Methods in Enzymology 469 - Biophysical, Chemical and Functional Probes of RNA Structure, Interactions and Folding: Interaction and Folding

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: 978-0-12-380922-3 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

Benjamin M. Akiyama Department of Molecular, Cell, and Developmental Biology, and Center for Molecular Biology of RNA, University of California, Santa Cruz, California, USA Nathan A. Baker Computational and Molecular Biophysics Program, and Department of Biochemistry and Molecular Biophysics; Center for Computational Biology, Washington University in St. Louis, St. Louis, Missouri, USA Robert T. Batey Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder, Colorado, USA Dana A. Baum Department of Chemistry, Saint Louis University, St. Louis, Missouri, USA Thu Betteridge Department of Biochemistry and Molecular Biology, Thomas Jefferson University, Philadelphia, Pennsylvania, USA Eric B. Brauns Department of Chemistry, University of Idaho, Moscow, Idaho, USA Pavol Cekan Department of Chemistry, Science Institute, University of Iceland, Reykjavik, Iceland Alan A. Chen Computational and Molecular Biophysics Program, and Center for Computational Biology, Washington University in St. Louis, St. Louis, Missouri, USA Shi-Jie Chen Department of Physics and Astronomy, and Department of Biochemistry, University of Missouri, Columbia, Missouri, USA Sebastian Doniach Department of Applied Physics, and Department of Physics, Stanford University, Stanford, California, USA David E. Draper Department of Biophysics, and Department of Chemistry, Johns Hopkins University, Baltimore, Maryland, USA xiii

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Contributors

R. Brian Dyer Department of Chemistry, Emory University, Atlanta, Georgia, USA Laura E. Easton MRC Laboratory of Molecular Biology, Cambridge, United Kingdom Kenneth D. Finkelstein Cornell High Energy Synchrotron Source, Cornell University, Ithaca, New York, USA Andrew D. Garst Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder, Colorado, USA Max Greenfeld Department of Chemical Engineering and Biochemistry, Stanford University, Stanford, California, USA Kathleen B. Hall Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, Missouri, USA Daniel Herschlag Departments of Biochemistry and Chemistry, Stanford University, Stanford, California, USA Ya-Ming Hou Department of Biochemistry and Molecular Biology, Thomas Jefferson University, Philadelphia, Pennsylvania, USA Amanda Y. Keel Department of Biochemistry and Molecular Genetics, and University of Colorado Denver, Aurora, Colorado, USA Jeffrey S. Kieft Howard Hughes Medical Institute, and Department of Biochemistry and Molecular Genetics, University of Colorado Denver, Aurora, Colorado, USA Eda Koculi Department of Biochemistry, Molecular Biology and Cell Biology, Northwestern University, Evanston, Illinois, USA Dominic Lambert Department of Chemistry, Johns Hopkins University, Baltimore, Maryland, USA Desirae Leipply Department of Biophysics, Johns Hopkins University, Baltimore, Maryland, USA

Contributors

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David M. J. Lilley Cancer Research UK Nucleic Acid Structure Research Group, MSI/WTB Complex, The University of Dundee, Dundee, United Kingdom Jan Lipfert Kavli Institute of Nanoscience, Delft University of Technology, Delft, The Netherlands Cuiping Liu Department of Biochemistry and Molecular Biology, Thomas Jefferson University, Philadelphia, Pennsylvania, USA Peter J. Lukavsky MRC Laboratory of Molecular Biology, Cambridge, United Kingdom Marcelo Marucho Department of Biochemistry and Molecular Biophysics, and Center for Computational Biology, Washington University in St. Louis, St. Louis, Missouri, USA Somdeb Mitra Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York, USA Suzette A. Pabit School of Applied and Engineering Physics, Cornell University, Ithaca, New York, USA Rohit V. Pappu Computational and Molecular Biophysics Program, and Center for Computational Biology; Department of Biomedical Engineering, Washington University in St. Louis, St. Louis, Missouri, USA Lois Pollack School of Applied and Engineering Physics, Cornell University, Ithaca, New York, USA Peter Z. Qin Department of Chemistry, University of Southern California, Los Angeles, California, USA Francis E. Reyes Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder, Colorado, USA Olav Schiemann Centre for Biomolecular Sciences, Centre of Magnetic Resonance, University of St Andrews, St Andrews, United Kingdom Xuesong Shi Department of Biochemistry, Stanford University, Stanford, California, USA

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Contributors

Snorri Th. Sigurdsson Department of Chemistry, Science Institute, University of Iceland, Reykjavik, Iceland Scott K. Silverman Department of Chemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois, USA Sergey Solomatin Department of Biochemistry, Stanford University, Stanford, California, USA Michael D. Stone Center for Molecular Biology of RNA, and Department of Chemistry and Biochemistry, University of California, Santa Cruz, California, USA Zhi-Jie Tan Department of Physics, Wuhan University, Wuhan, Hubei, China Sarah A. Woodson T.C. Jenkins Department of Biophysics, Johns Hopkins University, Baltimore, Maryland, USA Xiaojun Zhang Department of Chemistry, University of Southern California, Los Angeles, California, 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 modernday 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 xvii

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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

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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 VOLUME 469. Biophysical, Chemical, and Functional Probes of RNA Structure, Interactions and Folding: Part B Edited by DANIEL HERSCHLAG

C H A P T E R

O N E

Large-Scale Native Preparation of In Vitro Transcribed RNA Amanda Y. Keel,* Laura E. Easton,† Peter J. Lukavsky,† and Jeffrey S. Kieft*,‡ Contents 4 5 6 6 9 12 14 15 16 16 19 20 24 24

1. Introduction 2. Native Purification of RNA: Affinity Chromatography Method 2.1. Materials 2.2. Preparation of HMM protein 2.3. Preparation of DNA templates by PCR 2.4. Transcription and purification of RNA 3. Native Purification of RNA: Anion-Exchange Chromatography 3.1. Materials 3.2. Cloning of the plasmid DNA template 3.3. Cell culture and plasmid purification 3.4. In vitro RNA transcription 3.5. Weak anion-exchange FPLC Acknowledgments References

Abstract Biophysical studies of RNA require concentrated samples that are chemically and structurally homogeneous. Historically, the most widely used methods for preparing these samples involve in vitro transcription, denaturation of the RNA, purification based on size, and subsequent refolding. These methods are useful but are inherently slow and do not guarantee that the RNA is properly folded. Possible mis-folding is of particular concern with large, complexly folded RNAs. To address these problems, we have developed methods for purifying in vitro transcribed RNAs in their native, folded states. These methods also have the advantage of being rapid and readily scaled to virtually any size RNA or transcription amount. Two methods are presented: the first is an affinity * Department of Biochemistry and Molecular Genetics, University of Colorado Denver, Aurora, Colorado, USA MRC Laboratory of Molecular Biology, Cambridge, United Kingdom { Howard Hughes Medical Institute, University of Colorado Denver, Aurora, Colorado, USA {

Methods in Enzymology, Volume 469 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)69001-7

#

2009 Elsevier Inc. All rights reserved.

3

4

Amanda Y. Keel et al.

chromatography approach and the second is a weak ion-exchange chromatography approach. Both use equipment and materials readily available to almost any lab and hence should provide flexibility for those seeking alternate approaches to large-scale purification of RNA in the folded state.

1. Introduction Biophysical and structural studies of RNA often require concentrated, highly pure samples of a specific RNA molecule. Techniques such as X-ray crystallography or nuclear magnetic resonance (NMR) spectroscopy often demand a large amount of RNA, and thus obtaining suitable samples is often challenging and time-consuming. In addition, the RNA in these samples must not only be chemically pure, but structurally homogenous. Traditional methods to prepare large amounts of RNA have relied largely on in vitro transcriptions followed by denaturation of the RNA molecule and then purification based on size (Doudna, 1997; Milligan et al., 1987). Although useful, these methods require the RNA being refolded into its native structure before experiments are conducted. For many smaller, less structurally complex RNAs, this is accomplished readily. However, for large and more structurally complex RNAs, refolding into a structurally homogenous population can be difficult (Uhlenbeck, 1995). In addition, denaturing purification methods are inherently slow and time-consuming, and thus preparation of a pure sample is the rate-limiting step for many biophysical and structural studies (Doudna, 2000). In recent years, there has been considerable interest and experimentation toward developing higher throughput, nondenaturing methods for purifying large amounts of RNA for structural and biophysical studies (Batey and Kieft, 2007; Easton and Lukavsky, 2009; Kieft and Batey, 2004; Kim et al., 2007; Lukavsky and Puglisi, 2004; McKenna et al., 2007). Several useful methods have emerged and are now being employed and optimized. In some cases, these methods have resulted in RNA that subsequently has been used to solve high-resolution structures of RNA or RNA–protein complexes, demonstrating their utility (Batey et al., 2004; Lukavsky et al., 2003; Seif and Hallberg, 2009). Here, we present two methods that are used in our labs to purify RNA in a ‘‘native’’ or nondenatured state. Both methods use DNA templates for large-scale in vitro transcriptions, but differ in how the RNA is purified. The first method uses an affinity chromatography approach, the second an ionexchange chromatography approach. Both methods are designed to yield a highly pure, structurally homogeneous, concentrated RNA sample that can be taken directly to biophysical or structural studies. Also, both methods use standard laboratory reagents and equipment and thus should be accessible

5

Native Purification of RNAs

to most. Each method has advantages and disadvantages, and thus we leave it to the reader to assess which is most suitable for their application. We also emphasize that like all methods, it is important to optimize the protocol for each individual lab’s equipment and use.

2. Native Purification of RNA: Affinity Chromatography Method This method relies on the addition of a specific RNA sequence at the 30 -end of the RNA transcript that serves as an affinity tag by binding to a specific protein, which then binds to a column matrix. This tag is eliminated during purification (Fig. 1.1). The advantages of this method are that it is rapid, and many RNAs can be purified in parallel using a set of reusable, commercially available gravity-flow columns. The procedure can also be scaled down to make use of disposable ‘‘spin columns.’’ Furthermore, we have developed a ‘‘cloning-free’’ PCR-based method of generating DNA templates for in vitro transcription, which further increases the throughput of RNA purification. Disadvantages of this method include the requirement that the HMM protein be expressed and purified first. Additional information regarding this method can be found in other references (Batey and Kieft, 2007; Edwards et al., 2009). 1. DNA template T7 promoter RNA X 5¢

glmS ribozyme

MS2

3¢

MS2

3¢

2. RNA from in vitro transcription 5¢ RNA X

glmS ribozyme

3. Affinity immobilization 5¢ RNA X

glmS ribozyme MS2

MS2/ 3' MBP

6XHis NiNTA

4. GlcN6P cleavage and elution 5¢

RNA X

6XHis Ni+ 5¢ glmS ribozyme MS2 MS2/ 3' NTA MBP

5. Column regeneration 5¢ glmS ribozyme MS2 MS2/ 3' MBP

6XHis

NiNTA

Figure 1.1 Affinity purification scheme. ‘‘RNA X’’ denotes the desired target RNA to be purified; ‘‘6 His’’ indicates the hexahistidine tag that is added to the MS2/MBP fusion protein to yield protein HMM. Figure is adapted from Batey and Kieft (2007).

6


2.1. Materials LB-kanamycin agar plates LB-kanamycin media IPTG MilliQ water (ddH2O) and RNase free water Sodium phosphate (mono- and dibasic) NaCl MES Tween 20 detergent Imidazole Glycerol Supplies for SDS–PAGE (acrylamide, SDS, etc.) T7 RNA polymerase (6 His-tagged) Magnesium chloride HEPES KCl Ammonium sulfate Magnesium sulfate Triton X-100 Taq and Pfu polymerase 1.25 mM dNTP mix 1 M Tris–HCl, pH 8.1 at 37 C Spermidine Dithiothreitol (DTT) 100 mM stocks of each NTP mix pH 7.5 (Sigma) Inorganic pyrophosphatase (Sigma) Ni-NTA resin (Qiagen) Amicon spin concentrators Glucosamine-6 phosphate (Sigma) (GluN6P) Supplies for denaturing urea polyacrylamide acrylamide, etc.)

gels

(urea,

TBE,

2.2. Preparation of HMM protein The His-tagged MBP-MS2 coat fusion protein (HMM) used in this method was created to allow the protein to be immobilized on both Ni2þ or amylose affinity resins. HMM is a 59-kDa protein containing an N-terminal hexahistidine (6 His) tag, a central maltose-binding protein (MBP) domain, and a C-terminal MS2 coat protein containing the V29/dIFG mutations, which prevent protein multimerization and increase its affinity for RNA (Lim and Peabody, 1994). The protein is expressed in E. coli from plasmid pHMM, which confers kanamycin resistance, and purified using affinity chromatography.


7

2.2.1. Buffers and media Bacteria growth media: LB þ 10 mg/ml kanamycin Lysis buffer: 50 mM sodium phosphate, pH 8.0, 300 mM NaCl, 0.5% Tween 20, 10 mM imidazole, 10% glycerol Elution buffer: 50 mM sodium phosphate, pH 8.0, 300 mM NaCl, 0.5% Tween 20, 250 mM imidazole, 10% glycerol Sepharose column dialysis buffer: 25 mM Na–MES, pH 6.0, 25 mM NaCl Sepharose column elution buffers: 25 mM Na–MES, pH 6.0, with either 25 mM or 1 M NaCl Storage buffer: 25 mM Na–HEPES, pH 7.5, 200 mM NaCl, 10% glycerol 2.2.2. Procedures 1. Transform chemically competent Rosetta (BL21)/pLysS cells with the pHMM plasmid and plate on LB þ kanamycin agar plates, then incubate at 37 C overnight (or until colonies appear). Using a picked colony, begin a 50 ml ‘‘starter culture’’ with LB þ 10 mg/ml kanamycin and grow at 37 C with vigorous shaking (220 rpm) overnight. 2. Inoculate 1 l of LB þ 10 mg/ml kanamycin with 5 ml of starter culture and incubate at 37 C with vigorous shaking until the OD600 nm is at 0.6–0.7 AU. It is helpful to check the OD600 nm at least once an hour, and more often as the desired OD600 nm approaches. Once the desired OD600 nm is reached, induce expression by adding IPTG to 0.5 mM. Allow the cells to grow for 3 h at 37 C. 3. Harvest cells by centrifuging at 5000g for 15 min at 4 C. Remove the supernatant and resuspend the cell pellet in 50 ml of lysis buffer þ 500 ml of bacterial protease inhibitor cocktail (Sigma). At this point, the cells can be frozen and stored at 80 C. 4. Lyse the cells using a sonicator at 80% power using 15 s bursts with 45 s intervals between bursts. Repeat this five times, keeping the cells on ice. Transfer the lysate to centrifuge tubes and pellet the cell debris by centrifugation in a JA-20 rotor (Beckman) for 30 min at 35,000g. Immediately transfer the supernatant to a fresh beaker or tube. 5. Apply the supernatant onto a Ni-NTA affinity column (10 ml volume) that has been thoroughly equilibrated with 10 column volumes (CV) of lysis buffer. If desired, the flow-though can be saved and analyzed later by SDS–PAGE to verify protein binding to the column. 6. Wash the column with at least 10 CV of lysis buffer, then elute the protein in 10 ml fractions (1 CV each) of elution buffer. Collect at least 4 fractions, and then analyze each fraction using SDS–PAGE (12% acrylamide) to check for the presence of the protein. In general, the majority of the protein elutes in the first 3 fractions. 7. Pool the fractions containing the HMM protein and dialyze against Sepharose column dialysis buffer overnight at 4 C. Apply the protein

8


to a Hi-Prep 16/10 SP-Sepharose column (GE Healthcare) and wash the column with dialysis buffer until the A280 returns to baseline, then elute with a gradient of 0.025–1 M NaCl over a 200-ml volume (the protein elutes around 0.2–0.3 M NaCl). Collect fractions and use 12% SDS–PAGE to identify those containing HMM protein. 8. Pool fractions containing HMM and dialyze exhaustively against storage buffer at 4 C, then divide into 1 ml aliquots and store at 20 C. Determine concentration by measuring A280 and use a molar extinction coefficient of 83,310 M1 cm1. Typical yields (averaged over five individual preparations of the protein on the 2–4 l cell culture size) are 100–120 mg/l culture (Fig. 1.2). Notes and hints:

To avoid repeated transformations of competent cells and the need for a starter culture, a frozen cell stock can be created after step 1. To do this, combine 250 ml of the starter culture with 250 ml of storage media (LB þ kanamycine, 30% glycerol) and store at 80 C. This can be used to inoculate future 1 l cultures. 1

2

3

MW

120 100

*

80 70 60 50 40 30

20 15 10

Figure 1.2 Stained 4–12% SDS–PAGE of expression and purification of HMM protein. Lane 1 is the total soluble fraction from induced and harvested cells, lane 2 is the elution from the Ni-NTA column, and lane 3 is the sample after SP-sepharose purification. An asterisk indicates the HMM protein. Figure is reprinted with permission from Batey and Kieft (2007).

9


Because HMM contains an MBP domain, affinity chromatography using amylose resin also could be used as an alternate or additional purification step, although we have not done this.

2.3. Preparation of DNA templates by PCR In vitro transcriptions using T7 RNA polymerase are the standard way to generate large amounts of a specific RNA sequence (Milligan et al., 1987). DNA templates for this reaction can be generated in several ways. We have optimized a method to produce DNA templates by PCR that is used directly in transcription reactions to produce RNA that can be purified by affinity chromatography (Fig. 1.3) (Edwards et al., 2009). The advantage of this approach is that many DNA templates can be generated rapidly and in parallel, without the need for cloning. Our method uses two rounds of PCR comprising three reactions, which in the end yields a DNA template with a T7 promoter that encodes the desired RNA product, the activatable glmS ribozyme, and the MS2 hairpin affinity tag. In a later section, we present an alternate method to produce and use linearized DNA plasmids as templates in transcription reactions. 2.3.1. Reagents and buffers Universal DNA primers: 50 -GEN- 50 -GCGCGCGAATTCTAATACGACTCACTATAG-30 50 -GLMS- 50 -AGCGCCCGAACTACCGGT-30 30 -MS250 -CAGACCCTGATGGTGTCTGAA-30 0 3 -TAG- 50 -ACCGGTACCGGTAGTTCGGGCGCT-30 pRAV23 plasmid Primers A and B, specific to desired RNA product (Fig. 1.3) MW 1a 1b

2

First PCR step: Reaction 1a

Reaction 1b

T7 promoter RNA 5¢-GEN Primer A

glmS 5¢-GLMS

Primer B Second PCR step: (Reaction 2): 5¢-GEN 5¢ T7 promoter

glmS ribozyme

2 ´ MS2 3¢-MS2

3¢-TAG Joint

RNA

3¢-MS2

glmS ribozyme Cleavage site

2 ´ MS2

Figure 1.3 At left is a schematic of the PCR-based method of generating DNA templates for in vitro transcription. The three-reaction method yields the DNA template shown at the bottom. Transcription from this DNA results in the desired RNA product linked to the affinity purification tag. At right is an ethidium bromide stained agarose gel showing representative results from reactions 1a, 1b, and 2. Reprinted with permission from Edwards et al. (2009).

10


10 HEPES PCR buffer: 200 mM Na–HEPES, pH 8.8, 100 mM KCl, 100 mM ammonium sulfate, 20 mM magnesium sulfate, 0.1% Triton X-100 2.3.2. Procedures 1. PCR reaction 1a (50 ml): This reaction generates a DNA product that contains the sequence of the desired RNA under control of the T7 promoter, as well as a ‘‘joint’’ on the 30 -end that allows it to be combined with the product of reaction 1b (in reaction 2) to generate the full-length product. To generate this first product, two universal primers are combined with two primers specific for the desired RNA. These two specific primers are designed as partially overlapping DNA sequences that produce the final sequence (Fig. 1.3).

5 ml 10 HEPES PCR buffer 10 ml dNTP mix (1.25 mM each dNTP) 1 ml 50 -GEN primer (100 mM stock) 1 ml 30 -TAG primer (100 mM stock) 1 ml specific primer A (2 mM stock) 1 ml specific primer B (2 mM stock) 1 ml Pfu polymerase 30 ml ddH2O

Thermocycler protocol: Initial denaturing at 94 C for 5 min; 25 cycles of 94 C for 30 s, 55 C for 30 s, 68 C for 1 min; final extension 72 C for 7 min. 2. PCR reaction 1b (50 ml): This reaction generates a DNA product that contains the glmS ribozyme and the MS2 hairpin affinity tag, as well as a ‘‘joint’’ on the 50 -end that allows it to combine with the product of reaction 1a (in reaction 2) to generate full-length product. The reaction uses two universal primers and the pRAV23 plasmid as the template of the reaction. The product of this reaction can be used with any desired RNA sequence, and hence can be stored as a ‘‘stock’’ solution. 5 ml 10 HEPES PCR buffer 10 ml dNTP mix (1.25 mM each dNTP) 1 ml 50 -GLMS primer (100 mM stock) 1 ml 30 -MS2 primer (100 mM stock) 2 ml pRAV23 plasmid DNA, from a standard 5 ml ‘‘miniprep’’ diluted 1:10 1 ml Pfu polymerase 30 ml ddH2O

Thermocycler protocol: Initial denaturing at 94 C for 5 min; 25 cycles of 94 C for 30 s, 55 C for 30 s, 68 C for 1 min; final extension 72 C for 7 min.


11

3. PCR reaction 2 (1 ml): In this final reaction, the products from reactions 1a and 1b join via the ‘‘joint’’ sequences, and this joined DNA is amplified by two universal primers, resulting in the complete DNA template for in vitro transcription. In general, this reaction is done in very large scale, with at least 1 ml of reaction volume, in order to generate enough DNA template for a large-scale transcription.

100 ml 10 HEPES PCR buffer 200 ml dNTP mix (1.25 mM each dNTP) 20 ml 50 -GEN primer (100 mM stock) 20 ml 30 -MS2 primer (100 mM stock) 20 ml product from reaction 1a 1 ml product from reaction 1b 20 ml Taq or Pfu polymerase 619 ml ddH2O

Thermocycler protocol: Initial denaturing at 94 C for 5 min; 25 cycles of 94 C for 30 s, 55 C for 30 s, 68 C for 1 min; final extension 72 C for 7 min. After the thermocycling protocol is complete, 5–10 ml of the reaction is analyzed on an agarose gel with an appropriate size marker ladder to verify that a product of the desired length has been produced and to verify that the product is pure (Fig. 1.3). The desired DNA product will be 200 bp larger than the size of the final desired RNA product (i.e., for an RNA product of 100 nt, the size of the DNA template made by this method will be 300 bp). Notes and hints:

It is important that Tris-containing buffers not be used in these reactions, particularly reaction 2. The reason is that Tris will induce slow selfcleavage of the glmS ribozyme (Batey and Kieft, 2007; McCarthy et al., 2005; Roth et al., 2006). Small amounts of Tris introduced into the transcription reaction from the PCR reaction should be avoided and can be alleviated by using HEPES-based buffers. Note that most commercial PCR buffers contain Tris. Although agarose gel purification or commercial ‘‘PCR clean-up’’ kits can be used to process the DNA produced in these PCR reactions, we have found this to be unnecessary in most cases. However, if yields of DNA are very low, or substantial amounts of products other than the desired products are obtained from the reactions, purification of the DNA products may be necessary. Both Taq and Pfu polymerases are suitable for this protocol. In general, Taq gives more consistent and greater yields but can also introduce undetected mutations. Pfu is less error prone, but we find it to be more sensitive to specific DNA sequences and conditions.

12


2.4. Transcription and purification of RNA 2.4.1. Reagents and buffers 10 HEPES transcription buffer: 300 mM Na–HEPES, pH 8.0, 100 mM DTT, 20 mM spermidine, 0.1% Triton X-100 100 mM each NTP, pH adjusted to 7.4 with NaOH T7 RNA polymerase (10–15 mg/ml) DNA from PCR reaction 2 of Section 2.4 1 mM MgCl2 RNA column wash buffer: 50 mM K–HEPES, pH 7.5, 150 mM NaCl, 10 mM MgCl2, 10 mM imidazole RNA column regeneration buffer: 50 mM K–HEPES, pH 7.5, 150 mM NaCl, 10 mM MgCl2, 250 mM imidazole 2.4.2. Procedures 1. RNA is made by using the PCR-generated DNA template directly in an in vitro transcription. We generally conduct a transcription reaction of 5 ml final volume using the 1 ml PCR reaction 2 DNA from above, using the following conditions:

1 HEPES transcription buffer 32 mM MgCl2 4 mM each NTP T7 RNA polymerase to a final concentration of 50 mg/ml 1 unit/ml inorganic pyrophosphatase (optional) 1 ml DNA template from PCR reaction 2, above RNase-free water to a final volume of 5 ml

2. The reaction is assembled in 15 ml conical tubes, and then is incubated for 2–3 h at 37 C. If no inorganic pyrophosphatase is added, the solution will turn cloudy over time as pyrophosphate is released. If pyrophosphatase is not used, then at the completion of the reaction, this precipitate must be pelleted in a tabletop clinical centrifuge for 10 min and the supernatant immediately removed. 3. To prepare the transcription reaction for affinity purification, 1.6 mg of HMM protein is added directly to the reaction and allowed to incubate for 10 min on ice to allow the protein to bind to the MS2 hairpin affinity tag. This reaction is applied to a gravity flow column containing 1 ml Ni-NTA resin (QIAGEN) per 2 ml of transcription reaction at room temperature, and the solution passed through with a slow drip. To facilitate complete binding of the RNA/protein complex to the column, the flow-through can be passed through a second time. 4. Wash the column four times, each wash containing 4 CV of RNA column wash buffer, to remove excess protein, nucleotides and RNA abortive transcription products.

13


5. To elute the product RNA from the column, add 1 CV of RNA column wash buffer þ 1 mM glucosamine-6-phosphate (GlcN6P) to the column and allow this to pass through, then close the column valve and allow the column to sit for 10 min at room temperature. Apply a second CV of column wash buffer þ 1 mM GlcN6P, open the column valve, and collect this elution fraction. Collect two subsequent 1 CV elutions. Usually, most of the RNA comes out in the first two elution fractions (Fig. 1.4). 6. The HMM 30 -tag complex is removed from the column using regeneration buffer to regenerate the Ni-NTA resin. Add 5 CV and allow column to sit for 10 min. Drain column and repeat with another 5 CV of this buffer. Rinse the column with 4 CV RNA column wash buffer to prepare for another use. A

Tx FT W1 W2 W3 E1 E2

S a b c

B

Tx FT W1 W2 W3 E1 E2 E3 S

a b

c

Figure 1.4 Results from the affinity purification method. (A) Purification of a 94-nt RNA from a 100-ml transcription reaction, using a QIAGEN Ni-NTA spin column. Lane ‘‘Tx’’: the raw transcription; lane ‘‘FT’’: column flow-through; lanes ‘‘W1’’– ‘‘W3’’: column washes; lanes ‘‘E1’’ and ‘‘E2’’: column elutions; lane ‘‘s’’: imidazole regeneration. Bands ‘‘a’’, ‘‘b’’, and ‘‘c’’ indicate the full-length transcript, 30 -tag, and product RNA, respectively. (B) Purification of the same RNA as panel (A), but from a larger-scale (3.25 ml) reaction using 3 ml of Ni-NTA resin in a gravity flow column. Labeled identically to (A), but with a third elution step. Figure is reprinted with permission from Batey and Kieft (2007).

14


7. To determine which elution fractions contain RNA, a small amount of each fraction (20 ml) can be analyzed by denaturing PAGE and staining with ethidium bromide. Combine RNA-containing fractions and then concentrate the RNA and exchange into the desired buffer using Amicon spin concentrators of the appropriate molecular weight. In general, we concentrate to >10 mg/ml in RNase-free water and store the RNA at 20 C. Notes and hints:

It is highly recommended (especially the first few times this method is used) that fractions be saved from each step of the purification procedure and small aliquots of these fractions analyzed by denaturing PAGE. Problems with the quality of the HMM protein, the transcription yields, binding to the column, etc. can be detected and appropriate troubleshooting conducted. If binding of the RNA to the column appears to be weak, the above procedure can be adjusted. Transcription and addition of HMM protein are done as described above, but then the solution is passed over the NiNTA resin at 4 C and allowed to sit for 5–10 min; to ensure complete binding of the RNA/protein complex to the column, the flow-through can be passed through a second time. In the cold room, the column is washed four times with 4 CV of cold RNA column buffer. To cleave the product RNA from the tag and elute from the column, move the columns to room temperature then add 2 CV of room temperature RNA column buffer þ 1 mM GlcN6P. Seal the column well and place it on slushy ice for 10 min. Then remove the column from the ice and open it to collect the eluate (repeat). Two subsequent elutions with cold buffer were allowed to flow through the column to remove further RNA. Always make sure to thoroughly clean the columns after each use, and if columns are not to be used in the next few days, store them in 20–30% ethanol at 4 C. It is best to keep several Ni-NTA columns that are dedicated to RNA purification. Using these columns for protein purification can introduce many foreign proteins to the columns, potentially including RNases.

3. Native Purification of RNA: Anion-Exchange Chromatography Two other powerful methods for native RNA purification, one based on size-exclusion chromatography and the other using affinity purification, have been published in the past. The former employs separation of the different species in a transcription reaction (NTPs, small abortive transcripts, the transcribed RNA, and the plasmid DNA) based on size rather than charge (Kim et al., 2007; Lukavsky and Puglisi, 2004; McKenna et al., 2007).


15

Similar to our protocol based on anion-exchange chromatography, this method allows efficient recycling of unincorporated rNTPs and separation of mono- and multimeric RNA species, but still requires removal of the T7 polymerase from the reaction mixture using tedious phenol/chloroform extraction and desalting of the sample prior to chromatography, which takes about 1–2 h. In our protocol, the T7 polymerase is simply separated from the RNA during chromatography, which is more convenient and saves time. In the latter protocol, the desired RNA transcript is fused to an activatable ribozyme and an affinity tag for purification (Batey and Kieft, 2007; Kieft and Batey, 2004). This elegant method, which is described in detail in the first protocol, allows production of milligram amounts of native RNA with homogenous 30 -ends, which can be crucial for crystallographic applications. In contrast to size-exclusion or weak anion-exchange chromatography, no separation of mono- and oligomeric species generated during transcription is achieved using this protocol, but this can be easily achieved using either method as a subsequent purification step. For NMR spectroscopic application though, this method is less desirable, since a significant amount (>100 nt) of the transcript comprises the ribozyme and affinity tag resulting in a significantly lower final RNA yield, which in turn increases the cost of isotopically labeled RNA oligonucleotides. The next sections describe a detailed protocol for RNA preparation by in vitro transcription from linearized DNA plasmids and RNA purification using weak anion-exchange chromatography.

3.1. Materials PCR supermix (Invitrogen) or similar PCR reagents pUC18 plasmid Restriction enzymes—HindIII, BbsI, and EcoRI T4 DNA ligase DH5a cells TYE-ampicillin agar plates 2 TY-ampicillin or -carbenicillin media QIAfilter plasmid MEGA or GIGA kit 70% ethanol RNase free water 4.9 M magnesium chloride (Sigma) 25 mM NTP mix (Sigma) T7 RNA polymerase (6 His-tagged) 1 M Tris–HCl, pH 8.1 at 37 C 100 mM spermidine 500 mM dithiothreitol 10% Triton X-100 Inorganic pyrophosphatase (Sigma)

16


8% acrylamide 8 M urea gels 0.1% toluidine blue solution 500 mM EDTA, pH 8.1 at 37 C

3.2. Cloning of the plasmid DNA template In order to achieve a high yield of the plasmid DNA template coding for the RNA oligonucleotide of interest, a high copy number vector such as pUC18 should be used. The DNA construct should be engineered with a 50 HindIII restriction site preceded by a 4-nt overhang to allow efficient cleavage by the enzyme, the T7 promoter sequence (TAATACGACTCACTATA), the coding sequence of the desired RNA followed by two spacer nucleotides (usually TT), and finally BbsI and EcoRI restriction sites again with a 4-nt overhang for efficient enzyme cleavage as described (Lukavsky and Puglisi, 2004). The 2-nt linker between the RNA coding sequence and the BbsI site is necessary for cleavage 2 and 6 nucleotides upstream of the restriction site, thereby cutting directly at the last nucleotide position of the DNA template coding for the desired RNA sequence, which is necessary to perform ‘‘runoff’’ transcriptions. If there is an internal BbsI restriction site in the DNA template sequence, the BsaI restriction site can be used as an alternative, cutting 1 and 5 nucleotides upstream and therefore requiring only a 1-nt linker between DNA template sequence and restriction site. The DNA construct can be prepared using standard polymerase chain reaction (PCR) methods and for longer DNA constructs overlapping primers can be used as described elsewhere (Lukavsky and Puglisi, 2004). The resulting DNA fragment is then digested with HindIII and EcoRI, ligated into the pUC18 vector digested with the same enzymes, and the plasmid is transformed into DH5a cells, grown at 37 C and spread on a TYE-ampicillin agar plate. The plasmids from any resulting colonies should be sequenced using a primer further upstream of the T7 promoter (PLUHIII ¼ CTTCGCTATTACGCCAG) to be able to confirm the correct T7 promoter sequence, before beginning the preparation of milligram quantities of the plasmid required for in vitro RNA transcription.

3.3. Cell culture and plasmid purification 1. Streak a TYE-ampicillin agar plate with cells containing the correct plasmid DNA construct and grow at 37 C overnight. Inoculate 2 5 ml 2TYampicillin or -carbenicillin (50 mg/l) media with single colonies and grow in an orbital incubator at 37 C to 0.5 OD600. Inoculate 2 1 l 2TYampicillin or carbenicillin with 1 ml of the 0.5 OD600 culture and grow for 16–18 h in an orbital incubator at 37 C, harvest the cells by centrifugation and either store at 20 C immediately or begin the plasmid extraction and purification.


17

When growing the cells for the plasmid preparation, it is advised not to over-grow the culture as this leads to lower plasmid yield due to cell lysis which can be observed in the cell pellet as dark veins. The plasmid can be extracted and purified from the harvested cells using the following protocol based on QIAfilter plasmid MEGA and GIGA protocols. The volumes of buffers used should always correspond to the GIGA protocol. However, the lysate can also be loaded onto a QIAGEN-tip 2500 intended for the MEGA protocol, which is sufficient to purify up to 6 mg of pure plasmid DNA at a cheaper price than the GIGA QIAGEN-tip 10000. The washing procedures should be performed according to the GIGA protocol (300–400 ml buffer QC), but elution and precipitation volumes again correspond to the MEGA protocol. All procedures should be performed at room temperature, which should not exceed 25 C. 2. Add 125 ml chilled buffer P1 to the cells and gently resuspend using a 25 ml pipette to remove the cells from the wall of the tube and to remove any lumps of cells. Lyse the cells by adding 125 ml buffer P2, gently mix by inverting four to six times and incubate at room temperature for 5 min. Add 125 ml chilled buffer P3 and mix well by inverting four to six times to neutralize buffer P2 and stop lysis. Pour the lysate into a MEGA–GIGA filter cartridge and leave to settle for 5–10 min. 3. During this period equilibrate a QIAGEN-tip 2500 with 35 ml buffer QBT by gravity flow. 4. Filter the solution to clear the cell lysate from the cell debris. When the cell debris reaches the filter, switch off the vacuum, add 50 ml wash buffer FWB, and gently stir into the cell debris using a plastic pipette, then continue to filter until the cell debris strains against the filter, usually 350–400 ml. 5. Apply the lysate to the QIAGEN-tip 2500 by gravity flow, then wash with 6–8 QIAGEN-tip 2500 volumes of buffer QC. Place the QIAGEN-tip 2500 on a clean centrifuge tube and elute the DNA by adding 35 ml buffer QF. Once all the buffer has passed add 24.5 ml room temperature isopropanol to precipitate the DNA. 6. Centrifuge at 15,000g, 4 C for 15 min. Set a low brake so that the DNA pellet remains attached to the tube wall after the spin. Carefully decant the supernatant and wash the pellet with 7 ml 70% ethanol before centrifuging as before for 10 min, decant the supernatant and repeat the 70% ethanol wash. This step is especially important, as washing the DNA pellet twice with 70% ethanol reduces the salt content of the final plasmid DNA, which could interfere with the restriction enzyme digestion for plasmid linearization. 7. Decant the supernatant, pipette off any excess ethanol and air dry at room temperature for 1 h or at 4 C overnight.

18


The DNA pellet dries colorless, which might be difficult to see when trying to resuspend, therefore, it is useful to mark on the outside of the tube the position of the DNA pellet while it is still visible. 8. Add 2 ml RNase-free water to the pellet and allow softening of the DNA pellet at room temperature for 10 min before resuspending, then transfer into a 15-ml tube and rinse the centrifuge tube with another 1 ml of RNase-free water and add to the 2 ml resuspension. 9. Measure the concentration and dilute to 700 mg/ml by adjusting the volume with appropriate volumes of the recommended 10 BbsI buffer and RNase-free water. Linearize the plasmid by incubating with 50 U/ml BbsI at 37 C overnight. 10. Analyze the digestion by loading 10 ml of 7 mg/ml undigested and digested samples on a 1% agarose gel. The linearized plasmid now can be used directly to perform in vitro RNA transcription. Notes and hints:

The buffers (except P1 and P3) and QIAGEN-tips 2500 required for the large-scale plasmid purification should be stored at room temperature (15–25 C). Temperatures exceeding 25 C lead to cell debris passing through the filter which leads to cloudy lysates potentially congesting the QIAGEN-tip 2500. Lower storage temperature causes SDS precipitation in buffer P2, but this can be rectified by warming the buffer or by preparing fresh buffer according to the manufacturer’s instructions. Storing buffers for a long period of time can also lead to salt precipitation in buffers QC and QF and fresh buffers should be prepared according to manufacturer’s instructions. We also noticed that plasmids prepared with the QIAGEN MEGA kit at temperatures exceeding 25 C are poorly linearized and often partially degrade during the enzyme digest, probably due to higher salt concentration and nuclease contamination. It is also important to add the isopropanol to the completely eluted sample rather than allowing the sample to elute into the isopropanol, since the initially very high isopropanol concentration once again can lead to increased salt precipitation interfering with the subsequent plasmid linearization. Problems with plasmid linearization can be caused by several additional factors. The plasmid yield might be higher than measured or the activity of BbsI might be lower than anticipated and therefore the plasmid remains partially undigested after the overnight incubation period at 37 C. This can be resolved easily by addition of more restriction enzyme and continued incubation for a few hours and also by storage of BbsI at 80 C until needed. Another problem might be that the salt concentration of the plasmid DNA solution is too high, because the isopropanol used for precipitation was not at room temperature or both 70% ethanol


19

washes were not performed. If problems with incomplete and slow digestion occur repeatedly, small-scale trial digestions should be performed to determine the optimal salt concentration for the large-scale cleavage of the remaining plasmid.

3.4. In vitro RNA transcription It is desirable to maximize the RNA yield for each sample and this can be achieved by determining the optimal magnesium chloride concentration required for in vitro transcription. A series of 13 small-scale (25 ml) reactions with a range of magnesium chloride concentrations from 4 to 52 mM, increasing by 4 mM increments each, are typically used for magnesium chloride optimization. 1. Prepare the series of reactions containing 4 mM each NTP, 70 mg/ml linearized plasmid, 1200 U/ml T7 RNA polymerase, 40 mM Tris–HCl (pH 8.1 at 37 C), 1 mM spermidine, 5 mM dithiothreitol, 0.1% Triton X-100, 1 U/ml inorganic pyrophosphatase and 4–52 mM magnesium chloride and incubate in a 37 C water bath for 1 h. 2. Analyze the yield by loading 2 ml of each reaction on a denaturing PAGE (8% acrylamide, 8 M urea) and visualizing by UV shadowing or staining with 0.1% toluidine blue solution. A large-scale reaction to produce milligram quantities of RNA can now be performed in a 20 ml reaction using the optimal magnesium chloride concentration. 3. Prepare the reaction using the same conditions as the small-scale reactions and incubate in a 37 C water bath for 2–4 h before comparing the yield against the optimal small-scale reaction by loading 2 ml of each on a denaturing PAGE. 4. Stop the reaction by the adding 0.5 M EDTA (pH 8.1 at 37 C) to a final concentration of 50 mM. The transcription reaction can be stored frozen at 20 C or directly purified. Notes and hints:

Typically we use NTPs from Sigma, and most RNA transcriptions give the best yield in the range of 20–25 mM final magnesium chloride concentration. NTPs from other sources might require different concentrations and isotopically labeled NTPs prepared in-house following published methods (Batey et al., 1992) are usually obtained as magnesium salts and therefore much lower magnesium chloride concentrations are required (8–12 mM ) for optimal transcription yield. The inorganic pyrophosphate (PPi) which builds up during the transcription traps magnesium at a molar ratio of 2:1 of Mg:PPi and thereby

20


1

2

3

4

5

6

DNA RNA

NTPs

Figure 1.5 Addition of pyrophosphatase improves transcription yield. Denaturing PAGE analysis of large scale transcriptions in the absence (lanes 1–3) or presence (lanes 4–6) of 1 U/ml pyrophosphatase. The yield is analyzed by loading 2 ml of the transcription reaction after 45 (lanes 1 and 4) and 90 min (lanes 2, 3, 5, 6) without or with addition of magnesium after 45 min (compare lanes 2 and 3 with 5 and 6). RNA bands are visualized by staining with 0.1% toluidine blue. The best yield is obtained in the presence of pyrophosphatase without addition of magnesium (lane 5). Reprinted with permission from Easton and Lukavsky (2009).

inhibits the rate of transcription (Kern and Davis, 1997). In the past, we therefore added additional magnesium to the reaction after 45 min, which slightly improved the final yield (Fig. 1.5, compare lanes 3 and 5). Lanes 2 and 3 should be compared for +/- additional MgCl effect without inorganic pyrophosphatase. The better choice though is to perform the transcription in the presence of inorganic pyrophosphatase (1U/ml) without further addition of magnesium, which improves the yield by about 20–30% (Fig. 1.5, compare lanes 4 and 6). Prolonged incubation times (up to 4 h) of the in vitro RNA transcription reactions can help to improve the yield. However, the likelihood of hydrolysis will also increase.

3.5. Weak anion-exchange FPLC For fast and simple purification of RNA oligonucleotides from crude transcription reactions, we use an AKTA prime FPLC system equipped with a 50-ml superloop and three 5 ml HiTrap diethylaminoethyl (DEAE) sepharose FastFlow columns (GE Healthcare) connected in series. The DEAE columns are equilibrated with 3 CV of buffer A (50 mM sodium phosphate, pH 6.5, 150 mM sodium chloride, and 0.2 mM EDTA) at room temperature. Buffer B contains the same components with 2 M sodium chloride. Both buffers can be prepared in large quantities, sterile filtered and stored at 4 C (buffer A) or room temperature (buffer B) to avoid precipitation of sodium chloride.

21


1. Load the stopped transcription reaction into the 50 ml superloop and perform weak anion-exchange chromatography using the following gradient collecting 10 ml fractions: 0–70 ml 70–100 ml 100–380 ml 380–410 ml 410–455 ml 455–485 ml

0% B to 10% B to 30% B to 100% B 100% B to 0% B

1 ml/min 2 ml/min 2 ml/min 4 ml/min 4 ml/min 4 ml/min

2. The fractions are analyzed by denaturing PAGE (8% acrylamide, 8 M urea) loading 5 ml of each fraction. Unincorporated NTPs and the T7 polymerase usually elute in fractions 3–7 and small abortive oligonucleotides in fractions 8–12 (Fig. 1.3). RNA oligonucleotides elute depending on the overall phosphate charge per molecule starting with fractions 15–16 (30 nt, 400 mM NaCl) up to fraction 28 (500 nt, 570 mM NaCl), while the plasmid DNA template elutes later over several fractions (>630–700 mM NaCl). Therefore, a very shallow gradient is required especially for larger RNAs. 3. Pool the fractions corresponding to the clean RNA oligonucleotide and concentrate using 15 ml Centriprep centrifugal devices with 10 kDa MWCO-cutoff for RNAs larger than 30 nt or 3 kDa cutoff for smaller RNA oligonucleotides. Concentrate the RNA samples to 1 ml and then equilibrate them into the appropriate buffer using three to five consecutive 15 ml buffer exchanges. Concentrate the RNA to 1 ml at each step to maximize the efficiency of the buffer exchange (two spins) and then thoroughly mix with 14 ml fresh buffer for another round of centrifugation. The final RNA sample is best stored frozen at 20 C. Purification of RNA using weak anion-exchange chromatography relies on the difference in the overall phosphate charge per molecule. Therefore, unincorporated NTPs and small abortive transcripts bind only weakly to the DEAE matrix while RNA binds with medium affinity depending on the size and the plasmid DNA binds strongly and elutes as a broad peak towards the end of the shallow gradient (Fig. 1.6). Using this method, we routinely purify RNA samples ranging from 30 to 500 nt in length. For small RNAs (30–40 nt), we sometimes observe slight contamination with abortive transcripts, but they are removed easily during the buffer exchanges with Centriprep centrifugal devices. When purifying RNA samples larger than 500 nt, care has to be taken not to pool fractions contaminated with the linearized plasmid DNA and therefore only the utmost peak fractions (3–5 fractions) are pooled avoiding the long tail fractions, which are more likely to contain the linearized plasmid. In all cases, the RNA is

22


A A260 (O.D./ml) and fraction buffer B (%)

100 80 60

C

Small aborts

20

DNA

RNA

3

4

5

5

10

98 RNA

DNA

15 20 25 30 35 Fraction number

40

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

B

2

188

40

0

MW 1

NTPs

45

62 49 38 28 17 14 6 3

Figure 1.6 RNA purification using weak anion-exchange FPLC. (A) Elution profile of a 48-nt RNA obtained from a 20-ml in vitro transcription reaction using DEAE sepharose chromatography. Unincorporated NTPs, small abortive transcripts, and the plasmid DNA are well separated from the desired RNA product. (B) Denaturing PAGE analysis loading 5 ml of the eluted fractions. RNA and DNA bands are visualized by staining with 0.1% toluidine blue. The crude transcription reaction is shown in lane 1, NTPs and small abortive transcripts are loaded in lanes 2–8, the eluted RNA fractions are in lanes 9–12, and the eluted plasmid DNA is in lanes 13–15 visible as a very faint band in lanes 14 and 15. (C) Denaturing SDS–PAGE analysis of the eluted fractions shows that the purified RNA is free of T7 RNA polymerase. Resuspended pellets from TCA-precipitation of 1 ml of the crude transcription reaction (lane 2), the pooled flow-through (lane 3), abortive transcripts (lane 4), and the pooled RNA fractions (lane 5) are loaded. T7 RNA polymerase (lane 1) and a molecular weight marker are loaded as references and the molecular weight is indicated on the left. T7 RNA polymerase bands are visualized by coomassie-staining. Reprinted with permission from Easton and Lukavsky (2009).

free of the T7 RNA polymerase, which does not bind to DEAE sepharose matrix at salt concentrations higher than 100 mM (Fig. 1.6). This method also helps to reduce the preparation cost of isotopically labeled RNA samples for NMR spectroscopic studies, since the expensive unincorporated NTPs can be pooled, lyophilized, and desalted using boronate-affinity chromatography as described (Batey et al., 1992). An additional benefit of this purification method is that different RNA species can be separated, if the charge per molecule is significantly different, as for instance between mono- and multimeric RNA species. During purification of a 74-nt RNA, four-way junction from the

A260 (O.D./ml) and fraction buffer B (%)

A

B

100

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

DNA

80 60 40

RNA

NTPs Small aborts

20

RNA DNA

0

5

10

15

20 25 30 35 Fraction number

40

45

Figure 1.7 Separation of monomeric from oligomeric RNA species using weak anion-exchange FPLC. (A) Elution profile of an 80-nt RNA obtained from a 20-ml in vitro transcription reaction using DEAE sepharose chromatography. The RNA product elutes in two distinct peaks at lower and higher NaCl concentration. (B) Denaturing PAGE analysis loading 5 ml of the eluted fractions. RNA bands are visualized by staining with 0.1% toluidine blue. The crude transcription reaction is shown in lane 1, NTPs and small abortive transcripts in lanes 2–4, and the eluted RNA fractions corresponding to fractions 20–30 from Fig. 1.7A are loaded in lanes 5–15. While all the fractions contain the clean RNA product, only fractions 20–22 contain the monomeric RNA species (lanes 5–7). Other fractions (lanes 8–15) represent mixtures of mono-, di-, and tetrameric RNA species as judged by gel filtration (data not shown). Reprinted with permission from Easton and Lukavsky (2009).

24


classical swine fever virus (Pestova et al., 1998), we observed that the RNA eluted in two distinct peaks along the shallow salt gradient (Fig. 1.7). Analysis of the individual fractions by gel filtration revealed that the main peak contains the monomeric RNA species while the fractions eluted at higher salt concentration represent di- and tetrameric species, which could be separated efficiently from the desired monomer based on their overall charge difference (Fig. 1.7).

ACKNOWLEDGMENTS Development of native purification methods in the Kieft Lab has been supported by a grant from the Butcher Foundation, a grant from the University of Colorado Technology Transfer Office, and NIH grant R03 AI072187. This work in the Lukavsky Lab has been supported by MRC and a HFSP grant RGP0024/2008-C. JSK is a Howard Hughes Medicinal Institute Early Career Scientist.

REFERENCES Batey, R. T., and Kieft, J. S. (2007). Improved native affinity purification of RNA. RNA 13, 1384–1389. Batey, R. T., Inada, M., Kujawinski, E., Puglisi, J. D., and Williamson, J. R. (1992). Preparation of isotopically labeled ribonucleotides for multidimensional NMR spectroscopy of RNA. Nucleic Acids Res. 20, 4515–4523. Batey, R. T., Gilbert, S. D., and Montange, R. K. (2004). Structure of a natural guanineresponsive riboswitch complexed with the metabolite hypoxanthine. Nature 432, 411–415. Doudna, J. A. (1997). Preparation of homogeneous ribozyme RNA for crystallization. Methods Mol. Biol. 74, 365–370. Doudna, J. A. (2000). Structural genomics of RNA. Nat. Struct. Biol. 7(Suppl.), 954–956. Easton, L. E., and Lukavsky, P. J. (2009). Rapid, nondenaturing purification of RNA using weak anion-exchange FPLC. RNA (in press). Edwards, A. L., Garst, A. D., and Batey, R. T. (2009). Determining structures of RNA aptamers and riboswitches by X-ray crystallography. Methods Mol. Biol. 535, 135–163. Kern, J. A., and Davis, R. H. (1997). Application of solution equilibrium analysis to in vitro RNA transcription. Biotechnol. Prog. 13, 747–756. Kieft, J. S., and Batey, R. T. (2004). A general method for rapid and nondenaturing purification of RNAs. RNA 10, 988–995. Kim, I., McKenna, S. A., Viani Puglisi, E., and Puglisi, J. D. (2007). Rapid purification of RNAs using fast performance liquid chromatography (FPLC). RNA 13, 289–294. Lim, F., and Peabody, D. S. (1994). Mutations that increase the affinity of a translational repressor for RNA. Nucleic Acids Res. 22, 3748–3752. Lukavsky, P. J., and Puglisi, J. D. (2004). Large-scale preparation and purification of polyacrylamide-free RNA oligonucleotides. RNA 10, 889–893. Lukavsky, P. J., Kim, I., Otto, G. A., and Puglisi, J. D. (2003). Structure of HCV IRES domain II determined by NMR. Nat. Struct. Biol. 10, 1033–1038. McCarthy, T. J., Plog, M. A., Floy, S. A., Jansen, J. A., Soukup, J. K., and Soukup, G. A. (2005). Ligand requirements for glmS ribozyme self-cleavage. Chem. Biol. 12, 1221–1226.


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McKenna, S. A., Kim, I., Puglisi, E. V., Lindhout, D. A., Aitken, C. E., Marshall, R. A., and Puglisi, J. D. (2007). Purification and characterization of transcribed RNAs using gel filtration chromatography. Nat. Protoc. 2, 3270–3277. Milligan, J. F., Groebe, D. R., Witherell, G. W., and Uhlenbeck, O. C. (1987). Oligoribonucleotide synthesis using T7 RNA polymerase and synthetic DNA templates. Nucleic Acids Res. 15, 8783–8798. Pestova, T. V., Shatsky, I. N., Fletcher, S. P., Jackson, R. J., and Hellen, C. U. (1998). A prokaryotic-like mode of cytoplasmic eukaryotic ribosome binding to the initiation codon during internal translation initiation of hepatitis C and classical swine fever virus RNAs. Genes Dev. 12, 67–83. Roth, A., Nahvi, A., Lee, M., Jona, I., and Breaker, R. R. (2006). Characteristics of the glmS ribozyme suggest only structural roles for divalent metal ions. RNA 12, 607–619. Seif, E., and Hallberg, B. M. (2009). RNA-protein mutually induced fit: structure of Escherichia coli isopentenyl-tRNA transferase in complex with tRNA(Phe). J. Biol. Chem. 284, 6600–6604. Uhlenbeck, O. C. (1995). Keeping RNA happy. RNA 1, 4–6.

C H A P T E R

T W O

Assembly of Complex RNAs by Splinted Ligation Benjamin M. Akiyama*,† and Michael D. Stone†,‡ Contents 1. Introduction 2. General Considerations for Splinted RNA Ligation 3. Preparation of Unmodified RNA Ligation Precursor Molecules 3.1. In vitro transcription of telomerase RNA 3.2. Targeted RNase H cleavage of telomerase RNA 4. Preparation of Modified (Dye Labeled) RNA Ligation Precursor Molecules 4.1. Protocol 3: Dye-labeling and HPLC purification of synthetic RNA oligonucleotides 5. RNA Ligation Methods 5.1. Protocol 4: Splinted RNA ligation method for producing FRET-labeled telomerase RNA 6. Application: Single-Molecule FRET Measurements Acknowledgments References

28 29 31 31 33 37 39 40 42 44 45 45

Abstract Mechanistic studies of RNA enzymes (ribozymes) and ribonucleoprotein (RNP) complexes such as the ribosome and telomerase, often seek to characterize RNA structural features, either dynamic or static, and relate these properties to specific catalytic functions. Many experimental techniques that probe RNA structure–function relationships rely upon site-specific incorporation of chemically modified ribonucleotides into the RNA of interest, often in the form of chemical cross-linkers to probe for sites of protein–RNA interaction or small organic fluorophores to measure dynamic structural properties of RNAs. The ability to arbitrarily modify any RNA molecule has been greatly enabled by modern RNA synthesis techniques; however, there remains a practical size * { {

Department of Molecular, Cell, and Developmental Biology, University of California, Santa Cruz, California, USA Center for Molecular Biology of RNA, University of California, Santa Cruz, California, USA Department of Chemistry and Biochemistry, University of California, Santa Cruz, California, USA


#


27

28

Benjamin M. Akiyama and Michael D. Stone

limitation ( 70 bases). Consequently, experimental approaches involving specific chemical modifications of larger RNAs require the use of RNA ligation methods. The aim of this chapter is to describe a general approach for covalently joining multiple site-specifically modified RNA fragments, drawing from our fluorescence-based structural studies of telomerase RNA as an example.

1. Introduction The essential role of RNA as a cellular messenger of genetic information has been well appreciated since the early 1960s. However, recent discoveries in the fields of RNA structure, RNA-mediated catalysis, and ribonucleoproteins (RNP) have demonstrated a far more diverse set of physiological functions for RNA. For example, catalytic RNAs termed ribozymes, such as the group I self splicing intron from Tetrahymena, can speed up the rate of chemical reactions by many orders of magnitude (Cech, 1990). Functional contributions of RNA within the context of large multisubunit ribonucleoprotein complexes have also been demonstrated. One striking example comes from the recent tour de force determination of the high-resolution structure of the ribosome, which verified the central role of RNA during protein synthesis (Cate et al., 1999). Since the first demonstration of the catalytic properties of RNA, intense effort has been focused on the development of methods which probespecific structural characteristics of biological RNAs. Researchers have long utilized direct UV cross-linking methods to study RNA–protein interactions (Pinol-Roma et al., 1989); however, this approach can often be extremely inefficient. Advances in RNA synthesis techniques have opened new avenues for RNA research by facilitating the construction of RNA substrates possessing site-specific chemical modifications that may be used to probe the structure and conformational dynamics of RNA molecules (Scaringe et al., 1998). Some of the more common chemical modification strategies enlisted in structural studies of RNA include: (1) directed cleavage of RNA by site-specifically tethered EDTA–Fe (Han and Dervan, 1994); (2) the use of specific thiol moieties to form disulfide cross-links under nonreducing conditions (Cohen and Cech, 2001); (3) the incorporation of 4-thiouridine as an intrinsically UV-activated cross-linker (Yu, 2000); and (4) labeling with small organic fluorophores which provide a means to study dynamic structural properties of RNA (Hengesbach et al., 2008). Despite the obvious utility of using synthetic RNA in structural studies, there exists a practical size limit (70 bases) for RNA generated by routine chemical synthesis. Many biological RNA molecules of interest are much larger than this, leading to a general demand for robust methodologies that produce large RNA molecules harboring site-specific modifications.

Assembly of Complex RNAs by Splinted Ligation

29

Here, we describe an optimized method for generating site-specifically modified RNAs of arbitrary length. This method builds upon previously described RNA ligation techniques (Moore and Query, 2000), and includes several modifications to existing protocols that have substantially improved ligation efficiencies in our hands. The protocol described herein should provide experimentalists with large quantities of modified RNA substrates for a wide variety of applications. We demonstrate the utility of the method with a description of the design and fabrication of a 177-nucleotide multiply dye-labeled RNA construct derived from the Tetrahymena thermophila telomerase RNA. Our protocol for constructing telomerase RNA for fluorescence analysis requires a three part splinted RNA ligation reaction to covalently couple 50 - and 30 -terminal dye-labeled fragments onto an in vitro transcribed RNA insert fragment (Stone et al., 2007). In Section 6, we briefly present representative data from single molecule Fo¨rster resonance energy transfer (smFRET) experiments conducted on dye-labeled RNA molecules prepared by splinted RNA ligation.

2. General Considerations for Splinted RNA Ligation RNA ligation approaches typically employ one of several naturally occurring RNA ligase enzymes to catalyze the ATP-dependent joining of a ‘‘donor’’ fragment possessing a 50 -monophosphate and an ‘‘acceptor’’ fragment terminating in a 30 -hydroxyl group. All ligase enzymes produce products with a naturally occurring 30 –50 -phosphodiester bond at the ligation site. T4 RNA ligase was identified early on as a candidate enzyme for producing oligoribonucleotides of defined sequence, due to its activity on a wide variety of RNA substrates (England and Uhlenbeck, 1978; Walker et al., 1975). However, the strong preference of T4 RNA ligase for single-stranded substrates can produce a variety of unwanted side-products such as RNA circles and homodimers (see Section 5) (Romaniuk and Uhlenbeck, 1983). In contrast, splinted RNA ligation techniques have a fundamental advantage over T4 RNA ligase-catalyzed joining reactions, due to the dependence upon a properly annealed preligation complex, and therefore provide a powerful quality control step which biases the reaction products toward the desired RNA sequence and topology (Moore and Query, 2000; Moore and Sharp, 1992, 1993). For this reason, splinted RNA ligation has become the method of choice for generating large complex RNA molecules. The assembly of complex RNA molecules, harboring site-specific modifications, is a multistep procedure that can be subdivided into three stages (Fig. 2.1): (i) preparation of unmodified RNA ligation precursor fragments, (ii) preparation of modified RNA ligation precursor fragments, and

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Preparation of unmodified RNA ligation precursor fragments Clone telomerase RNA gene behind T7 RNA polymerase promoter

PCR amplify telomerase RNA gene with T7 RNA polymerase promoter

In vitro transcribe telomerase RNA using PCR amplicons as template

Preparation of modified RNA ligation precursor fragments Chemically synthesize RNA with 5-N-U modification at desired labeling site

Dye-label synthetic RNA fragments

Purify telomerase RNA transcripts with desalting column

Deprotect dye-labeled RNA fragments

Targeted RNase H digestion

PAGE purify dye-labeled RNA fragments

PAGE purify RNase H digestion products

HPLC purify dye-labeled RNA

RNA ligation Splinted RNA ligation of unmodified and dye-labeled RNA fragments

PAGE purify RNA ligation products

Verify by diagnostic PAGE and quantify by UV–vis spectrometry

Figure 2.1 General procedure for preparation of dye-labeled Telomerase RNA.

(iii) splinted RNA ligation and product purification. Efficient splinted RNA ligation requires that each precursor fragment possess the desired sequence and terminal functional groups. Therefore, purification procedures and


31

diagnostic analysis should be performed at each stage of the procedure. Of course, one pays a price for purity by taking losses at each purification step. Therefore, to ensure sufficient quantities of final product, each reaction step should be scaled up enough to account for the expected losses.

3. Preparation of Unmodified RNA Ligation Precursor Molecules There are two preferred methods for producing unmodified RNA ligation precursor fragments: chemical synthesis and in vitro transcription. For shorter RNA molecules, wherein ligation precursor fragments do not exceed 60–70 ribonucleotides, it is possible to construct complex RNA molecules entirely from chemically synthesized precursor fragments. However, while commercially available RNA synthesis has become more affordable, it still remains prohibitively expensive for some research groups. In these situations, as well as when RNA ligation precursor fragments must be greater than 70 nucleotides in length, in vitro transcription becomes the method of choice for producing large quantities of unmodified RNA ligation precursor molecules.

3.1. In vitro transcription of telomerase RNA Run-off transcription by one of several phage polymerases can readily produce milligram quantities of RNA fragments in excess of 1 kb in length (Fig. 2.2). The most common phage polymerase used for in vitro transcription is from the bacteriophage T7, which can be purchased commercially or overexpressed and purified from E. coli (Zawadzki and Gross, 1991). Detailed description of in vitro transcription techniques exist elsewhere (Wyatt et al., 1991); therefore, in this section we only highlight a few important points. Duplex DNA templates for T7 RNA polymerase reactions must possess the T7-specific promoter sequence, and can be generated either by linearization of plasmid DNA or via the polymerase chain reaction (PCR). We prefer to produce transcription templates by PCR with a highfidelity polymerase because it facilitates the production of RNA transcripts terminating at an arbitrary sequence. The efficiency of any given transcription reaction will vary significantly and should be optimized by titrating the concentration of Mg2þ, rNTPs, and DNA template. 3.1.1. Protocol 1: In vitro transcription of full-length telomerase RNA 3.1.1.1. Reagents

Phusion Hot Start High Fidelity Polymerase, New England Biolabs (Finnzymes) (F-540L) 5 Phusion HF reaction buffer (F-518)

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Telomerase RNA gene T7 RNA polymerase promoter

DNA plasmid

PCR amplify

In vitro transcribe

Purify on desalting column

Early fractions: Telomerase RNA

Late fractions: Free NTPs

Figure 2.2 Schematic diagram of in vitro transcription reaction. The telomerase RNA gene is cloned behind a T7 RNA promoter. PCR is used to generate linear DNA and the PCR amplicons are in vitro transcribed with T7 RNA polymerase. The RNA is purified away from free nucleotide on a desalting column.

Plasmid DNA PCR template in 10 mM Tris–HCl (pH 8) Custom DNA primers (100 mM ), Integrated DNA Technologies dNTP mix (40 mM stock), New England Biolabs (N0447L) Nuclease Free Water for PCR, Ambion (AM9932) Qiagen MiniElute PCR Purification Kit (28004) T7 RNA polymerase, New England Biolabs (M0251L) or laboratory purified T7 RNA polymerase reaction buffer (B9012S) (6 mM MgCl2, 10 mM dithiothreitol, 2 mM spermidine, 40 mM Tris–HCl, pH 7.9) rNTPs (80 mM), New England Biolabs (N0466S)


33

100% ethanol 3.3 M Na acetate (pH 5.2) PD10 desalting columns (for removing excess rNTPs after transcription reaction), GE Biosciences

3.1.1.2. Preparation of DNA template for transcription reaction by PCR Prepare PCR DNA template using standard plasmid purification techniques. In a total reaction volume of 500 ml prepare the following reaction mix: 50 ng template DNA, 1 mM forward PCR primer, 1 mM reverse PCR primer, 0.2 mM dNTP mix, 5 ml Taq Phusion enzyme in 1 Phusion HF buffer, adding the enzyme last. Aliquot into five PCR reaction tubes (100 ml/tube) and perform the PCR reaction in a thermocycler. 3.1.1.3. In vitro transcription Transcription reactions require 25 ng/ml linear DNA as a template for T7 RNA polymerase. In a total reaction volume of 500 ml, add 12.5 mg PCR template DNA, 0.5 mM NTP mix, and 50 ml T7 RNA polymerase in 1 T7 RNA polymerase buffer. During preparation the reaction mixture and enzyme should be kept on ice. Incubate the reaction at 37 C for 2 h and stop reaction with 10 ml 0.5 M EDTA. 3.1.1.4. Purification of telomerase RNA transcripts Due to the presence of short abortive transcripts within in vitro transcription reactions, it is typically advisable to gel purify desired RNA products by polyacrylamide gel electrophoresis (PAGE). However, the losses incurred during PAGE purification of large RNAs are significant (>50%); therefore, we try to minimize the number of gel purification steps during the construction of complex RNA molecules. Since full-length telomerase RNA transcripts will be further processed by targeted RNase H digestion (see below), we simply purify in vitro transcription reaction products by ethanol precipitation with 0.1 M NaOAc (pH 5.2), followed by a desalting column (PD-10, GE Biosciences) to remove residual unincorporated rNTPs.

3.2. Targeted RNase H cleavage of telomerase RNA Although extremely powerful, the use of in vitro transcription techniques for producing RNA ligation precursor fragments is limited by several intrinsic properties of phage RNA polymerases. First, the efficiency of the transcription reaction is very sensitive to the 50 -sequence of the nascent RNA transcript, typically requiring a guanosine at the þ1 position. Second, the propensity of T7 RNA polymerase to append nontemplated ribonucleotides onto the 30 -ends of nascent transcripts results in a heterogeneous distribution of transcript lengths (so called N þ 1 products). Several approaches have been employed to circumvent these shortcomings, with

34


the aim of producing RNA fragments of arbitrary sequence with homogeneous 50 - and 30 -termini (Fig. 2.3). The use of cis- and trans-acting ribozymes to remove 50 - and 30 -heterogeneities from large scale RNA preparations has been described (FerreD’Amare and Doudna, 1996; Price et al., 1995) (Fig. 2.3A). This method has the benefit of low cost and facile increase in scale, but suffers from being sequence sensitive and the laborious design/fabrication process for each construct to be produced. As many applications for site-specifically modified A

Telomerase RNA 5¢

Ribozyme cleavage Cis-ribozyme

DNAzyme cleavage

RNase H cleavage

DNAzyme

Chimeric 2¢-O-me RNA/DNA targeting oligo

Trans-ribozyme 3¢

5¢

3¢

5¢

5¢

3¢ 5¢

3¢

3¢

RNase H

Cleavage product RNA 5¢

3¢

B

C 3¢

2¢-O-me RNA DNA 2¢-O-me RNA 5¢

5¢

3¢

1

2

3

4

+

Site of Roche RNase H Site of NEB RNase H cleavage cleavage

Figure 2.3 Preparation of unmodified RNA ligation precursor fragments. (A) Schematic diagram of cleavage strategies for generating RNA ligation precursor fragments of specific length. Open triangles indicate desired site of RNA cleavage. (B) Cleavage site specificity of RNase H enzymes from different vendors. Enzyme obtained from Roche cleaves between the third and fourth base pair of the RNA–DNA duplex (read 50 –30 from the RNA). RNase H from New England Biolabs cleaves after the fourth base pair. (C) Denaturing PAGE gel of RNase H digestion products stained with ethidium bromide. Lane 1: No chimeric targeting oligonucleotide added. Lanes 2 and 3: Single chimeric targeting oligonucleotide added. Lane 4: Double cleavage directed by two chimeric targeting oligonucleotides.


35

RNA molecules require the production of several different RNA variants, we prefer more versatile approaches for postprocessing products of in vitro transcription reactions. Two alternatives to ribozyme-based approaches for site-specifically cleaving RNA transcripts are targeted cleavage by laboratory engineered DNAzymes (Breaker and Joyce, 1994; Silverman, 2008) or by ribonuclease H (RNase H) (Lapham and Crothers, 1996) (Fig. 2.3A). DNAzymes are oligodeoxyribonucleic acids that have been engineered to possess RNA hydrolyzing activity, with a cleavage specificity targeted by base pair complementarity between the DNAzyme and the RNA target. Although DNAzymes are both cost-effective and elegant, quantitative cleavage of target RNA substrates requires multiple annealing/cleavage cycles which can often result in detectable RNA degradation. For this reason, we rely upon targeted RNase H cleavage to generate large quantities of unmodified RNA ligation precursor fragments with homogeneous termini. RNase H is capable of cleaving RNA site-specifically using a 20 -Omethyl RNA/DNA chimeric oligonucleotide to direct the site of cleavage via Watson–Crick base pairing with the target RNA (Inoue et al., 1987). Thus, one can readily alter the cleavage specificity by simply redesigning a short synthetic 20 -O-methyl RNA/DNA chimeric oligonucleotide (Fig. 2.3A and C). Our general design for RNase H targeting oligonucleotides includes four deoxynucleotide bases flanked by ten 20 -O-methyl ribonucleotides on each side. RNase H can be purchased from a number of commercial sources. However, a cryptic variation in cleavage position specificity has been observed by our group and several others, depending on the enzyme source (Lapham et al., 1997). To circumvent this confusion in the past, we have processed all RNA ligation precursor fragments (unmodified and modified) by RNase H cleavage using a single source of enzyme. However, more recently we have mapped cleavage site preferences for two available sources of RNase H enzymes (Fig. 2.3B), eliminating the requirement for RNase H cleavage of chemically synthesized RNA ligation precursor fragments. The extent of site-specific RNA cleavage will vary with the efficiency of annealing between the chimeric RNase H targeting oligonucleotide and the RNA substrate. Sites with a high degree of secondary structure are typically more difficult to cleave to completion; however, in our experience slight modifications in the position of the cleavage site, length of the chimeric RNase H targeting oligonucleotide or salt concentration during the annealing step usually results in greater than 80% cleavage. Furthermore, we have found that once an efficient RNase H cleavage site has been determined, the efficiency of subsequent splinted RNA ligation at this site is also high. To construct dye-labeled telomerase RNA molecules for our FRET measurements, we first prepare an 89 nucleotide long RNA insert fragment by

36


targeting RNase H cleavage to two different sites within full-length telomerase RNA produced by in vitro transcription as described above. 3.2.1. Protocol 2: Targeted RNase H cleavage of unmodified RNA ligation precursor fragment 3.2.1.1. Reagents

Ribonuclease H, New England Biolabs (M0297L) RNase H reaction buffer, New England Biolabs (B0297S) (75 mM KCl, 3 mM MgCl2,10 mM dithiothreitol, 50 mM Tris–HCl, pH 8.3) Custom synthetic chimeric 20 -O-methyl RNA/DNA oligonucleotides, Integrated DNA Technologies, Inc. Full-length telomerase RNA prepared by in vitro transcription (see above) RNasin Plus RNase inhibitor, Promega (N2615) Urea loading buffer (8 M urea, 25 mM EDTA, 5 mM Tris, pH 8.0) Fluor-Coated TLC Plates, Ambion (AM10110) KONTES RNase-Free Pellet Pestle, VWR (749521-1590)

3.2.1.2. Annealing chimeric RNase H targeting oligos and telomerase RNA Set up the annealing reaction with 100 mg of RNA from the previous transcription reaction. Add to this a fivefold molar excess of your custom synthetic chimeric 20 -O-methyl RNA/DNA oligonucleotides. Bring up to a total volume of 250 ml with nuclease-free H2O. Denature the reaction for 4 min at 90 C. Incubate for 10 min at 37 C to allow the chimeric oligonucleotides to anneal to the RNA. 3.2.1.3. RNase H cutting reaction In a separate tube on ice add 50 ml 10 RNase H buffer, 12.5 ml RNasin Plus RNase inhibitor, and 50 ml RNase H enzyme to 137.5 ml nuclease-free H2O. Add this to the annealing reaction. Incubate at 37 C for 2 h. Stop the reaction with 1/20th volume 0.5 M EDTA, pH 8.0. 3.2.1.4. Purification of RNase H cleavage products Ethanol precipitate the RNA and resuspend the pellet in 1 urea loading buffer. Run on an 8% denaturing PAGE gel. Transfer the gel to a sheet of saran wrap and place over a fluorescent TLC plate. Excise the band containing the desired product by UV shadowing and place in a 15-ml conical tube. Add 5 ml gel elution buffer (10 mM Tris–HCl, pH 8.0 þ 100 ml phenol:chloroform: isoamyl Alcohol) and crush the excised gel slice with an RNase-free pellet pestle or a pipet tip. Agitate in a shaker or tape to a vortexer left on overnight. Spin down tube and transfer as much solution as possible to a new tube, avoiding the crushed pieces of acrylamide. Butanol extract the volume down to about 200 ml. Ethanol precipitate the RNA and save the pellet.


37

4. Preparation of Modified (Dye Labeled) RNA Ligation Precursor Molecules In our laboratory, we have primarily utilized splinted RNA ligation methods to generate fluorescently labeled telomerase RNA molecules (Fig. 2.4A); however, the method outlined in this chapter is applicable to the construction of large RNA molecules for use in a variety of RNA structure probing experiments. In principle, any base modification that can be introduced through chemical synthesis can be site-specifically incorporated into a larger RNA construct via RNA ligation approaches. If it is necessary to perform further chemical treatment of modified synthetic RNA fragments (i.e., dye labeling) it is desirable to purify away the fraction of RNA that was not properly functionalized. This can often be done using a HPLC system with the appropriate column. Once purified, the modified synthetic RNA fragments may be ligated to in vitro transcribed RNA to generate full-length RNAs with one or more site-specific labels. For the purposes of this chapter, we describe the labeling of telomerase RNA with two dye labels, Cy3 and Cy5, for use in single-molecule FRET experiments. To generate RNA ligation precursor fragments harboring site-specific dye modifications, synthetic RNAs are purchased possessing a 5-amino-allyluridine base at the desired position. Under basic conditions, monoreactive Cy3 and Cy5 dyes (GE Life Sciences) containing the dye conjugated to an N-hydroxysuccinimide-activated carboxylic acid, will react with amines and amides to covalently attach the RNA to the dye. As the dye packs are reactive with primary amines, buffers that contain amines such as Tris will quench the coupling reaction and should therefore be avoided. It is important to note that any synthetic oligoribonucleotide that will serve as a ‘‘donor’’ during the ligation step must have a 50 -monophosphate introduced, which may be directly incorporated during synthesis or added enzymatically using T4 polynucleotide kinase. We have found it unnecessary to PAGE purify synthetic RNA fragments prior to dye-labeling, as there will be a subsequent PAGE purification step. Furthermore, to ensure stability of the RNA during the labeling reaction, we perform the dye coupling reaction with protected RNA, and deprotect after the RNA is labeled. In our experience, shorter synthetic RNA oligonucleotides with a minimum of predicted secondary structure have a higher labeling efficiency. The length of synthetic RNA ligation precursors will typically also be dictated by other factors such as the size of the target RNA and the desired position of the modification. Lastly, ligation efficiencies will vary significantly from one position to another; therefore, it may be necessary to extend the length of a synthetic fragment in order to improve yields. The removal of unmodified RNA fragments following the dye-labeling step is accomplished by reverse-phase high-pressure liquid chromatography (HPLC) (Fig. 2.4B) (Walter, 2003). Reversed-phase HPLC uses a column

38


B Absorbance (A.U.)

A Chemically synthesize RNA with 5-N-U modification at desired labeling site

Dye-label 5-N-U labeled RNA with amino-reactive Cy3 or Cy5

EtOH precipitate to remove excess dye

1500

Cy5-RNA

1000

ABS650

500 0

ABS260

8

10

12

14

16

18

mL

C Cy5-RNA

Cy3-RNA

Deprotect dye-labeled RNA Cy5 EtOH precipitate Cy3

PAGE purify and gel extract

Butanol extract, EtOH precipitate, and resuspend in HPLC buffer A (100 mM TEAA, pH 7.0)

HPLC purify dye-labeled RNA by reverse phase chromatography

Figure 2.4 Preparation of modified (dye labeled) RNA ligation precursor fragments. (A) General protocol for dye-labeling synthetic RNA fragments. (B) HPLC trace of purification for a Cy5-labeled RNA oligonucleotide. Unlabeled synthetic telomerase RNA fragment elutes 10 ml (ABS260 peak) and Cy5-labeled RNA elutes at 15 ml (overlapping ABS260 and ABS650 peaks) (C) Diagnostic denaturing PAGE gel of HPLCpurified Cy5-RNA and Cy3-RNA fragments imaged on Typhoon scanner (GE Life Sciences).

with a hydrophobic stationary phase and an aqueous mobile phase. As the mobile phase is gradually combined with a nonpolar solvent, molecules are separated by their polarity in the column with hydrophobic molecules eluting later in the gradient. Since Cy5 and Cy3 add a large hydrophobic surface to the RNA, reversed-phase HPLC is ideal for separating labeled from unlabeled RNA.


39

4.1. Protocol 3: Dye-labeling and HPLC purification of synthetic RNA oligonucleotides 4.1.1. Reagents

3.3 M NaOAc, pH 5.2 Sodium bicarbonate Custom synthesized RNA oligonucleotides, Thermo Scientific/ Dharmacon Cy5 and Cy3 monoreactive dye packs, GE Healthcare (PA25001 and PA23001) 100% ethanol RNA deprotection buffer, Thermo Scientific/Dharmacon (B-001000DP-018)—100 mM acetic acid, adjusted to pH 3.8 with TEMED KONTES RNase-Free Pellet Pestle, VWR (749521-1590) Urea loading buffer (8 M urea, 25 mM EDTA, 5 mM Tris, pH 8.0) 100 mM triethylammonium acetate (TEAA), pH 7.0 100% acetonitrile Eclipse XDB-C8 HPLC column, Agilent (990967-906) AKTA purifier

4.1.2. Dye-labeling reaction Resuspend 200–400 mg synthetic RNA in 200 ml 10 mM Tris, pH 8.0, and ethanol precipitate synthetic RNA with 0.1 M NaOAc, pH 5.2. Resuspend the pellet in 200 ml 0.1 M sodium bicarbonate solution. Add solution to Cy5 or Cy3 monoreactive dye pack tube. Incubate for 1 h at 37 C and ethanol precipitate. Resuspend dye-labeled RNA pellets in 200 ml deprotection buffer. Incubate at 60 C for 30 min and ethanol precipitate. Resuspend pellet in 100 ml 1 urea loading buffer containing no loading dye. Run on an 8% denaturing PAGE gel. Excise the cyan- or pink-colored band, avoiding any smears below, and transfer to a 15 ml conical tube. Crush the gel slice in 5 ml TE using a pipet tip or RNase-free pellet pestle. Agitate in a shaker or tape to a vortexer left on overnight. Spin down tube and transfer as much solution as possible to a new tube, avoiding the crushed pieces of acrylamide. Butanol extract volume down to about 200 ml. Ethanol precipitate RNA and save pellet. 4.1.3. HPLC purification of Dye-Labeled RNA Equilibrate column in 100% acetonitrile until pressure stabilizes ( 20 ml acetonitrile). Run a gradient to 100% 100 mM TEAA, pH 7.0 over 5 min and stay at 100% 100 mM TEAA. Wait for pressure to stabilize. Resuspend labeled RNA pellets from previous PAGE purification in 60 ml 100 mM TEAA, pH 7.0. Load sample into a 100 ml loop, and inject. Set a gradient to 100% acetonitrile over 35 column volumes. Begin collecting fractions when

40


the UV 260 absorbance begins to climb. The unlabeled RNA should elute first (Fig. 2.4B), whereas the second large peak should contain the labeled RNA with additional absorbance at 650 nm for Cy5 or 550 nm for Cy3. If the HPLC model does not have a built-in UV–vis detector, a spectrophotometer can be used to identify dye-labeled fractions. Dye-labeled RNA-containing fractions should be pooled and butanol extracted to a volume of 200 ml. They should then be ethanol precipitated and the pellet should be stored at 20 C for ligation to the in vitro transcribed precursor RNA fragment. RNA labeling and purification may be confirmed by UV spectrophotometry and diagnostic PAGE analysis (Fig. 2.4C).

5. RNA Ligation Methods In our laboratory, we have used three different enzymes to catalyze the ATP-dependent ligation of a ‘‘donor’’ RNA fragment possessing a 50 monophosphate to an ‘‘acceptor’’ RNA fragment having a 30 -hydroxyl group: T4 RNA ligase, T4 DNA ligase, and T4 RNA ligase 2 (Fig. 2.5A). The robust ability of T4 RNA ligase to covalently join individual oligoribonucleotides makes this enzyme an obvious choice for the assembly of complex RNA molecules (England and Uhlenbeck, 1978; Walker et al., 1975). However, the utility of T4 RNA ligase for this purpose is limited by its strong preference for single-stranded RNA fragments, which can often lead to unwanted side products including RNA circles and dimers (Fig. 2.5A, left) (Romaniuk and Uhlenbeck, 1983). To circumvent this shortcoming, researchers more typically employ T4 DNA ligase, which can covalently couple two RNA fragments when annealed to a complementary DNA splint (Fig. 2.5A, right) (Moore and Query, 2000; Moore and Sharp, 1992). Splinted RNA ligations have several important advantages over T4 RNA ligase-catalyzed joining reactions. First, the requirement for a complementary DNA splint virtually eliminates formation of unwanted side products, making possible more complex multisite ligation reactions. Second, the precise geometry required to promote T4 DNA ligase-mediated ligation imposes strict constraints upon putative ligation complexes and provides an important level of quality control which bypasses any residual heterogeneity in the lengths of RNA ligation precursor fragments. Despite these clear advantages, the efficiency of splinted RNA ligation reactions catalyzed by T4 DNA ligase is typically limited by the amount of enzyme one can introduce into the reaction. This problem is derived from the poor turnover of T4 DNA ligase on DNA/RNA heteroduplexes. More recently, a variant of T4 RNA ligase, called T4 RNA ligase 2 (Nandakumar et al., 2004), has been identified. This enzyme has a pronounced preference

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A

RNA ligation precursor fragments Cy5-labeled RNA fragment

Unmodified RNA fragment

Cy5

Cy3-labeled RNA fragment Cy3

T4 RNA ligase

DNA-splinted RNA ligation

Target product

DNA splint

DNA splint T4 DNA ligase or T4 RNA ligase 2

+ Side products RNA circles Dimers

B

No ligase

T4 RNA T4 DNA ligase 2 ligase

* 1

2

3

4

5

6

Figure 2.5 Strategies for RNA ligation. (A) (left) T4 RNA ligase joins single-stranded RNA and can generate undesired side products such as RNA circles and dimers. (Right) T4 DNA ligase and T4 RNA ligase 2 prefer double-stranded nucleic acid. When directed by a DNA splint, they show greatly reduced side product formation. (B) T4 RNA ligase 2 is highly active. Lanes 1 and 2 are the same gel lane containing all RNA ligation precursor fragments but no ligase enzyme. Lane 1: Typhoon scanner image of ethidium bromide staining of in vitro transcribed telomerase RNA precursor fragments (asterisk indicates cross-excitation of Cy3-RNA fragment in Typhoon scanner). Lanes 3–6 are a titration of commercially available high-concentration preparations of T4 RNA ligase 2 and T4 DNA ligase imaged on Typhoon scanner. Cy3-RNA- and Cy5RNA-labeled fragments are present in twofold excess over in vitro transcribed unmodified RNA fragment. Lane 3: 1% (v/v) T4 RNA ligase 2. Lane 4: 10% (v/v) T4 RNA ligase 2. Lane 5: 1% (v/v) T4 DNA ligase. Lane 6: 10% (v/v) T4 DNA ligase.

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for joining RNA ends within a DNA/RNA duplex, and turns over products much faster than T4 DNA ligase. Thus, when using T4 RNA ligase 2 one can use far less enzyme to achieve comparable levels of RNA ligation to that of T4 DNA ligase-catalyzed reactions, making this approach significantly more cost-effective (Fig. 2.5B). For these reasons, we typically use T4 RNA ligase 2 for our splinted RNA ligation reactions. However, it is important to note that T4 RNA ligase 2 retains a detectable level of activity on single-stranded RNA fragments, and thus produces more side products than T4 DNA ligase. These side products can usually be eliminated by gel purification of the desired RNA ligation products. However, for certain joining reactions we have found it necessary to use the less efficient T4 DNA ligase enzyme to suppress unwanted reaction products. Before performing a large scale RNA ligation reaction, we recommend setting up a diagnostic reaction in a 20-ml reaction volume. Once the ligation is determined to be efficient, the reaction can then be scaled up linearly. When designing DNA splints, several factors should be considered. We find that DNA splints that are 25–30 bases long are typically sufficient to form a stable preligation complex. If possible, the site of ligation should be chosen so as to avoid stable secondary structure within the DNA splint strand. If annealing of the RNA ligation precursor fragments to the DNA splint is found to be inefficient, the length of the DNA splint may be increased. In addition, two ligation sites that are sufficiently close to each other may be annealed to a single DNA splint. The relative concentrations of the DNA splint and RNA precursor fragments will have a significant impact on the efficiency of preligation complex formation. While it is possible to ‘‘drive’’ the annealing reaction by increasing the amount of one of the RNA ligation precursor fragments, it is important that the molar concentration of DNA splint not exceed the concentration of the most abundant RNA ligation precursor fragment. The presence of a molar excess of DNA splint will inhibit the reaction by titrating away RNA ligation precursor fragments.

5.1. Protocol 4: Splinted RNA ligation method for producing FRET-labeled telomerase RNA 5.1.1. Reagents

T4 DNA ligase, New England Biolabs (M0202M) T4 RNA ligase 2, New England Biolabs (M0239L) T4 DNA ligase buffer, New England Biolabs (B0202S) (10 mM MgCl2,10 mM dithiothreitol, 1 mM ATP, 50 mM Tris–HCl, pH 7.5) T4 RNA ligase 2 buffer New England Biolabs (B0239S) (2 mM MgCl2, 1 mM DTT, 400 mM ATP, 50 mM Tris–HCl, pH 7.5)


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Unmodified RNA ligation precursor fragment (see Section 3) Dye-labeled RNA ligation precursor fragment (see Section 4) Custom DNA ‘‘splint’’ oligos, Integrated DNA Technologies, Inc. RNasin Plus RNase Inhibitor, Promega (N2615) Urea Loading buffer (8 M urea, 25 mM EDTA, 5 mM Tris, pH 8.0) KONTES RNase-Free Pellet Pestle, VWR (749521-1590) 25:24:1 phenol:chloroform:isoamyl alcohol Chloroform Glycogen for Molecular Biology, Roche (10901393001)

5.1.2. Annealing of DNA Splints to precursor RNA fragments The conditions listed in this protocol are for a diagnostic scale reaction. Once successful, these same conditions may be scaled up linearly to produce larger quantities of RNA ligation products. In a total reaction volume of 10 ml, add 20 pmol of the unmodified RNA ligation precursor fragment (from Section 3), 40 pmol Cy5-labeled RNA ligation precursor fragment (from Section 4), 40 pmol Cy3-laveled RNA ligation precursor fragment (from Section 4), and 40 pmol each of the corresponding DNA splint oligos. Denature at 90 C for 3 min, and anneal at 30 C for 10 min. 5.1.3. Ligation reaction In a separate tube, create a 10-ml mixture containing 2 ml T4 DNA ligase or T4 RNA ligase 2 and 1 ml RNasin Plus RNase inhibitor in 1 concentration T4 DNA ligase or T4 RNA ligase 2 buffer. Add this mixture to the annealing reaction above. Incubate at 30 C for 2 h. Bring to 200 ml with 10 mM Tris, pH 8.0, phenol–chloroform extract, and ethanol precipitate, adding 1 ml glycogen to help visualize the pellet. 5.1.4. PAGE purification Resuspend the pellet in 20 ml urea loading buffer and run on an 8% denaturing PAGE gel. Cover the gel while it runs to avoid photobleaching of the dyes. Excise the purple band on the gel corresponding to the Cy5 and Cy3 double-labeled RNA. Confirm the success of your excision using the fluorescence setting on a Typhoon gel scanner. Crush the excised band in 1 ml 10 mM TE pH 8.0 using a pipet tip or an RNase-free pellet pestle. Agitate in a shaker or tape to a vortexer left on overnight. Spin down to pellet the acrylamide and remove as much supernatant as possible, avoiding the acrylamide. Butanol extract supernatant down to about 200 ml. Ethanol precipitate and resuspend the pellet in 5 ml 10 mM Tris, pH 8.0. Quantify RNA concentration using a Nanodrop spectrophotometer.

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6. Application: Single-Molecule FRET Measurements We have developed the splinted RNA ligation procedure outlined in this chapter to generate site-specifically dye-labeled telomerase RNA constructs. These modified telomerase RNA constructs may be used to characterize dynamic RNA structural properties using Forster resonance energy transfer (FRET) (Stone et al., 2007). Our laboratory specializes in single molecule FRET measurements, which facilitates the direct observation of transient RNA structural states. The details of single molecule FRET A

Prism Quartz slide Sample FRET-labeled Telomerase RNA

Objective Fluorescence Acceptor (Cy5)

FRET =

IA IA + ID CCD

Donor (Cy3)

Donor Acceptor intensity (ID) intensity (IA)

C Intensity (A.U.)

B

250 200

1500 1000 500 0

150 100 FRET

Number of molecules

300

50 0 0

0.2

0.4 0.6 FRET

0.8

1.0

0

20

40

60 Time (s)

80

100

120

0

20

40

60 Time (s)

80

100

120

1.0 0.8 0.6 0.4 0.2 0

Figure 2.6 Application of splinted RNA ligation procedure: single molecule FRET. (A) Diagram of prism-type total internal reflection fluorescence microscope (TIRFM) for single molecule FRET measurements. (B) Distribution of single-molecule FRET values for dye-labeled telomerase RNA molecules generated by splinted RNA ligation. (C) Dye intensity and FRET traces of a single telomerase RNA molecule: Cy3 emission (green), Cy5 emission (red), FRET ratio (blue).


45

techniques have been described elsewhere (Roy et al., 2008), so we will only briefly discuss the technique to demonstrate one of many possible applications of splinted RNA ligation techniques. The protocols described in the previous sections enable us to produce extremely pure samples of Cy3–Cy5-labeled telomerase RNA. In addition, we often engineer the 50 -end of the RNA to possess a 15-base sequence extension that is annealed to a complementary DNA strand containing a 30 -biotin modification (Fig. 2.6A). The biotin moiety serves to immobilize the FRET-labeled telomerase RNA on a streptavidin-coated microscope slide. In this way, individual telomerase RNA molecules can be imaged onto a CCD camera using total internal reflection microscopy. A typical single molecule FRET measurement is conducted by illuminating the sample with a 532-nm laser, which directly excites the Cy3 (donor) dye. If the Cy3 dye is sufficiently close (2–9 nm) to the Cy5 (acceptor) dye, energy transfer can occur resulting in a mixture of emitted light coming from the donor and acceptor dyes. Because the emission spectra of Cy3 and Cy5 are sufficiently separated, fluorescence coming from each dye may be separated using a dichroic mirror, and the FRET ratio, defined as the intensity of the acceptor dye (IA) divided by the sum of the acceptor plus donor dyes (IA þ ID), may be determined. In this experimental geometry, FRET measurements can be conducted for many hundreds of telomerase RNAs simultaneously, allowing one to rapidly construct a histogram of observed FRET values (Fig. 2.6B). However, the true power of single molecule approaches is the ability to analyze FRET trajectories for individual RNA molecules (Fig. 2.6C), which has been used characterize structural dynamics of ribozymes (Zhuang et al., 2000) and ribonucleoprotein complexes (Cornish et al., 2008; Stone et al., 2007).

ACKNOWLEDGMENTS We thank our colleagues Xiaowei Zhuang, Mariana Mihalusova, John Y. Wu, and Kathleen Collins for their contributions to the work described in this chapter. We apologize to those scientists whose work was not cited due to space considerations. The research described in this chapter was supported by the National Institutes of Health, Howard Hughes Medical Institute, and University of California at Santa Cruz startup funds.

REFERENCES Breaker, R. R., and Joyce, G. F. (1994). A DNA enzyme that cleaves RNA. Chem. Biol. 1, 223–229. Cate, J. H., et al. (1999). X-ray crystal structures of 70 S ribosome functional complexes. Science 285, 2095–2104. Cech, T. R. (1990). Self-splicing of group I introns. Annu. Rev. Biochem. 59, 543–568. Cohen, S. B., and Cech, T. R. (2001). Engineering disulfide cross-links in RNA using thioldisulfide interchange chemistry. Curr. Protoc. Nucleic Acid Chem. Chapter 5, Unit 5 1.

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Cornish, P. V., et al. (2008). Spontaneous intersubunit rotation in single ribosomes. Mol. Cell. 30, 578–588. England, T. E., and Uhlenbeck, O. C. (1978). Enzymatic oligoribonucleotide synthesis with T4 RNA ligase. Biochemistry 17, 2069–2076. Ferre-D’Amare, A. R., and Doudna, J. A. (1996). Use of cis- and trans-ribozymes to remove 50 and 30 heterogeneities from milligrams of in vitro transcribed RNA. Nucleic Acids Res. 24, 977–978. Han, H., and Dervan, P. B. (1994). Visualization of RNA tertiary structure by RNA-EDTA. Fe(II) autocleavage: Analysis of tRNA(Phe) with uridine-EDTA.Fe(II) at position 47. Proc. Natl. Acad. Sci. USA 91, 4955–4959. Hengesbach, M., et al. (2008). RNA intramolecular dynamics by single-molecule FRET. Curr. Protoc. Nucleic Acid Chem. Chapter 11, Unit 11 12. Inoue, H., et al. (1987). Sequence-dependent hydrolysis of RNA using modified oligonucleotide splints and RNase H. FEBS Lett. 215, 327–330. Lapham, J., and Crothers, D. M. (1996). RNase H cleavage for processing of in vitro transcribed RNA for NMR studies and RNA ligation. RNA 2, 289–296. Lapham, J., et al. (1997). The position of site-directed cleavage of RNA using RNase H and 20 -O-methyl oligonucleotides is dependent on the enzyme source. RNA 3, 950–951. Moore, M. J., and Query, C. C. (2000). Joining of RNAs by splinted ligation. Methods Enzymol. 317, 109–123. Moore, M. J., and Sharp, P. A. (1992). Site-specific modification of pre-mRNA: The 20 -hydroxyl groups at the splice sites. Science 256, 992–997. Moore, M. J., and Sharp, P. A. (1993). Evidence for two active sites in the spliceosome provided by stereochemistry of pre-mRNA splicing. Nature 365, 364–368. Nandakumar, J., et al. (2004). RNA substrate specificity and structure-guided mutational analysis of bacteriophage T4 RNA ligase 2. J. Biol. Chem. 279, 31337–31347. Pinol-Roma, S., et al. (1989). Ultraviolet-induced cross-linking of RNA to proteins in vivo. Methods Enzymol. 180, 410–418. Price, S. R., et al. (1995). Crystallization of RNA-protein complexes. I. Methods for the large-scale preparation of RNA suitable for crystallographic studies. J. Mol. Biol. 249, 398–408. Romaniuk, P. J., and Uhlenbeck, O. C. (1983). Joining of RNA molecules with RNA ligase. Methods Enzymol. 100, 52–59. Roy, R., et al. (2008). A practical guide to single-molecule FRET. Nat. Methods 5, 507–516. Scaringe, S. A., et al. (1998). Novel RNA synthesis method using 50 -O-silyl-20 -O-orthoester protecting groups. J. Am. Chem. Soc. 120, 11820–11821. Silverman, S. K. (2008). Catalytic DNA (deoxyribozymes) for synthetic applications-current abilities and future prospects. Chem. Commun. (Camb.) 14(30), 3467–3485. Stone, M. D., et al. (2007). Stepwise protein-mediated RNA folding directs assembly of telomerase ribonucleoprotein. Nature 446, 458–461. Walker, G. C., et al. (1975). T4-induced RNA ligase joins single-stranded oligoribonucleotides. Proc. Natl. Acad. Sci. USA 72, 122–126. Walter, N. G. (2003). Probing RNA structural dynamics and function by fluorescence resonance energy transfer (FRET). Curr. Protoc. Nucleic Acid Chem. Chapter 11, Unit 11 10. Wyatt, J. R., et al. (1991). Synthesis and purification of large amounts of RNA oligonucleotides. Biotechniques 11, 764–769. Yu, Y. T. (2000). Site-specific 4-thiouridine incorporation into RNA molecules. Methods Enzymol. 318, 71–88. Zawadzki, V., and Gross, H. J. (1991). Rapid and simple purification of T7 RNA polymerase. Nucleic Acids Res. 19, 1948. Zhuang, X., et al. (2000). A single-molecule study of RNA catalysis and folding. Science 288, 2048–2051.

C H A P T E R

T H R E E

Methods of Site-Specific Labeling of RNA with Fluorescent Dyes Sergey Solomatin* and Daniel Herschlag† Contents 1. Introduction 2. Design of Labeled RNA Constructs 2.1. Selection of labeling sites and construct assembly methods 2.2. Design of the construct assembly 2.3. Selection of dyes for single molecule fluorescence studies 3. Dye Labeling of RNA Fragments 4. Notes on In Vitro Transcription with T7 RNA Polymerase 5. Assembly of Labeled RNA Constructs 6. Examples of Protocols 6.1. Protocol 1: Labeling of RNA oligos by with fluorescent dyes (NHS ester form) 6.2. Protocol 2: Ligation of large RNAs with T4 DNA ligase Acknowledgments References

48 49 49 49 51 53 55 56 58 58 61 66 66

Abstract Single molecule fluorescence techniques offer unique insights into mechanisms of conformational changes of RNA. Knowing how to make fluorescently labeled RNA molecules and understanding potential limitations of different labeling strategies is essential for successful implementation of single molecule fluorescence techniques. This chapter offers a step by step overview of the process of obtaining RNA constructs ready for single molecule measurements. Several alternative methods are described for each step, and ways of troubleshooting the most common problems, in particular, splinted RNA ligation, are suggested.

* Department of Biochemistry, Stanford University, Stanford, California, USA Departments of Biochemistry and Chemistry, Stanford University, Stanford, California, USA

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47

48

Sergey Solomatin and Daniel Herschlag

1. Introduction Over the past two decades, an ever increasing appreciation of multiple roles RNAs play in biology has led to an increasing interest in understanding the fundamental behavior of RNA. A broad variety of experimental approaches has been applied to studying structure and dynamics of RNA, including native gel electrophoresis, NMR, small angle X-ray scattering (SAXS), chemical structure probing, atomic force microscopy, UV, and fluorescence spectroscopy. Single molecule methods have become the latest frontier in studies of RNA dynamics (Weiss, 2000; Zhuang, 2005), revealing unique information about the behavior of RNA molecules hidden from bulk experiments by ensemble averaging (Downey et al., 2006; Ha et al., 1999; Hodak et al., 2005; Lee et al., 2007; Qu et al., 2008; Russell et al., 2002; Xie et al., 2004; Zhuang et al., 2000, 2002). Fluorescence-based techniques have two characteristics that make them particularly useful in single molecule implementation. First, they allow increased throughput of single molecule measurements through simultaneous observation of hundreds of individual molecules at the same time. Analysis of data from thousands of individual molecules is essential for bridging the gap between observations made on individual molecules and traditional bulk measurements made with ensembles of 1020 molecules. Second, an ability to label different parts of RNA molecules allows one to study the dynamics at a submolecular level and obtain increasingly detailed information about the RNA structure and motions. To take full advantage of these traits, one needs to incorporate fluorescent dyes site-specifically into an RNA of interest and to be able to do this at different selected position on an RNA molecule. This chapter outlines general strategies of preparing dye-labeled RNA constructs, in particular concentrating on double-labeled constructs for single molecule fluorescence resonance energy transfer (smFRET) measurements (Ha, 2001; Roy et al., 2008). These constructs are designed such that the dynamics of interest is revealed through changes of the distance between two dye labels, donor and acceptor, which in turn result in changes of the energy transfer efficiency from the donor to the acceptor. Anticorrelated changes of the donor and acceptor fluorescence, resulting from changes in energy transfer efficiency, are easy to distinguish from uncorrelated fluctuations of the fluorescence intensity arising from multiple possible sources. Because of this trait, FRET is the technique that has been most widely used for studying RNA dynamics (Weiss, 2000; Zhuang, 2005). A step by step overview of the process of obtaining RNA constructs ready for single molecule FRET is presented here. For each step, several alternative methods described in the literature are suggested. Means of

Methods of Site-Specific Labeling of RNA with Fluorescent Dyes

49

troubleshooting of the most common problems, in particular, splinted RNA ligation, are suggested based on the literature and on the authors’ personal experience.

2. Design of Labeled RNA Constructs 2.1. Selection of labeling sites and construct assembly methods Design of a labeled RNA construct is aimed at achieving two goals: 1. Obtaining the best possible FRET signal. 2. Minimally perturbing the behavior of the RNA. Observation of a FRET signal requires two dyes to be positioned on the RNA within the range of efficient energy transfer determined by the Fo¨rster distance for a particular dye pair (typically, 3–6 nm). A crystal structure is a great starting point for identifying appropriate labeling positions. A simple heuristic rule is to choose labeling sites that are remote in the secondary structure, but close in the tertiary structure. However, one should be careful not to perturb residues that might be involved in forming long-range tertiary interactions (e.g., Brion and Westhof, 1997, and references therein). Such residues are likely to conform to the rule above, but modifying them can severely destabilize the native structure of the RNA. On the other hand, labeling base paired regions adjacent to residues that make long-range tertiary contacts is safer, and it ensures that FRET will be observed. If a crystal structure is not available, phylogenetic ( Jaeger et al., 1994; Michel and Westhof, 1990), cross-linking (Chen et al., 1998), and biochemical (Lehnert et al., 1996) data can help guide the search for best labeling sites.

2.2. Design of the construct assembly Labeling with two different dyes typically requires the final RNA construct to be assembled from at least two fragments (Scheme 3.1). Depending on the choice of labeling sites, one of the following assembly strategies can be pursued: 1. RNA is made (synthetically or by in vitro transcription) in one piece, and labeled by base pairing to complementary-labeled oligos: (a) Two oligos are bound at the 50 and 30 ends (b) One of RNA ends is labeled directly, and an oligo is bound at the other end (c) One oligo is bound at an end and the other at an internal site

50


1

2

a)

b)

c)

d)

a)

b) Ligation

e)

Circular permutation

Scheme 3.1 Design strategies for making labeled RNA constructs. Numeration corresponds to the outline numbering in the text. Brown lines designate RNA, blue lines designate DNA oligos, stars designate fluorescent dyes.

(d) Two oligos are bound at internal sites. (e) New 50 and 30 ends are designed by circular permutations, and an oligo is bound at an internal site. 2. RNA is split into several fragments: (a) RNA is reassembled by base pairing of the fragments. (b) RNA is reassembled by covalent joining of the fragments. Assembly method 2.b (Lee et al., 2007; Sattin et al., 2008) reproduces exactly the same RNA as of the original unlabeled construct and can be considered the least perturbing method of labeling, as long as the positions of the dyes were appropriately chosen. However, it also remains the most technically challenging method, and it imposes the strictest requirements on the purity of RNA fragments, as discussed below. Other methods of assembly (1.a–e and 2.a) are easier to implement, but they restrict the choice of labeling sites to the ends of the molecule (1.a–c and 2.a), and/or require assumptions that modifications of the sequence— that is, binding of oligos at the ends and at internal sites (1.b–d), circular permutations (1.e) or nicks in the continuous backbone (2.a)—do not affect the dynamics of interest. Testing such assumptions can be nontrivial. The modular architecture of RNA structure lends some support to the assumption that adding duplexes at the ends of the molecule will not generally perturb its behavior (Brion and Westhof, 1997; Tinoco and Bustamante, 1999). Single molecule studies demonstrated that the overall folding rate, substrate docking rate and catalysis by the Tetrahymena group I


51

ribozyme were the same for the construct labeled by and oligo annealed to a 30 end extension as for unmodified construct (Russell et al., 2002; Zhuang et al., 2000, 2002). This method of labeling has been most widely used, but it has an obvious limitation of placing dyes in the vicinity of the ends of the molecule. In principle, the ends of an RNA molecule can be moved by making circular permutations (Pan, 2000). As it is known that circular permutations can strongly affect folding mechanisms of RNA (Lease et al., 2007; Pan et al., 1999), this approach may be mostly useful for studying structure and local dynamics of RNA. Also, after a circular permutation the ends of the molecule are expected to be right next to each other, so that the second dye has to be placed at some internal position. To place a dye at internal positions without breaking the RNA backbone, the Pan lab developed a method that involves replacing nonessential hairpin loops within RNA sequences with larger loops with specific sequences that are hybridized to labeled DNA oligos (Smith et al., 2005). These modifications had little effect on structure, as assayed by chemical footprinting, or catalytic efficiency of the catalytic domain of RNase P. The same method was successfully employed for studying ribosome dynamics (Dorywalska et al., 2005).

2.3. Selection of dyes for single molecule fluorescence studies Single molecule FRET experiments push the limits of sensitivity and time resolution of the detection systems, because the goal of these experiments is to get as much information as possible from the weakest possible light source. Good photophysical properties of dyes are essential for getting the most out of these experiments. The following properties are highly desirable for smFRET dyes: (1) high extinction coefficient and quantum yield (i.e., most of the excitation light is converted into useful signal); (2) high stability against photobleaching (i.e., each molecule can be observed for a long time); (3) stable fluorescent signal (i.e., no chemical or conformational transformations of the dye leading to large fluctuations of fluorescence such as blinking); (4) good spectral overlap of donor emission and acceptor excitation (allowing high maximum FRET efficiency); (5) good spectral separation of donor and acceptor emissions (i.e., easy to optically separate two signals and calculate the actual value of FRET); (6) donor and acceptor emission in the range of high quantum efficiency of the detection systems (e.g., for some CCDs quantum efficiency falls off sharply outside of the 450–850 nm window). Currently, one can choose from a broad variety of organic fluorophores covering the entire optical spectrum from UV to near IR that are commercially available (see Table 3.1). A lot of early smFRET work was performed

52


Table 3.1 Spectral properties of several selected fluorophores for RNA labeling Dye

Cyanine fluorphores Cy2 Cy3 Cy5 Cy5.5 Alexa fluorophores Alexa 350 Alexa 430 Alexa 488 Alexa 532 Alexa 555 Alexa 647 Alexa 700 ATTO fluorophores ATTO 425 ATTO 532 ATTO 647 ATTO 700 Other fluorophores Fluorescein (0.1 M NaOH) Tetramethyl rhodamine Texas Red

lex (nm)

lem (nm)

e (M 1 cm 1)

F

489 548 649 675

506 562 670 694

150,000 150,000 250,000 190,000

0.12 0.16–0.39 0.28 0.23

346 430 494 530 555 651 702

445 545 517 555 572 672 723

19,000 15,000 73,000 81,000 155,000 270,000 205,000

– – 0.92 0.61 0.1 0.33 0.25

436 532 645 700

484 553 669 719

45,000 115,000 120,000 120,000

0.90 0.90 0.20 0.25

495

519

79,000

0.79–0.95

557

576

103,000

0.2

589

615

139,000

0.9

with the cyanine dyes Cy3 and Cy5, and, while several lines of dyes were marketed recently as superior to cyanine dyes, the Cy dyes are still widely used because they have a highly desirable combination of properties. The fluorescence signal of Cy dyes is strong, long-lived and stable in oxygen-depleted environments in the presence of stabilizing agents,1 and they have well-separated emission spectra in the range ideal for most detection systems. Manufacturers of Alexa (Invitrogen), Dylight (Thermo Fisher Scientific), and ATTO (ATTO-Tec, also available from Sigma-Aldrich) dyes offer a broad choice of fluorophores covering the entire visible spectrum in small increments. Red dyes from these lines are reported to be significantly 1

Trolox (Rasnik et al., 2006) is an exceptionally good one, but other compounds, such as b-mercaptoethanol, n-propyl gallate, ascorbic acid, or chloramphenicol, have been used (Widengren et al., 2007).


53

more photostable than Cy5.2 Furthermore, ATTO dyes display exceptionally high brightness and have low intersystem crossing rates, which should favorably reflect on the stability of their fluorescence signal. Furthermore, some of the older, nonbranded, dyes, for example, Texas Red (Ha et al., 1996), fluorescein (Xie et al., 2004), or tetramethyl rhodamine (Lang et al., 2004), can work perfectly well in single molecule applications, and these dyes typically cost less than those noted above. All of the above dyes can be purchased in N-succinimide activated form and used for labeling of amino groups (see below). The choice of fluorophores are more limited for labeling of thiol groups with maleimide derivatives of dyes, and especially for full synthesis of oligos using dye-labeled phosphoramidites.

3. Dye Labeling of RNA Fragments Any strategy of assembly of a labeled RNA construct for smFRET requires obtaining two dye-labeled RNA fragments, typically two oligos. Dye-labeled RNA fragments can be obtained in one of the following ways: 1. Commercially labeled oligos. 2. In-house labeling of commercially synthesized oligos with amino- or thiol groups. 3. Full in-house oligonucleotide synthesis with 50 end incorporation of dye phosphoramidites. 4. Direct RNA labeling. One should keep in mind that not every labeling method is compatible with assembly strategies that require ligation of the oligos, for example enzymatic ligation with T4 DNA ligase would require 50 monophosphate group and a free hydroxyl group at the 30 of each junction to be ligated, whereas RNAs labeled by dye phosphoramidite incorporation or direct labeling at the 50 and 30 ends will have different end groups most likely incompatible with the ligation. 1. Commercially labeled oligos. Several commercial oligos manufacturers (e.g., International DNA Technology, or IDT, www.idtdna.com, Dharmacon, www.dharmacon.com, Gene Link, www.genelink.com) provide an option of purchasing a custom RNA oligos with a variety of dyes incorporated at the 50 end, 30 end or internally (currently only at uracils). While undeniably convenient, this strategy often is not the most 2

These comparisons (by the manufacturers) were probably made in oxygen-rich environments, as the differences appear to be small in oxygen-depleted solutions in the presence of Trolox (unpublished observations).

54


cost-effective because of higher prices and low yields of supplied oligos. Furthermore, the choice of dyes is limited to those few that available from the RNA manufacturer, and currently the selection is narrow for dyes to be incorporated at 30 and internal positions. 2. Labeling of amino- or thiol-modified oligos. In-house labeling of commercially synthesized oligos carrying reactive groups at specified positions combines the convenience of not having to synthesize the full oligo sequence with much broader flexibility of choosing one’s favorite dyes and placing them at any position along the oligo sequence. These factors, along with the lower cost of synthesis, makes in house labeling strategy our current favorite. Most fluorescent dyes are available as N-hydroxysuccinimide (NHS) or maleimide derivatives, ready for conjugation to primary amino- or thiol groups, respectively. As natural RNAs lack strongly nucleophilic aliphatic primary amino groups, the use of NHS-derivatives is typically easier and more straightforward than thiol modification by maleimides, which may require working in reducing or oxygen-free environments. Amino groups can be incorporated into custom RNA oligos during synthesis at either of the ends (through an aliphatic linker), internally at the uridine base as 5-aminoallyl uridine or at the backbone (Uni-LinkTM by IDT).3 The labeling reaction (Scheme 3.2) is easy to perform (see below), as long as certain precautions are taken. It is necessary to make sure that amines are not present in the reaction mix (e.g., TRIS-based buffers are incompatible with NHS labeling) and that the pH is optimal for labeling. At low pH most of the amino groups are protonated and their reactivity is low, but at high pH NHS esters can be hydrolyzed faster than they can react with the oligo, resulting in low yields. Carbonate or phosphate-based buffers at pH 8–9 typically work well. R2 NH2

O R1 O

N

R1

N R2 H Dye-labeled RNA

O

OH

O + O

O R1

OH Inactivated dye

O

OH

O H2O

N

+ O

N

O

Scheme 3.2 The labeling reaction of amino-modified RNA (R2) with NHS-activated dye (R1) and a competing side reaction of NHS hydrolysis.

3

20 -Amino ribose can also be used, but amino group in this position is much less reactive than a typical aliphatic amine.


55

Purification of the labeled oligo from excess dye can be accomplished by a combination of ethanol precipitation and PAGE or HPLC purification. Changes in the oligo mobility resulting from dye labeling are usually sufficient to purify the labeled oligo from the unlabeled material. Accomplishing this purification is beneficial for single molecule experiments, as it decreases the number of molecules that are not labeled, or labeled with a single dye only. 3. Full in-house oligonucleotide synthesis with 50 end incorporation of dye phosphoramidites. Due to wide availability of commercially synthesized oligonucleotides, full synthesis of RNA oligos in individual labs is not frequently carried out these days and choices of commercially available dye phosphoramidites are limited. Thermo Fisher Scientific (www. thermo.com) provides a selection of DyLight phosphoramidites that have spectral properties analogous to Cy3, Cy5, and Cy5.5 dyes. 4. Direct RNA labeling. Several methods of labeling the directly labeling transcribed RNA molecules been reported in the literature, but have not yet found wide applications. Dyes at the 50 were incorporated cotranscriptionally by initiating with a dye–guanosine conjugate (Fang et al., 1999), at the 30 end through oxidation of the terminal ribose to aldehyde form by sodium periodate with subsequent reaction with hydrazine derivatives of dyes (Proudnikov and Mirzabekov, 1996), and at the 20 hydroxyl in the middle of an RNA by deoxyribozyme-catalyzed ligation (Baum and Scott, 2007).

4. Notes on In Vitro Transcription with T7 RNA Polymerase RNA fragments longer than 40–50 nucleotides are most often synthesized by in vitro transcription with T7 RNA polymerase. Heterogeneity of transcripts obtained by this method must be recognized by researchers aiming to use such transcripts for ligations, as it can lead to serious artifacts that are especially notable in single molecule experiments. T7 RNA polymerase strongly prefers guanosine at the first and second transcribed positions, otherwise the transcription yields drastically decrease (Milligan and Uhlenbeck, 1989). However, if T7 polymerase encounters four or more guanosines in a row at the start site, it generates heterogeneity at the 50 end (Pleiss et al., 1998). A proper choice of the starting sequence appears to be sufficient to avoid this problem. However, heterogeneity at the 30 end is the rule rather than the exception for any RNA sequence transcribed by T7 polymerase. Run-off in vitro transcripts almost invariably contain a significant fraction of nontemplated nucleotides at the 30 end, with n þ 1, n þ 2, and n þ 3

56


transcripts being the main contaminants (Milligan and Uhlenbeck, 1989). For transcripts that are larger than 100 nt this contamination is not easy to recognize and essentially impossible to purify away. To obtain clean 30 ends, it is best to extend the transcribed sequence beyond the intended end and then cleave the RNA at the desired site. RNA can be extended with (a) a sequence that encodes one of small ribozymes (Ferre-D’Amare and Doudna, 1996; Price et al., 1995); (b) a recognition sequence for a DNAzyme (Santoro and Joyce, 1997); or (c) with a sequence complementary to a DNA oligo with subsequent cleavage of the hybrid by RNase H (Stone et al., 2007, see Akiyama and Stone, Chapter 2, this volume). Extending the 30 end sequence with a cis-cleaving hammerhead ribozyme is an easy and efficient way to obtain ‘‘clean’’ 30 ends, as this gives cotranscriptional cleavage. However, it does place certain constraints on the sequence at cleavage site (Birikh et al., 1997). If the desired 30 end sequence is not compatible with the hammerhead cleavage, another small ribozyme or other methods mentioned above can be used. Small ribozymes and DNAzymes leave 20 –30 cyclic phosphate at the site of cleavage, and this group must be removed before the ligation. This task can be accomplished by treating cleaved products with polynucleotide kinase (PNK) in the absence of ATP (Schurer et al., 2002).

5. Assembly of Labeled RNA Constructs 1. Noncovalent assembly by Watson–Creek base pairing. Assembly of larger RNA constructs through base pairing of complementary oligos, or oligos and in vitro transcripts, is the most widely used method in single molecule fluorescence field (Ha et al., 1999; Hodak et al., 2005; Xie et al., 2004; Zhuang et al., 2002). Its biggest advantage is the ease of the procedure, which usually involves simple mixing of the solutions of oligonucleotides and annealing via some combination of heating and cooling steps. Upon annealing, the efficiency of construct assembly can be tested using nondenaturing acrylamide gels. Purification of the fully assembled constructs can also be done using nondenaturing PAGE. 2. Covalent incorporation of labeled oligos. Full-length-labeled RNA molecules of essentially any size can be obtained by joining together labeled RNA oligos (typically obtained by synthesis) and either synthetic or in vitro transcribed RNA fragments comprising the rest of the sequence. Joining several RNA fragments into a single chain can be done using (i) protein ligases, (ii) deoxyribozyme ligase, and (iii) chemical ligations. (a) Enzymatic ligation using T4 DNA ligase (Moore and Sharp, 1992), remains the most often used process, offering significant advantages


57

over RNA ligase I as discussed by Moore and Query (2000). Recently discovered RNA ligase II (Ho and Shuman, 2002) does not suffer from many of the limitations of RNA ligase I and may become an efficient alternative to T4 DNA ligase. Heterogeneity of fragments that need to be ligated is one of the biggest obstacles for enzymatic ligations. Heterogeneity of in vitro transcripts was discussed above, and it must be avoided. Synthetic oligos are always contaminated by shorter products (n 1, n 2, etc.) because of

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