e48; Shcherbakova et al., 2007. Methods Cell Biol. 84, 589–615].

1. Introduction The term ‘‘footprinting’’ refers to an assay that probes changes in macromolecular structure by probing changes in the sensitivity of the individual residues to a modification or cleavage reagent. This technique was initially developed to probe specific DNA–protein interactions by DNase I (Galas and Schmitz, 1978), and the term arose from an association that a protein leaves ‘‘a footprint’’ on its cognate binding site on a DNA, visualized as a blank region lacking bands of cleaved DNA on the image of a sequencing gel. Quantitating changes in the band intensities within such ‘‘footprinted’’ regions as a function of the protein ligand concentration or as a function of time provided the binding affinities or interaction rates, respectively. Later, it was demonstrated that the same approach could be used with a variety of reagents to probe not only intermolecular (Tullius and Dombroski, 1986) but also intramolecular interactions (Latham and Cech, 1989) and was appropriate to probe changes in the macromolecular structure associated with macromolecular folding or assembly. Among the wide variety of footprinting reagents used to probe RNA structure, small chemicals that do not confer significant steric hindrances proved to be most useful. One of the smallest one, the hydroxyl radical ( OH), has been used extensively to probe changes in the tertiary structure of RNA in kinetic (Adilakshmi et al., 2005, 2008; Chauhan et al., 2009; Deras et al., 2000; Heilman-Miller and Woodson, 2003; Kwok et al., 2006; Laederach et al., 2007b; Lease et al., 2007; Nguyenle et al., 2006; Sclavi et al., 1997, 1998; Shcherbakova and Brenowitz, 2005; Shcherbakova et al., 2004; Silverman et al., 2000; Uchida et al., 2003) and equilibrium studies (Ralston et al., 2000; Schlatterer et al., 2008; Su et al., 2003; Swisher et al., 2002; Takamoto et al., 2002, 2004; Uchida et al., 2002, 2003). Falling only slightly short in its reactivity to elemental fluorine, this highly reactive species is capable of oxidizing virtually every biological molecule. In the case of nucleic acids, hydroxyl radicals cleave their backbones by abstracting hydrogens from the ribose or deoxyribose sugar rings (Hertzberg and Dervan, 1984). Because the sugars are ubiquitously located on the outer faces of base pairs in nucleic acid helices, the changes in OH cleavage do not reflect changes in the RNA secondary structure but are associated with the formation of tertiary interactions. An additional advantage is that OH radicals interact nonspecifically with nucleotides, thereby allowing us to monitor individual behavior of all residues that are involved in tertiary interactions.

Hydroxyl-Radical Footprinting to Probe Equilibrium Changes in RNA Tertiary Structure

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When the products of the cleavage reaction are separated on a highresolution gel or by capillary electrophoresis, this footprinting technique tracks changes in the reactivity of every nucleotide along the RNA chain. The small size of OH and its properties comparable to that of a water molecule, allow us to link the changes in the OH reactivity of a nucleotide to changes in its solvent accessibility, associated with conformational changes in an RNA molecule. This footprinting technique is appropriate for both in vitro and in vivo studies. The feasibility to footprint RNA structure in vivo was demonstrated in synchrotron footprinting experiments on 16S rRNA and RNase P (Adilakshmi et al., 2006). There are a wide variety of ways to produce OH for footprinting. In the in vivo footprinting experiments mentioned above the OH are produced by radiolysis of intracellular water by a high-energy X-ray beam generated by a synchrotron. However, for in vitro studies, the readily available chemical reagent hydrogen peroxide (H2O2) can generate OH without the need for a synchrotron beam. A simple way to decompose H2O2 in a controlled manner is to use Fenton chemistry (Fenton, 1894) in its Udenfriend version (Udenfriend et al., 1954) where EDTA-chelated iron(II) reduces peroxide to produce OH and the resulting iron(III)–EDTA is recycled back to iron (II)–EDTA by ascorbate:

Fe(II) – EDTA + H2O2 → Fe(III) – EDTA + •OH + OH – ascorbate The concentrations of the reagents can be adjusted so that the cleavage occurs during several seconds. The reaction is quenched by addition of thiourea or ethanol. In this version this reaction can be used to probe equilibrium changes in the RNA structure, as well as to track slow kinetic processes taking minutes to hours. The protocols for millisecond time-resolved footprinting are published elsewhere (Schlatterer and Brenowitz, 2009; Shcherbakova and Brenowitz, 2008; Shcherbakova et al., 2006, 2007). It was recently noticed that minutes-long incubation of RNA with only Fe(II)–EDTA caused a small amount of RNA to be uniformly cleaved, presumably due to the production of the OH by oxidative chemistry. This reaction, simpler in its execution (Takamoto et al., 2004), requires longer reaction times. However, it can be a valuable tool in case of molecules or a macromolecular complex that are highly sensitive to hydrogen peroxide, as was reported for the TFIIIA–DNA complex (Tullius et al., 1987). In this chapter, we present two protocols of equilibrium OH footprinting based on the peroxidative and the oxidative Fenton chemistry that we routinely use to probe changes in RNA tertiary structure during its cation-dependent folding. We discuss how the reaction conditions can be adjusted to accommodate a specific experimental requisite. We also note the available choices of techniques to separate and quantitate the reaction products and specifics of the data analysis of equilibrium footprinting experiments.

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2. Sample Preparation Our laboratory has over a decade’s experience in studying the folding pathways of the group I intron ribozyme derived from Tetrahymena thermophila, and we shall use it as an example in the subsequent sections to highlight the applications of equilibrium OH footprinting. The methods described here are very general and can be easily adapted for any structured RNA molecule. The RNA molecule under investigation can be directly end-labeled with a radioactive label, in which case the OH cleavage products can be separated on a high-resolution denaturing polyacrylamide gel. The dried gel is exposed to a phosphor storage screen and visualized by scanning the screen in a phosphor-imaging system (STORMÒ phosphorimager from GE Healthcare in our case). The intensities of cleavage products, appearing as bands in the gel image, are quantitated using the software SAFA (Das et al., 2005; Laederach et al., 2007a; see also Chapter 3). Alternatively, to avoid radioactivity and increase the throughput of product separation and data analysis, we have recently adopted an indirect fluorescent labeling strategy, described in Section 5. For the experiments with directly labeled RNA, we first separately label the 50 - and 30 -ends of RNA (Shcherbakova and Brenowitz, 2008) with 32P. Several reaction aliquots of the RNA in a buffered solution are prepared so that each reaction tube contains enough radioactively labeled RNA for visualization and quantitation of the bands of the reaction products (usually 10–20 pmol RNA per tube). For investigating mono- or divalent cation mediated equilibrium folding of the RNA each reaction sample is heated to denature the RNA (90 C for 3 min in our case) and gradually cooled to the desired folding temperature when a desired concentration of the cation is added to the RNA and the sample is allowed to equilibrate. Incubation at 50 C is often a necessary intermediate step for large RNA molecules to avoid misfolding. Depending on the folding temperature and the type of the cations, equilibration can take from several minutes to hours. For example, in monovalent ions and/or at high temperatures, the Tetrahymena ribozyme folding has been shown to be completed within seconds. However, the same ribozyme can be trapped in a long lived misfolded state at lower temperatures or in high-divalent cation concentrations, so that reaching the catalytically active native state can take hours. The RNA for the footprinting experiments should be prepared with a special care to minimize any prereaction cleavage of its backbone which generates high background noise that appears as high-intensity bands in sequencing gels or abnormally high peaks in automated sequencer readouts. The buffers should be chosen wisely to minimize quenching of OH by buffer components and to prevent side reactions that might cause random


35

RNA cleavage. Inorganic buffers such as sodium cacodylate buffer are ideal due to their low propensity to quench OH. (Note. Sodium phosphate buffer was reported to significantly increase nucleic acid cleavage; Tullius et al., 1987.) Whereas monovalent cations do not interfere with the Fenton chemistry and hence does not affect the OH cleavage reaction, high concentrations of divalent cations (higher than 50 mM) can reduce the extent of OH cleavage by displacing iron (Fe2þ) from its complex with EDTA (Tullius et al., 1987). Glycerol is a very potent OH quencher (Tullius et al., 1987) that should be omitted if possible. If glycerol must be present in a reaction buffer, its scavenging must be accommodated by a concomitant upscaling of OH production by increasing the concentrations of the Fenton reagents. We discuss below how to troubleshoot such problems by scaling the OH production.

3. Equilibrium OH Footprinting Based on Peroxidative Fenton Chemistry The standard set of Fenton reagents we use for footprinting experiments include ammonium iron(II) sulfate hexahydrate [(NH4)2Fe (SO4)26H2O] (Sigma Ultra, minimum 99% pure); 0.5 M EDTA solution, pH 8.0 (Ambion); (þ)Sodium L-ascorbate (Sigma), 30% Hydrogen Peroxide (Sigma Aldrich), thiourea (Sigma), and RNase-free water (e.g., the DEPCtreated water from Ambion). A 100 mM stock solution of ferrous ammonium sulfate should be freshly prepared for every experiment or kept frozen in small aliquots. The color of this solution can change to slight yellowish due to the iron oxidation. Do not use the solution that turns brown or orange, with some precipitate. The Fe–EDTA solution is prepared by mixing the ferrous ammonium sulfate with EDTA to have 1.1 times excess of EDTA over iron. Solutions of sodium ascorbate and dilutions of hydrogen peroxide should be prepared fresh for each experiment. Following equilibration of the RNA sample with the desired concentration of the cation at the folding temperature 2 ml drops of 5 mM Fe–EDTA, 50 mM sodium ascorbate, and 1.5% hydrogen peroxide solutions are separately placed on the inside wall of the microfuge tube containing the RNA. The Fenton reaction is initiated by vigorous vortexing of the tube, resulting in simultaneous addition of all the three components to the 100 ml of the RNA solution. After 15 s the reaction is quenched by addition of 20 ml of stop solution (prepared beforehand by mixing thiourea with EDTA to final concentrations of 100 and 200 mM, respectively). If a protein component was present in the reaction mix it can be removed by phenol/ chloroform treatment of the samples with subsequent ethanol precipitation of nucleic acid from the aqueous phase.

36


Alternatively if only nucleic acids are present in solution, the reaction can be quenched by addition of three times the reaction volume of ethanol. Following addition of ethanol, the reaction products are kept for at least 30 min at 20 C and then precipitated by spinning in a refrigerated centrifuge at 14,000 rpm. Formation of tertiary structure can require molar concentrations of monovalent cations that upon ethanol precipitation form large pellets. In the case of high concentrations of monovalent cations, the reaction solution should be diluted to adjust the monovalent ion concentration to 0.15–0.3 M prior to precipitation. We found that the salt pellets are significantly smaller if the samples are first diluted and then precipitated by an equal volume of isopropanol instead of ethanol. In the case of titration samples that contain less than 50 mM of monovalent ions additional monovalent salts should be added to 0.15–0.3 M prior to the precipitation step to facilitate precipitation of small RNA fragments generated during OH cleavage. The precipitation of the small RNA fragments can be additionally facilitated by addition of glycogen solution (AmbionÒ ) to 1 mg/ml. Following precipitation, the pellets are dried in a SpeedVac and resuspended in a gel loading buffer containing formamide, EDTA, bromophenol blue, and xylene cyanol (we use Gel loading buffer II from AmbionÒ ). The samples are then loaded into the wells of a denaturing sequencing gel and separated at constant power (70–80 W). In certain cases, when the reactions are performed in smaller volumes, the formamide gel loading buffer may be added directly to the reaction mix after quenching OH production and a small aliquot (usually up to 10 ml) is loaded directly on the gel. However, in these situations the radioactive counts of the labeled RNA should be high enough to allow visualization of the cleavage products. We do not recommend direct sample loading for the cation titration experiments since the high amount of salt in some lanes will interfere with the electrophoretic separation of fragments, thereby significantly diminishing data quality. We have also noted improved signal to noise ratio as well as unbiased distribution of large and small cleavage fragments when the reaction products are completely precipitated, washed with 70% ethanol and resuspended in small volumes of gel loading buffer (usually 12 ml for loading 4 ml in each of the three sequencing gels) prior to separation by electrophoresis.

4. Equilibrium OH Footprinting Based on Oxidative Fenton Chemistry Radioactively labeled RNA samples are equilibrated in 100 ml buffered solution as described above. The following fresh solutions are prepared (or thawed from frozen stock aliquots): 250 mM (NH4)2Fe(SO4)2, 250 mM


37

EDTA, and 500 mM sodium ascorbate. Just before initiating the cleavage reaction, the Fenton reaction mix is prepared by mixing together 2 ml of the (NH4)2Fe(SO4)2 solution, 2.2 ml of the EDTA solution, 62.5 ml of the ascorbate solution, and 158 ml of RNase-free water. This Fenton reaction mix contains 0.1 mM Fe (NH4)2(SO4)2, 0.14 mM EDTA, and 6.94 mM ascorbate. OH cleavage is initiated by adding 5 ml of the Fenton reaction mix to the RNA solution and allowing the cleavage reaction to proceed for 30 min at a desired temperature (we have not noticed significant temperature dependence of OH production). The reaction is quenched by addition of three times the volume of ethanol or equal volume of isopropanol and precipitated as described for the peroxidative Fenton reaction. The concentrations of the reagents listed in both protocols are used to probe structural changes during equilibrium RNA folding under ideal nonradical scavenging condition, namely, in the presence of sodium or potassium cacodylate buffer and various concentrations of mono- or divalent cations. Usually a small concentration of EDTA (0.1 mM ) is present in the solution to bind contaminant transition metals, thereby preventing them from binding to the RNA and cleaving its backbone. We recommend minimizing the amount of OH scavengers (glycerol, DTT, b-mercaptoethanol and organic buffers) in the buffer. If their presence is unavoidable and/or a relatively high concentration of biological components is present, the concentrations of the Fenton reagents should be scaled appropriately. The optimal OH cleavage under nonoptimal scavenging condition can be found by a ‘‘dose response’’ experiment. For this experiment, the RNA sample is prepared as described above and aliquoted into several tubes containing the reaction mix in question. Then Fenton reaction is performed such that the concentration of any one of the Fenton reagents increases from one test tube to another. The products of the reaction are separated by denaturing gel electrophoresis, while taking care that the smallest cleavage fragments do not run out of the bottom of the gel, so that they can be visualized with the uncut RNA band (at the top) on the same gel. The fraction of cleaved RNA is calculated from the ratio of the intensity for the contour enclosing all the cleaved fragments, to the total intensity of cleaved and uncut samples (Shcherbakova and Brenowitz, 2008). Single-hit kinetics is typically achieved when no less than 70% of the RNA remains intact (Tullius et al., 1987). The product profile should be relatively uniform. When the intensity is visibly increased from longer to shorter fragments it means that the RNA is cleaved multiple times and the concentration of OH should be decreased to achieve the single-hit kinetics condition. The reagent concentrations determined by the dose response experiments are then used for equilibrium folding studies of the RNA under the desired experimental condition.

38


5. Cleavage Product Separation The cleavage products of the peroxidative and oxidative Fenton reaction are processed identically. Following ethanol/isopropanol precipitation as described above, the pellets are washed with 70% ethanol, dried in a SpeedVac, dissolved in a formamide gel loading buffer, heated for 30 s at 95 C and loaded onto high-resolution 8% acrylamide/bisacrylamide sequencing gel containing 7 M urea (Tijerina et al., 2007). More than a single gel may be required to clearly separate the regions of interest. For every gel it is important to run a ladder for assigning nucleotide position numbers to the bands on the gel. We typically use a G ladder generated by T1 ribonuclease which, in the presence of 7.5 M urea at 55 C, cleaves after every guanosine residue along the RNA backbone. We found that one-meter-long electrophoresis gel plate can save some time especially in the case of long RNA molecules. In this case, the special long-range gel solutions (e.g., LongRanger Gel solution from CambrexÒ ) are recommended. It is critical to siliconize one of the long gel plates. The long gels have to be cut in half before transferring them to a gel paper. Gels are then dried, exposed to a phosphor storage screen and visualized by scanning the screen by a phosphor imager. A typical equilibrium footprinting gel obtained for a Mg2þ titration of the Tetrahymena ribozyme bearing the UUCG mutation of the L5b tetraloop is shown in Fig. 2.1. The ‘‘no OH’’ lane demonstrates a single RNA band of a low electrophoretic mobility that corresponds to the full-length RNA without any detectable degradation. The same band of a diminished intensity is present in all other lanes on this gel along with additional bands corresponding to the shorter cleavage products separated at single nucleotide resolution. For the lanes corresponding to the RNA incubated in the absence of Mg2þ, the backbone cleavage is nearly uniform. The decrease in the intensity of cleaved bands, corresponding to certain nucleotides, correlate with the decreased solvent accessibility of those nucleotides due to tertiary interactions formed upon addition of higher concentrations of mono- or divalent cations. Some bands display a reverse trend, that is, increase in intensity with increasing ionic strength. Increased reactivity to OH can be generally rationalized as an increase in the solvent exposure of some regions in the RNA molecule upon its folding (e.g., the increase in the cleavage of the nucleotide 122 reports on the formation of the characteristic structural feature of the hinge region in the P4–P6 domain). We have recently begun to routinely implement separation of the products of the cleavage reaction by capillary electrophoresis (Mitra et al., 2008). This method involves footprinting unlabeled RNA molecules followed by reverse transcription (RT) of the products into fluorophore-labeled cDNA

No •OH

T1

T1*


39

(Mg2+)

147–148 142 134 129 126 116–119 110–112 108 100 96 91–92

82–83 77 73

Figure 2.1 Hydroxyl-radical footprinting of the mutant of the Tetrahymena ribozyme bearing UUCG mutation of the L5b tetraloop. The equilibrium folding experiment was performed as a function of the increasing concentration of Mg2þ in the background of 100 mM KCl at 25 C buffered by 10 mM of potassium cacodylate, pH 7.0.

40


fragments using DNA primers labeled by a fluorescent dye (Cy5 in our case) that anneal to the 30 -end of the RNA molecules. Briefly, the unlabeled RNA molecule is subjected to OH cleavage subsequent to its cation mediated folding reaction. The cleaved products are precipitated, dried, and resuspended in 9 ml of a buffer containing 50 mM Tris–HCl pH 8.3, 60 mM NaCl, and 10 mM DTT. 0.5 ml of a 10 mM Cy5-labeled primer stock solution is added to each sample, heated at 85 C for 1 min and slowly cooled to 25 C for primer annealing. We then add 9 ml of reverse transcription mix (4 ml of 5 RT buffer supplied with Superscript III (InvitrogenÒ ), 1 ml of 0.1 M DTT, 2 ml of RNase Inhibitor, 2 ml of 10 mM dNTP mix) to each tube. The solutions are incubated at 55 C for 5 min before addition of 1 ml (200 units) of Superscript III to initiate RT reaction that occurs at 55 C for 15–30 min. Upon completion of RT extension, we degrade the RNA by adding 2 ml of 2 N NaOH and incubating at 95 C for 3 min and neutralize the solution by 2 ml of 2 N HCl. Finally, 3 ml of 3 M sodium acetate and 80 ml of ethanol is added and the cDNA is precipitated by centrifugation at 16,100g for 30 min. The pellets are washed with 70% ethanol, dried, and resuspended in 40 ml of the Sample Loading Solution (BeckmanÒ ). The dideoxy sequencing reactions (for ladder lanes) are performed in the same way except that we add 0.25 mM of one of the ddNTPs. The resuspended cDNA fragments are then separated in an automated DNA sequencing apparatus (BeckmanÒ CEQ 8000 in our case). The optimized parameters that, in our hands, produce high-resolution peak traces at single nucleotide resolution for about 300 nucleotides are electrokinetic injection voltage, 2 kV; electrokinetic injection time, 7 s; denaturation, 90 C for 150 s; separation voltage, 3 kV; and capillary temperature, 60 C. The fragment of a sequencer profile that demonstrate visible differences for unfolded and folded RNA are shown in Fig. 2.2.

6. Quantitation of the Changes in the Reactivity and Data Analysis The changes in the extent of cleavage for individual or grouped nucleotides can be teased out by quantitating the changes in the intensities of the corresponding bands. When the RNA fragments are separated by The oxidative Fenton chemistry was used to produce hydroxyl radicals. The radioactively labeled RNA fragments were separated on 8% gel electrophoresis with 7 M urea. Each band represents a single nucleotide. Location of the nucleotide in the RNA sequence is assigned from the G ladder generated by RNase T1 digestion of the same RNA, run on the same gel. Changes in solvent accessibility of individual nucleotides appear as decrease or increase in intensity of separate bands or their groups.

Activity (cnts)

20,000 17,500 15,000 12,500 10,000 7500 5000 2500 0 51.5

52.0

52.5

53.0 Time (min)

53.5

54.0

54.5

55.0

51.5

52.0

52.5

53.0 Time (min)

53.5

54.0

54.5

55.0

Activity (cnts)

20,000 17,500 15,000 12,500 10,000 7500 5000 2500 0

Figure 2.2 Capillary separation of the products of the OH-induced cleavage of the RNA backbone. Peak profiles of Cy5-labeled cDNAs corresponding to cleaved unfolded (upper panel) and folded (lower panel) T. thermophila ribozyme are shown by blue lines. Red peaks corresponding to the BeckmanÒ size standard ladder are used to correlate the peaks with the RNA nucleotides. The peaks that demonstrate significant decrease in reactivity upon folding are enclosed within gray contour.

42


denaturing gel electrophoresis, we quantitate these changes either semiautomatically using the SAFA software (Chapter 3, Laederach et al., 2008) or manually using a standard gel analysis software like ImageQuantÒ in which the bands demonstrating a visible change in intensity are enclosed within a contour whose area is reported (Shcherbakova and Brenowitz, 2008). When the cleaved fragments are separated by capillary electrophoresis, the peak traces of the sequencer readout (Fig. 2.2), stored as numerical values in arbitrary units, are exported in the form of text files (.txt), are then semiautomatically processed and analyzed using the CAFA software (Mitra et al., 2008). The data normalization procedure is described in detail in Chapter 3. Briefly, the data for each sample is normalized to account for the variations in sample loading and scaled to reflect the observed changes between 0 (initial state) and 1 (final state). This normalization can be executed either manually or automatically by gel or capillary electrophoresis data analysis software. In the case of the equilibrium studies the normalized data can be fit to the Hill equation: Y i ¼

KdinH ½Mþ nH ; 1 þ KdinH ½Mþ nH

where Kdi is the equilibrium dissociation constant, [Mþ] refers to the concentration of the reagent that caused the analyzed structural changes (e.g., Mg2þ in our studies of Mg2þ-induced folding), and nH is the Hill coefficient. An example of the equilibrium curves generated for several local reporters of folding from the gel electrophoresis shown in Fig. 2.1 is presented in Fig. 2.3. We report lower and upper limits calculated at the 65% confidence interval that corresponds to one standard deviation of the data. Depending on the length of the RNA and the extent of its tertiary structure, multiple titration curves can be generated for a single RNA molecule (e.g., for the 385 nucleotide Tetrahymena ribozyme we usually generate about 30 titration curves). A simple visual inspection of the curves can answer some important questions about equilibrium folding of the RNA, for example: How concerted is the equilibrium folding? Can several distinct groups of titration curves be distinguished? Are there any obvious equilibrium intermediates present? Which structural elements are involved in different phases of the process? In some cases, it is beneficial to complement this richly detailed local information by additional information about global changes in the nucleic acid structure (Kwok et al., 2006; Laederach et al., 2007b; Schlatterer et al., 2008; Takamoto et al., 2002, 2004). Analytical ultracentrifugation (see chapter 10 in volume 469 of these series) or small angle X-ray scattering can help set constrains on the conformational space for generating coarse-grained

Fractional saturation

1.0

1.0

Nucleotides 92–95

1.0

0.8

0.8 0.6 0.4 0.2

Kd = 0.33 (0.31; 0.35) nH = 2.45 (2.13; 2.84)

0.0 0.01

0.1

1

0.6

0.4

0.4

0.2

0.2 Kd = 0.47 (0.45; 0.49) nH = 4.54 (3.84; 5.40)

0.0 0.01

1.0

1.0

Nucleotide 164

0.8

0.6

10

Nucleotides 180–182 Fractional saturation

Nucleotides 108–111

0.1

1

Kd = 0.27 (0.26; 0.28) nH = 3.6 (3.2; 4.0)

0.0

0.01

10

0.1

1

10

Nucleotide 213

Nucleotides 200–201 1.0

0.8 0.8

0.8

0.6 0.6

0.6

0.4 0.4

0.4

0.2 0.2

0.2 Kd = 0.23 (0.22; 0.24) nH = 2.91 (2.67; 3.19)

0.0 0.01

0.1 1 (Mg2+), (mM)

10

0.0

Kd = 0.45 (0.42; 0.48) nH = 5.5 (4.4; 7.0)

0.01

0.1

1

(Mg2+), (mM)

10

Kd = 0.40 (0.39; 0.41) nH = 4.2 (3.9; 4.5)

0.0 0.01

0.1

1

10

(Mg2+), (mM)

Figure 2.3 A representative set of equilibrium titration curves obtained from the gel electrophoresis shown in Fig. 2.1. The data were normalized and scaled as described in Chapter 3. The Hill coefficient nH and equilibrium dissociation constant Kd (in mM) for each of the sites that demonstrate change in the reactivity associated with increase in the concentration of Mg2þ were determined when the data were fit to the Hill equation.

44


structural models of equilibrium folding intermediates (Baird et al., 2005; Laederach et al., 2007b; Lipfert et al., 2007).

7. Conclusions Hydroxyl-radical footprinting provides vast local information about formation of tertiary structure during equilibrium RNA folding. The same approach can be used to probe specific nucleic acid–protein interactions. This simple experiment combined with a separation technique capable of single nucleotide resolution can provide information about changes in solvent accessibility of every nucleotide along the nucleic acid backbone as a function of a reagent concentration. This information can be used by itself to characterize the sequence of events and intermediate structures during equilibrium folding (or molecular association) or can be melded with data obtained by other techniques, under identical, or closely related experimental conditions. The recent advances in the data analysis (see Chapter 3) and modeling of the RNA structure to be compatible with the experimental data ( Jonikas et al., 2009) provide a valuable set of tools to interpret this information in terms of coarse-grained models of the molecular structure.

ACKNOWLEDGMENTS The authors are members of the laboratory of Michael Brenowitz within which the studies described in this chapter were conducted. The writing of this chapter was supported by grant RO1-GM085130 from the National Institute of General Medical Sciences of the National Institutes of Health. We are thankful to Michael Brenowitz for critical reading of the manuscript and his valuable suggestions.

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C H A P T E R

T H R E E

Rapid Quantification and Analysis of Kinetic OH Radical Footprinting Data Using SAFA

Katrina Simmons,* Joshua S. Martin,* Inna Shcherbakova,† and Alain Laederach* Contents 1. Introduction 2. Using SAFA 2.1. Gel cropping 2.2. Lane definition and gel rectification 2.3. Band assignment and use of a sequence file 2.4. Single-peak fitting with SAFA 2.5. Data output 3. Data Normalization 3.1. Data normalization using invariant residues 3.2. Generating time–progress curves and obtaining rate constants 3.3. Data interpretation by inspection of the time–progress curves and comparison of rate constants 4. Data Visualization 5. Conclusion Acknowledgment References

48 50 52 52 54 55 57 57 57 58 60 61 64 64 65

Abstract The use of highly reactive chemical species to probe the structure and dynamics of nucleic acids is greatly simplified by software that enables rapid quantification of the gel images that result from these experiments. Semiautomated footprinting analysis (SAFA) allows a user to quickly and reproducibly quantify a chemical footprinting gel image through a series of steps that rectify, assign, and integrate the relative band intensities. The output of this procedure is raw band intensities that report on the relative reactivity of each nucleotide with the * Developmental Genetics and Bioinformatics, Wadsworth Center, Albany, New York, USA Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts, USA

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Methods in Enzymology, Volume 468 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)68003-4

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Katrina Simmons et al.

chemical probe. We describe here how to obtain these raw band intensities using SAFA and the subsequent normalization and analysis procedures required to process these data. In particular, we focus on analyzing timeresolved hydroxyl radical (OH) data, which we use to monitor the kinetics of folding of a large RNA (the L-21 T. thermophila group I intron). Exposing the RNA to bursts of OH radicals at specific time points during the folding process monitors the time progress of the reaction. Specifically, we identify protected (nucleotides that become inaccessible to the OH radical probe when folded) and invariant (nucleotides with constant accessibility to the OH probe) residues that we use for monitoring and normalization of the data. With this analysis, we obtain time–progress curves from which we determine kinetic rates of folding. We also report on a data visualization tool implemented in SAFA that allows users to map data onto a secondary structure diagram.

1. Introduction Chemical probes such as the OH radical (Latham and Cech, 1989), dimethyl sulfate (DMS) (Tijerina et al., 2007), N-methylisatoic anhydride (NMIA) (Wilkinson et al., 2006), and various RNA modifying and cleaving enzymes (Donis-Keller et al., 1977) are commonly used to determine aspects of RNA structure and dynamics. The general principle behind all of these probes is their selective reactivity with nucleotides in different conformations or structural states (Felden et al., 1996). The information provided by these experiments indicates how the local structure of the nucleic acid affects the reactivity of the chemical probe. For example, OH radicals will react more readily with nucleotides that are solvent exposed, reporting on the solvent accessibility of particular residues (Latham and Cech, 1989). DMS reacts selectively with adenines and cytosines that are not involved in base-pairing reactions (Lempereur et al., 1985), whereas NMIA creates 20 -O-adducts with highly flexible nucleotides in the RNA (Badorrek and Weeks, 2005). Chemical mapping experiments are relatively simple to carry out experimentally, and readout is often accomplished using gel electrophoresis (Laederach et al., 2008). Nucleic acids are readily separated on electrophoretic gels with single nucleotide resolution. Large electrophoretograms such as that illustrated in Fig. 3.1A will often contain several thousand individual bands, each representative of a nucleotide’s relative chemical reactivity. Each lane in the gel can represent the RNA’s reactivity under different conditions, or as is the case in Fig. 3.1A, at different times during a folding reaction. The ability to probe RNA structure under different conditions and at different time points is a major motivation for obtaining quantitative band intensities from the gel image shown in Fig. 3.1A. The gel image shown in Fig. 3.1A reports on the Mg2þ-induced folding reaction of a mutant (L5b) of the L-21 Tetrahymena thermophila group I

Rapid Quantification and Analysis of Kinetic OH Radical Footprinting Data

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Figure 3.1 (A) Gel image of a typical chemical mapping experiment on the L5b mutant of the T. thermophila group I intron analyzed by OH radical cleavage. Each band in the gel corresponds to a specific nucleotide and each lane, in this case, a different time point along the folding reaction. Nucleotides where a visual change in band intensity is seen as a function of time are magnified in (B) and (C). (D) A zoomed in version of the gel image on the invariant residues (113–115) used for normalization. This chapter focuses on the SAFA software, which is designed to rapidly quantify the relative band intensities from such a gel image (Das et al., 2005).

intron (Laederach et al., 2007). We will use these data (along with a similar gel reporting on the folding reaction of the wild-type RNA) to illustrate the process of obtaining accurate and reproducible time–progress curves for analysis. We analyze the Mg2þ-induced folding reaction as measured by OH radical footprinting for both a wild-type and mutant RNA, which have different folding rates (Laederach et al., 2007; Shcherbakova and Brenowitz, 2005). OH radicals measure the solvent accessibility of nucleotides by selectively reacting with highly accessible residues, indicating the sites of protection, and accessibility in the molecule (Latham and Cech, 1989). The radicals are generated either using synchrotron radiation or the Fenton chemical reaction (Fenton, 1894; Shcherbakova and Brenowitz, 2008; Shcherbakova et al., 2006). The basic premise of the data we are analyzing is a relative change in the accessibility of nucleotides as the RNA folds upon the addition of Mg2þ. As shown in Fig. 3.1B, certain bands become lighter (or darker) as a function of time, indicating folding of the RNA. By sequentially cleaving the RNA with OH radicals at different time points during the reaction, we measure the fractional saturation of the

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reaction. We will use these data to generate time–progress curves, from which we can then obtain local folding rates. SAFA is software designed to facilitate the analysis of RNA footprinting gels. We describe here the major steps in analyzing these data using the software, which can be downloaded along with the example files at http:// simtk.org/home/safa. SAFA runs on Mac OS X, Windows XP, and is written using the Matlab (The Mathworks) programming language. Heavy users of the system should consider acquiring the Matlab software (which is licensed campus-wide at most academic institutions) as performance of SAFA is significantly improved when used in conjunction with Matlab. When users choose to use SAFA in conjunction with Matlab, they must download the ‘‘source’’ version of the software independently of whether they are running Matlab on a Windows or Macintosh computer. Furthermore, if users have the Windows Vista or Home edition, using Matlab will solve many compatibility issues.

2. Using SAFA Using SAFA is relatively straightforward. Users will recognize familiar menus (such as the File Menu, indicated by red arrow in Fig. 3.2A). The basic workflow of SAFA is controlled through a series of buttons (marked by green arrows) on the right side of the program in Fig. 3.2A. Clicking on each button successively leads the user through the different steps of the procedure. Initially, the software requires as input both a gel image (either with a .gel or .tiff extension, 16-bit images are generally easier to analyze with the software, although 8-bit images are supported) and a sequence file (in FASTA, .fas format). The gel image will appear in the central SAFA window (Fig. 3.2A) while reading in a sequence file will open the sequence selection dialog box (Fig. 3.2B). In general, keyboard commands or keyboard shortcuts are displayed above the main gel window in the software. Several example data sets are available for download from http:// simtk.org/home/safa including all the data presented in this chapter. Our goal is to obtain quantitative intensities for all bands shown in Fig. 3.1. As can be seen in Fig. 3.1B and C, certain bands (in this case corresponding to nucleotides C177 and G160, respectively) become significantly lighter as the folding reaction progresses. This indicates that as the molecule is folding these regions are becoming progressively more protected. By plotting the relative change in band intensity as a function of time, we can generate time–progress curves that report on the local rate of conformational change in the RNA molecule. Several steps are required to obtain these data from the gel image. The first step we describe here

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Figure 3.2 Screenshots of the SAFA software. (A) The main SAFA window used to control all aspects of the software. The analysis procedure is carried out by successively clicking on each of the buttons to the right side the main window in which the gel image is displayed. (B) The sequence selection tool allows the user to define how the semiautomated procedure of band assignment will be carried out. In this case, the sequence selection tool is setup to use a T1 ladder, which indicates the positions of guanosines on the gel. (C) The quantification progress window appears when the software is automatically fitting lanes and is used to monitor the progress and accuracy of the single-peak fitting procedure (Takamoto et al., 2004).

involves integrating the band intensity using a procedure called ‘‘singleband peak fitting’’ (Takamoto et al., 2004). Single-band peak fitting fits a model to the raw image data, but requires accurate initial guesses of the relative positions of the bands on the gel. The inevitable imperfections in the gel matrix yield gel images with nonparallel lanes and ‘‘smiles’’ (or ‘‘frowns’’) that make the problem of estimating band positions quite difficult. SAFA includes functionality to ‘‘rectify’’ the gel image, effectively correcting the imperfections so as to have parallel lanes, and no smiles (or frowns) in the gel image. Details of the procedure for gel rectification can be found in previous publications (Laederach et al., 2008) and a video demonstration of the process can be downloaded from

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http://simtk.org/home/safa. We recommend viewing the video tutorials, as the gel rectification process is fundamentally a visual process, requiring accurate user input. For the purposes of this method, we will briefly outline the procedure for the example data set we use here.

2.1. Gel cropping The gel image must be initially cropped. In Fig. 3.3A, we show a screenshot of the main SAFA window in which we have loaded the gel image into SAFA. Using the cursor, we have drawn a cropping box around the right half of the gel image, which reports on the folding progress of the L5b mutant of the T. thermophila group I intron. We are careful to include in our cropping the T1 digest lane, which will help us assign the bands to the sequence in later steps. In general, a large enough portion of the gel image should be kept so that all lanes are encompassed.

2.2. Lane definition and gel rectification The next step in the SAFA procedure involves defining lanes. This procedure is used to vertically align the gel by identifying lane boundaries. Such lane boundaries are generally trivial to identify by eye, but present a significant computational challenge when attempted in a fully automatic way. SAFA therefore uses a semiautomated approach to resolve this problem. The user initially defines at least one lane by drawing the boundaries of that lane using a click and draw procedure, analogous to a line tool in a computer graphics program. The user can then invoke an automated procedure for ‘‘guessing’’ the next lane boundaries, which uses the previous lane definitions as a starting point. This automated procedure is invoked by typing ‘‘G.’’ The procedure of guessing lane boundaries is continued until all lanes are correctly selected, at which point the user invokes a vertical rectification procedure by typing ‘‘Z.’’ The vertical rectification procedure uses a linear interpolation routine that preserves the total pixel count in the gel image. The result of this procedure is illustrated in Fig. 3.3B, where each lane is properly defined by red vertical lines. SAFA then identifies the center of the lanes as illustrated by dotted lines in Fig. 3.3C. Once the lanes have been defined, the gel now requires horizontal alignment. The horizontal alignment procedure in SAFA is analogous to the vertical one. In this case, horizontal lines are drawn through corresponding bands in the gel image as shown in Fig. 3.3D. These lines should trace through the bands in the gel and will act as anchor lines for the horizontal rectification procedure. This procedure is manual, and the user should choose several well-resolved bands throughout the gel image as shown in Fig. 3.3D. The user can zoom in and out by right clicking to more accurately trace through


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Figure 3.3 Illustration of the gel rectification procedure implemented in SAFA. (A) Screenshot of the main SAFA window with a gel image loaded; red box delineates the user-defined cropping boundary for subsequent analysis. (B) Screenshot of the lane definition procedure. Red line is manually drawn that marks the boundaries of each lane. Once a few lanes have been manually defined, SAFA is able to ‘‘guess’’ the outline of the following lanes, which is done by typing the letter ‘‘G.’’ (C) Screenshot of the completed and recorded defined lanes that occur after the user types ‘‘Z’’ or ‘‘Q.’’

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the bands in the gel. The number of such anchor lines required and their position is dependent on the extent of horizontal deformation within the gel image. The ultimate result (Fig. 3.3E) is a gel that is both vertically and horizontally aligned, making the process of band assignment straightforward.

2.3. Band assignment and use of a sequence file As noted above, SAFA requires two input files, a gel image and a FASTA sequence file. The FASTA file for the L-21 T. thermophila group I intron is shown below: >Group I Intron, T. thermophila GGAGGGAAAAGUUAUCAGGCAUGCACCUGGUAGCUAGU CUUUAAACCAAUAGAUUGCAUCGGUUUAAAAGGCAAGACC GUCAAAUUGCGGGAAAGGGGUCAACAGCCGUUCAGUACCA AGUCUCAGGGGAAACUUUGAGAUGGCCUUGCAAAGGGUAU GGUAAUAAGCUGACGGACAUGGUCCUAACCACGCAGCCAA GUCCUAAGUCAACAGAUCUUCUGUUGAUAUGGAUGCAGU UCACAGACUAAAUGUCGGUCGGGGAAGAUGUAUUCUUCU CAUAAGAUAUAGUCGGACCUCUCCUUAAUGGGAGCUAGCG GAUGAAGUGAUGCAACACUGGAGCCGCUGGGAACUAAUUU GUAUGCGAAAGUAUAUUGAUUAGUUUUGGAGU In general, the first line of the FASTA file contains some descriptive text of the sequence preceded by a ‘‘greater than’’ character (>). It is also important to input the RNA sequence (replacing T with U) if the molecule being studied is in fact RNA. SAFA can also be used to study DNA, and therefore distinguishes between T and U in the sequence. Most importantly, the file should be saved as a ‘‘Western ASCII’’ text file, especially if it is being edited in a word processing application. SAFA assumes that the first nucleotide in the file corresponds to the first nucleotide in the sequence, unless an offset is set in the sequence selection tool (Fig. 3.2B). The sequence selection tool (Fig. 3.2B) defines how the user will assign bands on the gel. It is invoked by clicking on ‘‘load sequence’’ (Fig. 3.2A, green arrow marked 1) in the main SAFA window. The general concept The color of the lines changes from red to green and a dotted line marks the middle of the lane when the procedure in completed. (D) Screenshot of the horizontal alignment procedure. The gel is aligned horizontally to ensure the bands are parallel. Similar to the lane definition process, the user identifies bands across the gel image with a horizontal line. Only one line should be drawn per band. The left button of the mouse begins the line and marks sequential points throughout the row and the right button ends the line. The number of rows to mark is at the users’ discretion; however, the more rows used will result in a more accurate an adjustment of the image. (E) Screenshot of the finished horizontally aligned gel. (F) Screenshot of the band assignment procedure.


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behind the sequence selection tool is that one or several ladder lanes will be run in a separate lane on the gel being analyzed. In the case of RNA, T1 digests are often used to identify guanosine nucleotides; this is the case in Fig. 3.1. Alternatively, sequencing ladders can also be run to identify any (or all) of the other nucleotides (Tijerina et al., 2007). Ultimately, with the gel rectified, the user will click on the bands they have chosen with the sequence selection tool. As is illustrated in Fig. 3.2B, the user has selected ‘‘G’’ and set an offset of 21, since the first nucleotide in the sequence file corresponds to the 21st residue of the T. thermophila group I intron. The user will only need to identify bands corresponding to G residues, and SAFA will guess the positions of all the residues in-between. This semi-automated procedure allows the user to precisely define bands in the gel across all lanes rapidly. Furthermore, by adjusting the parameters in the sequence selection tool, the user has the option of selecting one or multiple nucleotides for manual definition during the assignment process. In general, selecting one nucleotide is sufficient, as SAFA is able to extrapolate the positions of the other nucleotides accurately during the assignment process. The sequence selection tool also allows the user to specify that the RNA is 30 -labeled, indicating that the 50 -band will be at the bottom of the gel. This option can also be used if the gel to be analyzed is the result of reverse transcription and the user wishes to use the RNA sequence. SAFA will use a single-peak fitting algorithm to fit the data, which can handle overlapping bands. As a result, the user can choose to continue to assign bands in the overlapping region of the gel. How ‘‘high’’ to go in the gel will depend greatly on the quality of the gel image, desired quality of the data, and complexity of the experiment. Users will need to evaluate the reproducibility of the procedure on their own data to evaluate the degree to which they can analyze highly overlapping regions in the gel. In the example we provide along with this chapter, we analyze the gel conservatively, obtaining data for the first 100 bands.

2.4. Single-peak fitting with SAFA Peak fitting is a fully automated process in SAFA and is activated by clicking on the ‘‘quantify’’ button in the main SAFA window (Fig. 3.2A, green arrow marked 7). SAFA will only perform the peak-fitting analysis if all the previous steps have been completed. Progress in the peak-fitting procedure is monitored through a series of three graphs that appear when the peak-fitting procedure is run (Fig. 3.2C). SAFA proceeds through each lane of the gel successively, allowing the user to visualize the quality of the fit. Visualizing the resulting fit is key when troubleshooting the quantification procedure, as potential errors in the fitting procedure can easily be seen when looking at these results. SAFA will plot all of the fits again once the fitting procedure is complete, to encourage users to look carefully at the traces.

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We illustrate in Fig. 3.4A a successful single-peak fitting procedure on Lane 4 of the gel shown in Fig. 3.1A. In Fig. 3.4A, SAFA plots the individual peaks in red, and the overall lane profile in blue. In this case, the individual Lorentzian peaks are evenly spaced, and have uniform widths. In general, SAFA optimizes the fit so as to have regularly spaced peaks with linearly varying widths. However, if the initial positions of the peaks as defined by the band assignment procedure (Fig. 3.3F) are inaccurate, SAFA will find a solution with large differences in peak widths, as is illustrated in Fig. 3.4B. In this case, the user has incorrectly assigned bands and the solution found by SAFA has irregularly spaced peaks and highly variable

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Figure 3.4 SAFA performs a peak-fitting procedure to determine the relative band intensity in the gel image. When the fitting procedure has converged, SAFA will plot the results as shown here. Red lines represent individual Lorentzians that when summed yield the lane profile shown in blue. (A) A successful and accurate singlepeak fitted graph for Lane 15 of the gel shown in Fig. 3.1. (B) Visual inspection of the fitted profile can be used to determine if the user was not accurate in assigning bands (Fig. 3.3F). In this example, there are large differences in the peak widths and irregularly spaced peaks. These are signs of inaccurate results.


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peak widths. SAFA will replot the fitted data as illustrated in Fig. 3.4 for each lane after the fitting procedure, and it is recommended to visually inspect these plots to identify any fits that look like Fig. 3.4B, as these will indicate potential inaccuracy in the results. Although SAFA will check for convergence of the fitting procedure and provide an error message if the convergence criteria are not met, it is often still possible to obtain convergence on an incorrect solution as is illustrated in Fig. 3.4B. Visual inspection is thus an important final step of the fitting procedure.

2.5. Data output At this stage, SAFA will output a text file with the raw band intensities determined through the single-peak fitting procedure. The text file is tab delimited, with the first column containing the residue numbers, and the next columns the actual raw band areas, as determined by peak fitting. It should be noted that the data are output in scientific notation, and when the file is opened in a spreadsheet program like Excel, the nucleotide numbers are displayed in scientific notation as well. Interpretation of the raw data, however, requires further analysis, normalization, and visualization. Although some users at this stage may prefer to perform these analyses in other software, SAFA offers several other tools for data visualization and normalization. The automated normalization functionality of SAFA is described in detail in a previous publication (Laederach et al., 2008); we choose here instead to analyze the time-resolved data manually to better illustrate the role of normalization in obtaining time–progress curves from these data (Shcherbakova et al., 2006). We also turn our focus to a previously unpublished tool built into SAFA for the visualization/projection of chemical mapping data on RNA secondary structure diagrams. These two approaches are critical for the correct interpretation of chemical mapping data.

3. Data Normalization 3.1. Data normalization using invariant residues The first step of data normalization requires correcting for nonuniform loading of the RNA into each lane. This procedure involves identifying invariant residues, that is, nucleotides where the relative density changes reflect the variations in the amount of RNA loaded per lane. The choice of the invariant residues is based on visual inspection of the gel image to identify bands that demonstrate no visible systematic change in intensity. Furthermore, any available structural information can be used to ensure that the selected residues belong to a region where structural changes are not

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expected. To verify whether the choice of invariant bands was correct, we plot the invariant band intensities as a function of reaction time to rule out a systematic increase or decrease in the band intensities. Usually, a single set of invariant residues can be used for all the protections and sites of the enhanced reactivity on the same gel. In the case of our example, we chose nucleotides 113–115 (Fig. 3.1D), which are in the P4 helix of L-21 T. thermophila, a highly stable region of the molecule that does not change conformation during folding. In general, helical segments work best as invariant residues. To normalize the time-resolved data, we divide the intensities of the all the bands by the average intensity of the invariant residues. The result of this procedure is illustrated in Fig. 3.5 in which the raw data (Fig. 3.5A) and invariant normalized data (Fig. 3.5B) for the wild-type Tetrahymena ribozyme (black) and L5b mutant is plotted (red symbols). The effect of this simple procedure is dramatic, resulting in uniform time–progress curves which can be subjected to further analysis. We provide the MicrosoftÒ Excel spreadsheet used to normalize these data at http://simtk.org/home/ safa. It should be noted that small variations in sample loading are easily corrected in this way. However, if large variations in sample loading are observed, this will lead to much greater noise in the data regardless of which invariant residues are chosen. The second step of data normalization aims to determine the extent of protection as a fractional value between 0 (initial state) and 1 (final state). This is accomplished by dividing the band intensity by the absolute in band intensity between the final and initial states. Following this transformation, the intensities are individually scaled to fractional saturation and this is illustrated in Fig. 3.5C. This transformation creates uniform kinetic data, which allows us to compare changes in intensity between different molecules. In this case, it is clear that the wild-type and mutant RNA (black and red in Fig. 3.5, respectively) have different rates of formation. For the three protections, we illustrate in Fig. 3.5, the L5b mutant appears to fold slower.

3.2. Generating time–progress curves and obtaining rate constants The normalized data are best visualized in Fig. 3.5C by plotting the fractional saturation of the individual sites that demonstrate changes in the reactivity as a function of time. The reaction time is calculated as a sum of the quench-flow delay time and half of the cleavage (or modification) reaction time. To visualize the details of the fastest events the X-axis is often set to a logarithmic scale. The collection of the individual time points can be fit to an exponential function to discern rate constants for the processes that are reflected in the time-resolved changes in the local reactivity to the footprinting reagent

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Figure 3.5 Time–progress curves following SAFA quantification. The wild-type data are shown in black and the L5b mutant in red. (A) When the data that are not normalized are plotted as a function of time, it is difficult to identify any trends in the changes in reactivity.

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(Fig. 3.5C, lines). These changes are best described in terms of an exponential function: X Y ¼ 1 ai expðki tÞ; i¼1

where ai and ki are the amplitude and rate constant, respectively, of the ith kinetic phase. When fitting the data we start with a single exponential parameter (i ¼ 1). If a systematic error arises as a result of the fitting two rate constants are fit. We usually report lower and upper limits calculated at the 65% confidence interval that corresponds to one standard deviation of the data.

3.3. Data interpretation by inspection of the time–progress curves and comparison of rate constants In many cases, a working hypothesis can be discerned from a simple inspection of the time–progress curves and the comparison of the rate constants. For example, for the limited number of time–progress curves illustrated in Fig. 3.5, we conclude that not all the time–progress curves demonstrate the same kinetic behavior. The significant offset in the reactivity (or burst) between initial state and the first data point for the wild-type data (Fig. 3.5, black curves) indicates that the onset of the folding process is faster than the dead time of the experiment. In this case better time resolution would be beneficial to characterize the fastest events of the folding under this condition. Improved time resolution is achieved by Fast Fenton footprinting (Shcherbakova et al., 2006) and will reveal the multiple kinetic phases of the local changes in the reactivity at shorter time points. The data we present here, however, were collected using synchrotron radiation to generate OH radicals, which did not have the time resolution to resolve the fastest events in the folding of the wild-type molecule (Ralston et al., 2000). We can nonetheless conclude from this analysis that the mutation of the L5b tetraloop causes significant changes in the folding mechanism, which yield slower rates (Fig. 3.5C). The presence of a distinct hierarchical folding mechanism for the wild-type ribozyme is gleaned from the comparison of the rate constants and the amplitudes of the fastest folding events: formation of the hinge region of the P4–P6 domain, reported by increase in the reactivity for nucleotide 122, precedes to the formation of junction region (B) When invariant residue normalization is applied (in this case based on invariant residues 113–115), much clearer changes in reactivity are revealed. (C) When further normalization procedures are applied based on setting the initial and final fractional saturation to zero for t ¼ 0 and one for the final state, we are able to compare the data between experiments and determine rate constants for the transitions.


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between P4–P6 domain and the catalytic core (nucleotides 109–112), whereas the catalytic core (nucleotides 95–97) is the slowest to fold. A complete analysis of these data is described in Laederach et al. (2007) where we model the folding pathways of the RNA based on these experiments. An automated approach to modeling this data kinetically is the Kinfold software described in detail by Martin et al. (2009).

4. Data Visualization SAFA outputs a large table of numbers, which alone can be difficult to interpret. If the data measure time progress of the folding reaction, time– progress curves can be generated and visualized as described above (Fig. 3.5). Often, however, the ability to project the data onto the twodimensional structure of the RNA is valuable in identifying regions of the molecule that are highly reactive to a particular probe. We have therefore developed a way to quickly and clearly map the quantified data generated through SAFA analysis onto the molecule’s secondary structure. Our idea is based on the fact that each lab has their favorite secondary structure diagram, and that a tool able to map data directly onto this diagram will be most valuable in interpreting the data within the structural context of the experiment. Other tools such as VARNA (Ferre et al., 2007) and S2S ( Jossinet and Westhof, 2005) generate automated secondary structure representations on which users may map data. These representations, however, do not always correspond to the context in which the data are being visualized, and as such we decided a tool that can facilitate plotting of data onto a userdefined diagram would have value. This tool can be accessed from the Secondary Structure Plot menu item under Visualize Data in SAFA. The visualization tool requires two inputs, SAFA data and a highresolution (>150 dpi) image of the secondary structure. SAFA reads either JPEG, GIF, or TIFF formatted pictures with the preferred format TIFF (without any form of compression). The image should have some space reserved on one side for a color legend. This procedure allows the user to manually define the relative positions on the image once and plot any data onto the image thereafter. It is therefore worthwhile obtaining a highquality image before carrying out the manual parts of the procedure. There are four steps required to generate a figure: 1. The image (in JPEG, GIF, or TIFF format) of the molecule’s secondary structure is loaded using the SecStruct Image button (Fig. 3.6A, red arrow) When the image loads correctly, ‘‘loaded’’ should appear to the right of the SecStruct Image button, and the image will appear in the main window as shown in Fig. 3.6B.

(A)

(E)

(B)

(F)

(C)

(G)

(D)

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Figure 3.6 The data visualization component allows users to map their data onto secondary structure diagrams. (A) Screenshot of the starting window that opens when Visualize Data ! Secondary Structure Plot is selected. Selecting the SecStruct Image button (highlighted by red arrow) allows the user to load a JPEG, GIF, or TIFF image of their secondary structure. (B) Once the image is loaded, it will appear in the window and the next step is to pick points on the image. The Pick Points button indicated by blue arrow begins the procedure where the user defines the location of each individual nucleotide on the diagram. Magenta arrow indicates the Square Width button, which determines the size of the box that is placed over the selected residues, and green arrow indicates the Offset button that allows the user to account for any offset in the start nucleotide of the nucleic acid. (C) A new window appears when the Pick Points button is pushed. Indications for keyboard shortcuts are printed above the upper X-axis along with the next box number indicated by orange arrow. (D) An example of pick points procedure on the T. thermophila group I intron secondary structure diagram. (E) An example of a completed procedure on the same figure. Each nucleotide in the diagram is covered by a square. (F) Once the procedure of picking points is completed, the image and corresponding boxes are saved


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2. The Pick Points (Fig. 3.6B, blue arrow) button will become active once an image is loaded. Clicking the Pick Points button will begin the procedure for indicating the location of nucleotides on the secondary structure diagram. The user can place squares over each residue in the corresponding region of the molecule. SAFA assumes that the first square to be placed corresponds to nucleotide number one, unless the user specifies an offset (Fig. 3.6B, green arrow). Each box added by a left click corresponds to a particular numbered nucleotide starting from the offset and increasing by increments of one. To assist in determining which box corresponds to what nucleotide, the next box number is indicated by orange arrow in Fig. 3.6C. To place successive squares, the left button is clicked. The right button on the mouse zooms in and out and inaccurately placed boxes are removed by either clicking the middle button on the mouse or moving the crosshairs over the box and typing ‘‘E’’ on the keyboard. The size of the squares can be set with the Square Width button (magenta arrow in Fig. 3.6B). The value must be a whole number. An example of the pick points procedure on the 50 -region of the T. thermophila group I intron is shown in Fig. 3.6D. Once all the boxes have been matched with their corresponding nucleotides (as illustrated in Fig. 3.6E), clicking either Z or Q will end the pick points procedure. The figure will return to its original window. Here, red highlighted boxes will become blue, and this signifies the picking points procedure is done. Figure 3.6F illustrates the result of the completed procedure. 3. Once all the points have been picked, clicking the Protection Data button (indicated by black arrow in Fig. 3.6F) will invoke a dialog to input the text file. A new window will appear with a color plot of the data (Fig. 3.6G). The user can then choose the lane corresponding to the data that he/she wishes to visualize on the secondary structure diagram. The colors used to represent the data are defined in the following manner: White nucleotides correspond to ‘‘zero’’ intensity. The value of the zero level is initially set to be the average of the minimum and maximum intensities in the data. Each residue that has higher reactivity will be colored red with intensity on a linear scale from this zero level to a maximum value. The maximum value is also a default value and is plotted in solid red. Similarly, residues with and can also be reloaded. The boxes change color from red to blue signifying the points have been saved. The light blue arrows mark the Load SecStruct Data and Save SecStruct Data buttons that allow the user to save their progress and load their data. Black arrow highlights the Protection Data button that initiates the next step in the process where the user chooses the data to be displayed on the figure. The purple arrows indicate the Set Max/ Zero/Min Value buttons that allow the user to adjust these values. (G) Representation of the SAFA quantified data for the gel image shown in Fig. 3.1 in a color plot allowing the user to choose which lane to plot on the secondary structure. (H) Mapping of Lane 15 onto the secondary structure of the T. thermophila group I intron indicating nucleotides that are protected in blue and cleaved by OH radicals in red.

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reactivity values lower than the zero value will be colored blue with intensity on a linear scale from the zero level to a minimum value. The default minimum value is plotted as solid blue. The maximum, minimum, and zero values are adjusted by clicking the appropriate Set Max/Zero/Min Value buttons (Fig. 3.6F, purple arrows). 4. When the structure image, residue locations, and accessibility data have all been loaded or assigned, the Generate Figure button will become active. This generates a full-sized figure showing the reactivity levels plotted on the secondary structure of the molecule (Fig. 3.6H). The plot protection legend checkbox controls whether the color legend is displayed in the preview plot as well as in the full-sized figure. If a legend is being displayed, its location is indicated before saving the figure or exporting to the file format of choice. Any nucleotides where no data are available will be covered in a gray box (as illustrated in Fig. 3.6H). The secondary structure image and the points picked for the residue locations on that image can be loaded independently from previously saved data files using the SecStruct Image and Load Pickpoints buttons, and all fields can be saved and later loaded together from those data files using the Load/ Save SecStruct Data buttons (Fig. 3.6F, light blue arrows).

5. Conclusion SAFA is a software package specifically designed for obtaining quantitative and reproducible data from a gel image of a chemical mapping reaction. It was originally intended for use mostly with OH radical footprinting gels (Brenowitz et al., 2002; Gross et al., 1998; Shcherbakova et al., 2006; Woodson, 2008), but is now being used for many other applications in nucleic acid chemical mapping (Vicens et al., 2007; Wilkinson et al., 2005, 2006). Since most chemical mapping experiments use similar principles to OH radical footprinting for the analysis of the reactions, it is easy to adapt SAFA to a wide range of applications. The sequence browser (Fig. 3.2B) is instrumental in defining how the band assignment procedure will behave, and by using the correct sequence and selection of nucleotides; the user can define any arbitrary combination of nucleotides. This flexibility, along with the gel rectification procedure, provide a series of tools for the rapid and accurate quantification of chemical mapping gels (Das et al., 2005).

ACKNOWLEDGMENT This work is supported the US National Institutes of Health (NIGMS) through grants R00-79953 and R21-87336 to A. L.


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REFERENCES Badorrek, C. S., and Weeks, K. M. (2005). RNA flexibility in the dimerization domain of a gamma retrovirus. Nat. Chem. Biol. 1, 104–111. Brenowitz, M., Chance, M. R., Dhavan, G., and Takamoto, K. (2002). Probing the structural dynamics of nucleic acids by quantitative time-resolved and equilibrium hydroxyl radical ‘‘footprinting’’. Curr. Opin. Struct. Biol. 12, 648–653. Das, R., Laederach, A., Pearlman, S. M., Herschlag, D., and Altman, R. B. (2005). SAFA: Semi-automated footprinting analysis software for high-throughput quantification of nucleic acid footprinting experiments. RNA 11, 344–354. Donis-Keller, H., Maxam, A. M., and Gilbert, W. (1977). Mapping adenines, guanines, and pyrimidines in RNA. Nucleic Acids Res. 4, 2527–2538. Felden, B., Florentz, C., Westhof, E., and Giege, R. (1996). Usefulness of functional and structural solution data for the modeling of tRNA-like structures. Pharm. Acta Helv. 71, 3–9. Fenton, H. J. H. (1894). What species is responsible for strands scission in the reaction of Fe(II) EDTA2 with H2O2 with DNA? J. Am. Chem. Soc. 6, 899. Ferre, F., Ponty, Y., Lorenz, W. A., and Clote, P. (2007). DIAL: A web server for the pairwise alignment of two RNA three-dimensional structures using nucleotide, dihedral angle and base-pairing similarities. Nucleic Acids Res. 35, W659–W668. Gross, P., Arrowsmith, C. H., and Macgregor, R. B. Jr. (1998). Hydroxyl radical footprinting of DNA complexes of the Ets domain of PU.1 and its comparison to the crystal structure. Biochemistry 37, 5129–5135. Jossinet, F., and Westhof, E. (2005). Sequence to structure (S2S): Display, manipulate and interconnect RNA data from sequence to structure. Bioinformatics 21, 3320–3321. Laederach, A., Shcherbakova, I., Jonikas, M. A., Altman, R. B., and Brenowitz, M. (2007). Distinct contribution of electrostatics, initial conformational ensemble, and macromolecular stability in RNA folding. Proc. Natl. Acad. Sci. USA 104, 7045–7050. Laederach, A., Das, R., Vicens, Q., Pearlman, S. M., Brenowitz, M., Herschlag, D., and Altman, R. B. (2008). Semiautomated and rapid quantification of nucleic acid footprinting and structure mapping experiments. Nat. Protoc. 3, 1395–1401. Latham, J. A., and Cech, T. R. (1989). Defining the inside and outside of a catalytic RNA molecule. Science 245, 276–282. Lempereur, L., Nicoloso, M., Riehl, N., Ehresmann, C., Ehresmann, B., and Bachellerie, J. P. (1985). Conformation of yeast 18S rRNA. Direct chemical probing of the 50 domain in ribosomal subunits and in deproteinized RNA by reverse transcriptase mapping of dimethyl sulfate-accessible. Nucleic Acids Res. 13, 8339–8357. Martin, J. S., Simmons, K., and Laederach, A. (2009). Exhaustive enumeration of kinetic model topologies for the analysis of time-resoved RNA folding. Algorithms 2, 200–214. Ralston, C. Y., Sclavi, B., Sullivan, M., Deras, M. L., Woodson, S. A., Chance, M. R., and Brenowitz, M. (2000). Time-resolved synchrotron X-ray footprinting and its application to RNA folding. Methods Enzymol. 317, 353–368. Shcherbakova, I., and Brenowitz, M. (2005). Perturbation of the hierarchical folding of a large RNA by the destabilization of its Scaffold’s tertiary structure. J. Mol. Biol. 354, 483–496. Shcherbakova, I., and Brenowitz, M. (2008). Monitoring structural changes in nucleic acids with single residue spatial and millisecond time resolution by quantitative hydroxyl radical footprinting. Nat. Protoc. 3, 288–302. Shcherbakova, I., Mitra, S., Beer, R. H., and Brenowitz, M. (2006). Fast Fenton footprinting: A laboratory-based method for the time-resolved analysis of DNA, RNA and proteins. Nucleic Acids Res. 34, e48.

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Takamoto, K., Chance, M. R., and Brenowitz, M. (2004). Semi-automated, single-band peak-fitting analysis of hydroxyl radical nucleic acid footprint autoradiograms for the quantitative analysis of transitions. Nucleic Acids Res. 32, E119. Tijerina, P., Mohr, S., and Russell, R. (2007). DMS footprinting of structured RNAs and RNA–protein complexes. Nat. Protoc. 2, 2608–2623. Vicens, Q., Gooding, A. R., Laederach, A., and Cech, T. R. (2007). Local RNA structural changes induced by crystallization are revealed by SHAPE. RNA 13, 536–548. Wilkinson, K. A., Merino, E. J., and Weeks, K. M. (2005). RNA SHAPE chemistry reveals nonhierarchical interactions dominate equilibrium structural transitions in tRNA(Asp) transcripts. J. Am. Chem. Soc. 127, 4659–4667. Wilkinson, K. A., Merino, E. J., and Weeks, K. M. (2006). Selective 20 -hydroxyl acylation analyzed by primer extension (SHAPE): Quantitative RNA structure analysis at single nucleotide resolution. Nat. Protoc. 1, 1610–1616. Woodson, S. A. (2008). RNA folding and ribosome assembly. Curr. Opin. Chem. Biol. 12, 667–673.

C H A P T E R

F O U R

High-Throughput SHAPE and Hydroxyl Radical Analysis of RNA Structure and Ribonucleoprotein Assembly Jennifer L. McGinnis, Caia D. S. Duncan, and Kevin M. Weeks Contents 1. Introduction 2. Theory 3. Practice 3.1. RNA folding 3.2. Chemical probing of RNA secondary and tertiary structure 3.3. Primer extension to map modification and cleavage sites 3.4. cDNA analysis by capillary electrophoresis 3.5. Data analysis 3.6. Normalization 3.7. SHAPE-directed RNA structure prediction 4. Examples and Interpretation 4.1. High-throughput structure analysis of protein-assisted RNA folding of the bI3 RNA 4.2. Identification of a misfolded free RNA state using SHAPE 4.3. Identification of a protein cofactor that binds the ubiquitous RNA tetraloop–receptor motif 4.4. Mechanism of cooperative folding of the bI3 RNP 5. Perspectives and Conclusion Acknowledgments References

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Abstract RNA folds to form complex structures vital to many cellular functions. Proteins facilitate RNA folding at both the secondary and tertiary structure levels. An absolute prerequisite for understanding RNA folding and ribonucleoprotein (RNP) assembly reactions is a complete understanding of the RNA structure at Department of Chemistry, University of North Carolina, Chapel Hill, North Carolina, USA Methods in Enzymology, Volume 468 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)68004-6

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each stage of the folding or assembly process. Here we provide a guide for comprehensive and high-throughput analysis of RNA secondary and tertiary structure using SHAPE and hydroxyl radical footprinting. As an example of the strong and sometimes surprising conclusions that can emerge from highthroughput analysis of RNA folding and RNP assembly, we summarize the structure of the bI3 group I intron RNA in four distinct states. Dramatic structural rearrangements occur in both secondary and tertiary structure as the RNA folds from the free state to the active, six-component, RNP complex. As high-throughput and high-resolution approaches are applied broadly to large protein–RNA complexes, other proteins previously viewed as making simple contributions to RNA folding are also likely to be found to exert multifaceted, long-range, cooperative, and nonadditive effects on RNA folding. These protein-induced contributions add another level of control, and potential regulatory function, in RNP complexes.

1. Introduction RNA is actively involved in diverse cellular processes, including protein synthesis, gene regulation, and maintenance of genome stability (Gesteland et al., 2006). In many critical examples, RNA function is directly related to its ability to fold back on itself to form complex higher order structures, or in some cases, the ability to achieve multiple structures. For example, during mRNA processing, catalytic RNAs such as group I and II introns, RNase P, small ribozymes, and some riboswitches fold into highly ordered threedimensional structures to cleave RNA (Cochrane and Strobel, 2008; Scott, 2007; Torres-Larios et al., 2006). Other RNAs, especially riboswitches, sample multiple stable conformations that then allow these RNAs to function as metabolite sensors that regulate gene expression (Edwards et al., 2007; Winkler and Breaker, 2005). Many RNAs, especially larger RNAs like catalytic introns and the ribosome, also recruit proteins to facilitate fast and stable folding in the cell. The proteins that facilitate RNA folding are generally classified into two categories, chaperones and cofactors (Herschlag, 1995; Rajkowitsch et al., 2007; Weeks, 1997). Chaperone proteins interact transiently with RNA and function to allow an RNA to refold and achieve a thermodynamically favored structure. Cofactor proteins are defined by their ability to bind stably to an RNA to create a structurally well-defined ribonucleoprotein (RNP) complex. Cofactors often stabilize a specific, active, tertiary structure. The mechanisms by which proteins facilitate RNA folding can be complex and emerging work has begun to emphasize that some proteins do not fit clearly into the traditional definitions of either chaperones or cofactors. An absolute prerequisite for understanding RNA folding and RNP assembly reactions is a complete understanding of the RNA structure at

High-Throughput RNA Structure Analysis

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each stage of the predominant folding or assembly pathway. Information regarding the final, stable, structure of an RNA or RNA–protein complex can be obtained by high-resolution NMR or by crystallography or, in a few cases, by molecular modeling. However, for most initial and intermediate states, RNA secondary and tertiary structure must be inferred from either comparative sequence analysis or chemical probing approaches. The most successful method for determining an RNA secondary structure has been by phylogenetic comparative sequence analysis (Gutell et al., 2002; Michel and Westhof, 1990). However, covariation analysis only demonstrates that a pairing is likely to exist and not that a physical pairing occurs (Woese et al., 1980). Covariation analysis also does not reveal whether a specific physical pairing actually exists in any given RNA state or folding intermediate. Alternatively, RNA structure information can be inferred experimentally by treating an RNA with small molecule or enzyme reagents that are sensitive to local RNA structure. Two broad classes of experiments have proven especially useful. First, many chemical and enzymatic reagents react with partial selectivity toward single-stranded nucleotides. Reactivity patterns obtained using these reagents provide information useful for inferring base-pairing interactions (Brunel and Romby, 2000; Ehresmann et al., 1987). Second, Fenton chemistry can be used to generate short-lived hydroxyl radicals in solution, which then react with the RNA backbone in a way that is sensitive to solvent accessibility (Tullius and Greenbaum, 2005; Tullius et al., 1987). Identification of RNA backbone positions that are protected from the hydroxyl radical reagent can provide strong evidence for higher order tertiary interactions (Latham and Cech, 1989). Conventional approaches for analyzing RNA base pairing or tertiary interactions often lead to a view of an RNA structure that is incomplete in key features. Conventional secondary structure-selective chemical and enzymatic RNA mapping reagents tend to have a narrow dynamic range and typically yield information for one-half, or less, of the nucleotides in an RNA. In addition, both secondary structure-sensitive and hydroxyl radicalmediated solvent accessibility experiments have been analyzed predominantly by sequencing gel electrophoresis. The effort required to analyze long, intact, RNAs by this approach is very large. A useful approach can be to focus on short RNAs, on simplified models of larger RNAs, or on a small region within a large RNA. The challenge in focusing on a segment of a large RNA is that long-range and unanticipated interactions, critical for understanding how an RNA functions or how proteins affect an RNA folding process, will be missed. These challenges can be addressed by new chemistries and high-throughput analysis approaches. Two methods that, in principle, yield comprehensive single nucleotide resolution information about RNA secondary and tertiary structure are SHAPE (selective 20 -hydroxyl acylation analyzed by primer extension) (Merino et al., 2005; Wilkinson et al., 2005, 2006) and the aforementioned

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hydroxyl radical footprinting (Brenowitz et al., 2002; Latham and Cech, 1989; Shcherbakova and Brenowitz, 2008; Shcherbakova et al., 2008; Tullius and Greenbaum, 2005; Tullius et al., 1987). SHAPE chemistry interrogates local nucleotide flexibility, while hydroxyl radical footprinting assesses solvent accessibility and, thus, the global RNA fold. Both techniques are adaptable to a variety of reaction conditions, including the presence of proteins. Additionally, both techniques are insensitive to nucleotide identity and can potentially provide structural information at nearly every nucleotide position in an RNA. When used in tandem, SHAPE and hydroxyl radical footprinting comprehensively probe multiple levels of RNA structure and provide an impressively detailed view of an RNA or RNP folding state or assembly intermediate. Combining these sensitive structural probes with an experimental readout using capillary electrophoresis makes it possible to characterize 350–600 nucleotides (nt) in a single high-throughput experiment. Thus, large RNAs and complex RNP structures can be readily studied in detail. These developments in high-throughput in-solution RNA structure analysis are revealing unanticipated, and complex, mechanisms for how RNAs fold and interact with proteins to achieve a final active structure. In this review, we will focus on the bI3 group I intron RNA and its RNP complexes as an example of how these technological advances are leading to new insights into RNA folding and RNP assembly reactions.

2. Theory SHAPE chemistry is a robust and comprehensive method for analyzing local nucleotide dynamics in RNA (Merino et al., 2005; Wilkinson et al., 2005, 2006). In a SHAPE reaction, the RNA is treated with an electrophilic reagent like NMIA or 1M7 (Merino et al., 2005; Mortimer and Weeks, 2007) that selectively reacts with the ribose 20 -hydroxyl group at conformationally flexible nucleotides (Fig. 4.1A). SHAPE reactivity reports local nucleotide flexibility because unconstrained nucleotides (like those in single-stranded structures) are more likely to sample relatively rare conformations that make the 20 -hydroxyl more nucleophilic and promote the reaction that forms a 20 -O-adduct. All four RNA nucleotides show nearly identical intrinsic reactivities when not constrained by base pairing or tertiary interactions (Wilkinson et al., 2009). In contrast, nucleotides constrained by base pairing or other interactions react poorly with SHAPE reagents. SHAPE chemistry is not strongly sensitive to solvent accessibility and, for example, conformationally dynamic but solvent inaccessible nucleotides are generally still reactive (Gherghe et al., 2008; Merino et al., 2005). SHAPE reagents both modify flexible sites in RNA and also undergo self-inactivating hydrolysis.

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Figure 4.1 Schemes for (A) RNA SHAPE chemistry and (B) RNA hydroxyl radical footprinting chemistries.

As a practical matter, this autoinactivation means that reagent reactivity does not need to be quenched, as long as the 20 -O-adduct forming reaction is allowed to proceed to completion (Merino et al., 2005). These bulky RNA adducts can be detected as stops to primer extension. The current gold standard for mapping solvent accessibility at the RNA backbone is hydroxyl radical footprinting. Short-lived hydroxyl radicals (OH) are generated in situ in a reaction between Fe2þ and H2O2 (Brenowitz et al., 2002; Latham and Cech, 1989; Shcherbakova and Brenowitz, 2008; Shcherbakova et al., 2008; Tullius and Greenbaum, 2005; Tullius et al., 1987). The iron ion is chelated by EDTA, which prevents the ion from binding directly to RNA. Hydroxyl radicals are then generated at the periphery of the RNA, leading to RNA cleavage (Fig. 4.1B). Cleavage occurs preferentially at RNA positions accessible to solvent, although other factors also contribute to reactivity (Balasubramanian et al., 1998; Lu et al., 1990). Backbone cleavage sites can also be detected by primer extension. Both SHAPE and the hydroxyl radical experiment can be converted to high-throughput formats by detecting sites of 20 -O-adduct formation or of backbone cleavage, respectively, by reverse transcriptase-mediated primer extension, resolved by capillary electrophoresis (Fig. 4.2A–D). In addition to the RNA modification reaction, a control no-reagent experiment is used to assess background. These reactions are compared to dideoxy sequencing

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Figure 4.2 High-throughput RNA structure mapping. (A) The RNA is folded and modified with a SHAPE reagent (1M7 or NMIA) or cleaved by hydroxyl radical chemistry. (B) RNA modifications or cleavages yield stops to primer extension, which are detected using color-coded, fluorescently labeled primers. (C, D) Resulting cDNAs are resolved by automated capillary electrophoresis. (E, F) After quantifying the net reactivity at each nucleotide position, the data are normalized on a scale spanning 0–2, where 1.0 is defined as the average intensity at highly reactive positions. (G) Changes in local nucleotide dynamics and solvent accessibility that distinguish any two RNA folding or RNP assembly states are readily detected at single nucleotide resolution by difference plot analysis.


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markers to assign nucleotide positions. After data processing, each nucleotide is characterized by a quantitative SHAPE reactivity or hydroxyl radical cleavage intensity that directly reflects the underlying RNA structure at single nucleotide resolution (Fig. 4.2E and F). In the SHAPE experiment, single-stranded nucleotides are reactive while base paired nucleotides are generally unreactive (Fig. 4.2E). Among other applications, SHAPE data can be converted to pseudo-free energy change terms to constrain the output of a thermodynamic structure prediction program. This blended approach generally produces highly accurate RNA secondary structure models (Deigan et al., 2009; Wilkinson et al., 2008). For the hydroxyl radical experiment, inspection of the final reactivity versus position histogram makes it possible to identify compact tertiary interactions as solvent inaccessible elements in the context of a large RNA (Fig. 4.2F).

3. Practice Both SHAPE and hydroxyl radical footprinting advantageously yield a comprehensive and single nucleotide resolution view of an RNA state in a concise set of experiments. In outline, both SHAPE and hydroxyl radical footprinting involve folding the RNA, adding protein (if appropriate), incubating the RNA with either the SHAPE electrophile or the hydroxyl radical reagents, recovering the RNA, and mapping the sites of modified or cleaved nucleotides by primer extension (Fig. 4.2). The following experimental overview is based on the analysis of the structure of the free bI3 group I intron RNA and its complexes with protein facilitators.

3.1. RNA folding SHAPE and hydroxyl radical footprinting experiments are most commonly performed with RNAs that have been generated by in vitro transcription (Duncan and Weeks, 2008; Latham and Cech, 1989; Merino et al., 2005; Tullius and Greenbaum, 2005; Wilkinson et al., 2006). These RNAs require purification by denaturing gel electrophoresis and then must be renatured to achieve a biologically relevant conformation. RNA folding reactions involving the renatured RNA can be initiated in several ways, including a change in ionic strength or addition of a protein or small molecule ligand. This protocol describes a general method in which an RNA is first heated and snap cooled in a low ionic strength buffer to eliminate multimeric forms. RNA folding is initiated by adding monoand divalent ions to this solution. The RNA is folded in a single step and then separated into (þ) and () reagent reactions for either SHAPE or hydroxyl radical footprinting.

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1. Add 4 pmol RNA in 10 ml sterile water to a 0.65 ml (Eppendorf ) reaction tube. Note: This amount of RNA is based on using 2 pmol per (þ) and () reagent reaction. The amount of RNA needed for complete analysis may vary with the RNA. The useful range is 0.5–8 pmol per reaction. 2. Heat the RNA to 95 C for 1 min. Place the RNA on ice for 1 min. 3. Add 24 ml 5 folding buffer [5 ¼ 200 mM MOPS (pH 8.0), 400 mM potassium acetate (KOAc) (pH 8.0), 100 mM MgCl2]. Adjust final volume to 108 ml with water. 4. Incubate at 37 C for 10 min. This is the ‘‘free RNA reaction mix.’’ Note: For protein-binding experiments, add proteins and incubate at appropriate concentrations, temperature, and time for the system. Adjustment of the volume of added water will be necessary to ensure a final 30 ml reaction volume in the chemical probing step.

3.2. Chemical probing of RNA secondary and tertiary structure Modification and cleavage reactions should take place under conditions that yield single hits over the length of RNA to be analyzed by primer extension ( 1 modification per 300 nt). For SHAPE, too high or low a concentration of reagent results in a steep signal decay in the fluorescence trace or in a low signal-to-noise ratio in the (þ) reagent lane, respectively. For hydroxyl radical cleavage, excessive cleavage yields a complex cleavage pattern that is biased toward short RNA fragments. For both reactions, fresh reagents are imperative. 3.2.1. SHAPE 1. Aliquot 3 ml 50 mM 1M7 (or NMIA) in DMSO and 3 ml neat DMSO into two 0.65 ml reaction tubes. These will be the (þ) and () reagent reactions. We strongly recommend 1M7 for routine SHAPE analysis of RNA. 1M7 can be synthesized as described (Mortimer and Weeks, 2007); alternatively, the NMIA reagent can be used. Note: The optimal concentration of 1M7 can vary with RNA length. The useful range is 0.5–8 mM; 5 mM final is a good starting concentration. For longer RNAs, or very AU rich RNAs, use the lower end of these 1M7 concentrations. 2. Add 27 ml of the free RNA reaction mix to 3 ml 1M7 in DMSO and an additional 27 ml to the tube containing 3 ml neat DMSO (final volume ¼ 30 ml).


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3. Allow reaction mixtures to react for five hydrolysis half lives. This time is 70 s for 1M7 and 35 min for NMIA (Merino et al., 2005; Mortimer and Weeks, 2007). No additional quench step is necessary. Note: For protein-binding experiments, remove proteins by proteolysis and phenol–chloroform extraction (Duncan and Weeks, 2008) before proceeding to step 4. 4. Adjust volume to 96 ml with water and recover RNA by ethanol precipitation: add 1 ml 3 M NaCl, 1 ml 20 mg/ml glycogen, 2 ml 0.5 M EDTA (pH 8.0), and 400 ml ethanol to both (þ) and () reactions, mix, and incubate at 80 C for 30 min. Sediment the RNA by spinning at maximum speed in a microfuge at 4 C for 30 min. 5. Redissolve RNA in 10 ml sterile water. 3.2.2. Hydroxyl radical footprinting Note: Fresh solutions are imperative for efficient cleavage reactions. Weigh out ammonium iron(II) sulfate hexahydrate [(NH4)2Fe(SO4)26H2O] and ascorbic acid (sodium salt) into dry reaction tubes; wait to make the solutions until just before addition to the RNA. Prepare a solution of 7.5 mM iron(II)/11.25 mM EDTA (pH 8.0). Prepare a 0.3% H2O2 solution in water. Prepare a 150 mM sodium ascorbate solution in water. Transfer two 27 ml aliquots of the free RNA reaction mix to two sterile 0.65 ml reaction tubes. One tube will be used for the (þ) OH reaction and the other for the () OH reaction. 5. For the (þ) OH reaction, sequentially add 1 ml each of the iron(II)–EDTA, hydrogen peroxide, and ascorbate solutions to the reaction solution. 6. Repeat step 5 for the () OH reaction, substituting 1 ml water for the iron(II)–EDTA solution. 7. Quench the reactions after 5 min at 37 C with 10.5 ml 75% glycerol (v/v).

1. 2. 3. 4.

Note: For protein-binding experiments, remove proteins by proteolysis and phenol–chloroform extraction (Duncan and Weeks, 2008) before proceeding to step 8. 8. Recover RNA as in steps 4–5 for the SHAPE reaction procedure.

3.3. Primer extension to map modification and cleavage sites Long RNA regions can be analyzed in a single experiment by performing each primer extension using a primer labeled with a color-coded fluorophore. The resulting cDNA products (from the (þ) and () reagent reactions plus one or two dideoxy sequencing reactions) are combined and resolved in one multifluor run by automated capillary electrophoresis.

76


In a single read, quantitative RNA structural information at single nucleotide resolution can be routinely obtained for 350–600 nt. Analysis is simplified by choosing fluorescent dyes with similar electrophoretic mobilities. The dyes 6-FAM, TET, HEX, and NED require very little correction for fluorophore-induced mobility shifts. This correction is readily performed using the ShapeFinder software (Vasa et al., 2008). Primer extension protocols are identical for RNA analyzed by SHAPE or by hydroxyl radical footprinting. 1. Add 3 ml of 0.3 mM fluorescently labeled primer to the RNA solutions from (þ) and () reagent reactions. 2. For sequencing reactions, add 3 ml 0.3 mM fluorescently labeled primer to 1–2 pmol of RNA in 8 ml sterile water. 3. For all reactions, anneal the primer to the RNA by heating at 65 C for 5 min, reducing the temperature to 37 C for 1 min, and placing on ice. 4. Add 6 ml of Superscript III enzyme mix (the solution is a 4:1:1 mixture of 5 first strand buffer (from Invitrogen), 100 mM DTT, and a solution that is 10 mM in each dNTP) to the (þ) and () reagent reactions and to the sequencing reactions. Also add 1 ml of 5 mM of a selected ddNTP to each sequencing reaction. Note: For AU rich RNAs, better extension can be achieved by decreasing the deoxynucleotide concentration used for sequencing. For example, if using ddATP to sequence at T, reduce the dATP concentration in the reverse transcriptase enzyme mix to 5 mM. 5. Add 1 ml SuperScript III Reverse Transcriptase (Invitrogen) to each tube. Mix well and incubate at 37 C for 5 min, 52 C for 20 min, 60 C for 5 min, and then place on ice. 6. Quench the primer extension reactions by adding 4 ml of a 1:1 mixture of 4 M NaOAc and 100 mM EDTA (pH 8.0) and place on ice. 7. Combine 22 ml from each (þ) and () reaction and one or two sequencing reactions into a 1.5 ml reaction tube and recover cDNA products by ethanol precipitation (240 ml 100% ethanol). Incubate at 80 C for 15 min. Sediment the cDNA by spinning at maximum speed in a microfuge at 4 C for 15 min. 8. Wash with 800 ml 70% ethanol to remove excess salt (which leads to poor resolution during capillary electrophoresis). 9. Repeat step 8 and dry pellet by vacuum for 10 min. 10. Resuspend in 10 ml deionized formamide.

3.4. cDNA analysis by capillary electrophoresis 1. Load each 10 ml sample into separate input wells on a capillary electrophoresis DNA sequencing instrument and run. 2. Export raw traces into ShapeFinder (Vasa et al., 2008).


77

3.5. Data analysis Data analysis using ShapeFinder has been outlined in detail elsewhere (Vasa et al., 2008; Wilkinson et al., 2008). Most tools are straightforward to use. The Align and Integrate tool in ShapeFinder performs a whole trace Gaussian integration to quantify the intensity of every peak in the (þ) and () reagent lanes. This tool works best for data in which traces have been adjusted to remove fluorescence background and in which peaks corresponding to the same positions have been aligned to overlap consistently. 1. Analyze SHAPE and hydroxyl radical data using ShapeFinder, as described (Duncan and Weeks, 2008; Wilkinson et al., 2008). 2. ShapeFinder is available at: http://bioinfo.unc.edu/Downloads/index. html. In brief, adjust the fluorescent baseline (window of 40 pixels), perform a mobility shift to account for the effect of different dyes on cDNA mobility, correct for signal decay, and scale the (þ) and () reagent traces to make them equal to each other in regions in which reagent-induced reactivities are low. Integrate all peaks in the (þ) and () reagent traces. 3. Subtract the () from the (þ) reagent reactivities to create a table of net reactivity as a function of nucleotide position.

3.6. Normalization A consistent approach to normalization is important for comparing data sets and for making accurate secondary structure predictions. We normalize SHAPE and hydroxyl radical data to a scale starting at zero (no reactivity) and in which 1.0 is defined as the average intensity of highly reactive positions. 1. Identify the (usually small number of) highly reactive outliers. 2. For large data sets (>300 measurements), use a model-free box plot analysis (Deigan et al., 2009). Reactivities greater than 1.5 times the interquartile range are outliers (Chernick and Friis, 2003). If the box plot approach is used for small RNA data sets (

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