Methods in Enzymology Vol 461: Chemokines, Part B

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: 978-0-12-374907-9 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

Ronen Alon Department of Immunology, The Weizmann Institute of Science, Rehovot, Israel Isabel Alves Department of Chemistry, Universite Pierre et Marie Curie, Paris, France Shirley Appelbe Neuroscience and Molecular Pharmacology, University of Glasgow, Glasgow, Scotland, United Kingdom Tione Buranda Department of Pathology and Cancer Center, University of New Mexico Health Science Center, Albuquerque, New Mexico, USA Gabriele S. V. Campanella Center for Immunology and Inflammatory Diseases, Division of Rheumatology, Allergy and Immunology, Massachusetts General Hospital, Harvard Medical School, Charlestown, Massachusetts, USA Jonathan J. Cannon Department of Computer Science and Engineering, Washington University in St. Louis, St. Louis, Missouri, USA Percy H. Carter Research & Development, Bristol-Myers Squibb Company, Princeton, New Jersey, USA Jenna L. Cash Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom Hyeryun Choe Department of Pediatrics, Harvard Medical School, Perlmutter Laboratory, Children’s Hospital, Boston, Massachusetts, USA John Dempster University of Strathclyde, Institute for Pharmacy & Biomedical Sciences, Glasgow, Scotland, United Kingdom Michael Farzan Department of Microbiology and Molecular Genetics, Harvard Medical School, New England Primate Research Center, Southborough, Massachusetts, USA xi

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David R. Greaves Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom Damon J. Hamel Skaggs School of Pharmacy and Pharmaceutical Science, University of California, San Diego, La Jolla, California, USA Tracy M. Handel Skaggs School of Pharmacy and Pharmaceutical Science, University of California, San Diego, La Jolla, California, USA Richard Horuk Department of Pharmacology, UC Davis, Davis, California, USA Victor Hruby Department of Chemistry, and Department of Biochemistry and Molecular Biophysics, University of Arizona, Tucson, Arizona, USA Ariane Jansma Skaggs School of Pharmacy and Pharmaceutical Science, University of California, San Diego, La Jolla, California, USA Pia C. Jensen Department of Neuroscience and Pharmacology, Laboratory for Molecular Pharmacology, The Panum Institute, University of Copenhagen, Copenhagen, Denmark Francis Lin Center for Molecular Biology and Medicine, Veterans Affairs Palo Alto Health Care System, Palo Alto, California, USA, and Laboratory of Immunology and Vascular Biology, Department of Pathology, School of Medicine, Stanford University, Stanford, California, USA, and Department of Physics and Astronomy, University of Manitoba, Winnipeg, Manitoba, Canada Tina Y. Liu Department of Biochemistry, Medical College of Wisconsin, Milwaukee, Wisconsin, USA Tamara Loos Laboratory of Molecular Immunology, Rega Institute for Medical Research, Leuven, Belgium Andrew D. Luster Center for Immunology and Inflammatory Diseases, Division of Rheumatology, Allergy and Immunology, Massachusetts General Hospital, Harvard Medical School, Charlestown, Massachusetts, USA Mario Mellado Department of Immunology and Oncology, Centro Nacional de Biotecnologı´a/ CSIC, Madrid, Spain

Contributors

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Mark J. Miller Washington University School of Medicine, Department of Pathology and Immunology, St. Louis, Missouri, USA Graeme Milligan Neuroscience and Molecular Pharmacology, University of Glasgow, Glasgow, Scotland, United Kingdom Anneleen Mortier Laboratory of Molecular Immunology, Rega Institute for Medical Research, Leuven, Belgium Laura Martinez Mun˜oz Department of Immunology and Oncology, Centro Nacional de Biotecnologı´a/ CSIC, Madrid, Spain Christopher M. Overall Departments of Biochemistry and Molecular Biology, University of British Columbia, and Oral Biological and Medical Sciences, Centre for Blood Research, Life Sciences Institute, Vancouver, British Columbia, Canada Ian Parker Departments of Neurobiology and Behavior, and Physiology and Biophysics, University of California, Irvine, California, USA Francis C. Peterson Department of Biochemistry, Medical College of Wisconsin, Milwaukee, Wisconsin, USA Robert Pless Department of Computer Science and Engineering, Washington University in St. Louis, St. Louis, Missouri, USA Paul Proost Laboratory of Molecular Immunology, Rega Institute for Medical Research, Leuven, Belgium Amanda E. I. Proudfoot Merck Serono Geneva Research Centre, Geneva, Switzerland Jose´ Miguel Rodrı´guez-Frade Department of Immunology and Oncology, Centro Nacional de Biotecnologı´a/ CSIC, Madrid, Spain Mette M. Rosenkilde Department of Neuroscience and Pharmacology, Laboratory for Molecular Pharmacology, The Panum Institute, University of Copenhagen, Copenhagen, Denmark

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Antal Rot MRC Centre for Immune Regulation, Institute of Biomedical Research, University of Birmingham, UK Zdzislaw Salamon Department of Biochemistry and Molecular Biophysics, University of Arizona, Tucson, Arizona, USA Ziv Shulman Department of Immunology, The Weizmann Institute of Science, Rehovot, Israel India Sielaff Merck Serono Geneva Research Centre, Geneva, Switzerland Larry A. Sklar Department of Pathology and Cancer Center, University of New Mexico Health Science Center, Albuquerque, New Mexico, USA Amanda E. Starr Departments of Biochemistry and Molecular Biology, University of British Columbia, Centre for Blood Research, Life Sciences Institute, Vancouver, British Columbia, Canada Andrew J. Tebben Research & Development, Bristol-Myers Squibb Company, Princeton, New Jersey, USA Gordon Tollin Department of Chemistry, and Department of Biochemistry and Molecular Biophysics, University of Arizona, Tucson, Arizona, USA Brian F. Volkman Department of Biochemistry, Medical College of Wisconsin, Milwaukee, Wisconsin, USA Gemma E. White Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom David L. Wokosin Northwestern University, Department of Physiology, Chicago, Illinois, USA Yang Wu Department of Pathology and Cancer Center, University of New Mexico Health Science Center, Albuquerque, New Mexico, USA Bernd H. Zinselmeyer Washington University School of Medicine, Department of Pathology and Immunology, St. Louis, Missouri, USA

PREFACE

Secreted signaling molecules like chemokines facilitate complex intercellular communication by means of interactions with cell membrane–spanning receptors. There are approximately 50 identified mammalian chemokines, and all share a common monomeric fold. There are also approximately 20 chemokine receptors, all having the seven transmembrane domain topology of G-protein–coupled receptors. Despite this apparent homogeneity, the variety of signals sent and received by them defies simple explanation. Clearly, the devil is in the details, and the structure/function relationships that govern these seemingly similar interactions are sure to be subtle and nuanced, requiring thoughtful experimentation to tease them apart. With this in mind, we have focused Volume 461 of the Methods in Enzymology series on methods to probe the physical characteristics, dynamics, modifications, and interactions of chemokines and chemokine receptors. These proteins have presented researchers with many hurdles, from the difficulty in preparing functional receptors, to the complex posttranslational modifications, to the disparity in in vitro and in vivo functionality of some chemokine variants. Hence, the topics presented herein run the gamut of biochemical disciplines, from in vitro nuclear magnetic resonance and plasmon resonance methods, to receptor modeling in silico, to model systems for measuring and even simulating in situ cell migration. In 1997 Richard Horuk edited volumes 287 and 288 in the Methods in Enzymology series on chemokines and chemokine receptors, putting together the first comprehensive practical guide to studying these molecules. Volume 461 is part two of two new editions on chemokines and their receptors. The previous volume 460 focused on studying the roles of chemokines and chemokine receptors in disease states, atypical chemokine receptors, chemokine signaling, and chemokine-related proteins from pathogens. Compilations like this are assembled by the immense efforts of many individual researchers, and we enthusiastically offer our thanks and gratitude to all of the authors who contributed to making these volumes a reality. We would also like to thank the incredible staff at Elsevier, and especially Delsy Retchagar and Tara Hoey, for valiantly trying to keep us on track and on time. We could not have done it without you. DAMON J. HAMEL AND TRACY M. HANDEL xv

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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VOLUME 257. Small GTPases and Their Regulators (Part C: Proteins Involved in Transport) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 258. Redox-Active Amino Acids in Biology Edited by JUDITH P. KLINMAN VOLUME 259. Energetics of Biological Macromolecules Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 260. Mitochondrial Biogenesis and Genetics (Part A) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 261. Nuclear Magnetic Resonance and Nucleic Acids Edited by THOMAS L. JAMES VOLUME 262. DNA Replication Edited by JUDITH L. CAMPBELL VOLUME 263. Plasma Lipoproteins (Part C: Quantitation) Edited by WILLIAM A. BRADLEY, SANDRA H. GIANTURCO, AND JERE P. SEGREST VOLUME 264. Mitochondrial Biogenesis and Genetics (Part B) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 265. Cumulative Subject Index Volumes 228, 230–262 VOLUME 266. Computer Methods for Macromolecular Sequence Analysis Edited by RUSSELL F. DOOLITTLE VOLUME 267. Combinatorial Chemistry Edited by JOHN N. ABELSON VOLUME 268. Nitric Oxide (Part A: Sources and Detection of NO; NO Synthase) Edited by LESTER PACKER VOLUME 269. Nitric Oxide (Part B: Physiological and Pathological Processes) Edited by LESTER PACKER VOLUME 270. High Resolution Separation and Analysis of Biological Macromolecules (Part A: Fundamentals) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 271. High Resolution Separation and Analysis of Biological Macromolecules (Part B: Applications) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 272. Cytochrome P450 (Part B) Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 273. RNA Polymerase and Associated Factors (Part A) Edited by SANKAR ADHYA VOLUME 274. RNA Polymerase and Associated Factors (Part B) Edited by SANKAR ADHYA

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VOLUME 275. Viral Polymerases and Related Proteins Edited by LAWRENCE C. KUO, DAVID B. OLSEN, AND STEVEN S. CARROLL VOLUME 276. Macromolecular Crystallography (Part A) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 277. Macromolecular Crystallography (Part B) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 278. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 279. Vitamins and Coenzymes (Part I) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 280. Vitamins and Coenzymes (Part J) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 281. Vitamins and Coenzymes (Part K) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 282. Vitamins and Coenzymes (Part L) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 283. Cell Cycle Control Edited by WILLIAM G. DUNPHY VOLUME 284. Lipases (Part A: Biotechnology) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 285. Cumulative Subject Index Volumes 263, 264, 266–284, 286–289 VOLUME 286. Lipases (Part B: Enzyme Characterization and Utilization) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 287. Chemokines Edited by RICHARD HORUK VOLUME 288. Chemokine Receptors Edited by RICHARD HORUK VOLUME 289. Solid Phase Peptide Synthesis Edited by GREGG B. FIELDS VOLUME 290. Molecular Chaperones Edited by GEORGE H. LORIMER AND THOMAS BALDWIN VOLUME 291. Caged Compounds Edited by GERARD MARRIOTT VOLUME 292. ABC Transporters: Biochemical, Cellular, and Molecular Aspects Edited by SURESH V. AMBUDKAR AND MICHAEL M. GOTTESMAN VOLUME 293. Ion Channels (Part B) Edited by P. MICHAEL CONN

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VOLUME 294. Ion Channels (Part C) Edited by P. MICHAEL CONN VOLUME 295. Energetics of Biological Macromolecules (Part B) Edited by GARY K. ACKERS AND MICHAEL L. JOHNSON VOLUME 296. Neurotransmitter Transporters Edited by SUSAN G. AMARA VOLUME 297. Photosynthesis: Molecular Biology of Energy Capture Edited by LEE MCINTOSH VOLUME 298. Molecular Motors and the Cytoskeleton (Part B) Edited by RICHARD B. VALLEE VOLUME 299. Oxidants and Antioxidants (Part A) Edited by LESTER PACKER VOLUME 300. Oxidants and Antioxidants (Part B) Edited by LESTER PACKER VOLUME 301. Nitric Oxide: Biological and Antioxidant Activities (Part C) Edited by LESTER PACKER VOLUME 302. Green Fluorescent Protein Edited by P. MICHAEL CONN VOLUME 303. cDNA Preparation and Display Edited by SHERMAN M. WEISSMAN VOLUME 304. Chromatin Edited by PAUL M. WASSARMAN AND ALAN P. WOLFFE VOLUME 305. Bioluminescence and Chemiluminescence (Part C) Edited by THOMAS O. BALDWIN AND MIRIAM M. ZIEGLER VOLUME 306. Expression of Recombinant Genes in Eukaryotic Systems Edited by JOSEPH C. GLORIOSO AND MARTIN C. SCHMIDT VOLUME 307. Confocal Microscopy Edited by P. MICHAEL CONN VOLUME 308. Enzyme Kinetics and Mechanism (Part E: Energetics of Enzyme Catalysis) Edited by DANIEL L. PURICH AND VERN L. SCHRAMM VOLUME 309. Amyloid, Prions, and Other Protein Aggregates Edited by RONALD WETZEL VOLUME 310. Biofilms Edited by RON J. DOYLE VOLUME 311. Sphingolipid Metabolism and Cell Signaling (Part A) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN

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VOLUME 312. Sphingolipid Metabolism and Cell Signaling (Part B) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN VOLUME 313. Antisense Technology (Part A: General Methods, Methods of Delivery, and RNA Studies) Edited by M. IAN PHILLIPS VOLUME 314. Antisense Technology (Part B: Applications) Edited by M. IAN PHILLIPS VOLUME 315. Vertebrate Phototransduction and the Visual Cycle (Part A) Edited by KRZYSZTOF PALCZEWSKI VOLUME 316. Vertebrate Phototransduction and the Visual Cycle (Part B) Edited by KRZYSZTOF PALCZEWSKI VOLUME 317. RNA–Ligand Interactions (Part A: Structural Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 318. RNA–Ligand Interactions (Part B: Molecular Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 319. Singlet Oxygen, UV-A, and Ozone Edited by LESTER PACKER AND HELMUT SIES VOLUME 320. Cumulative Subject Index Volumes 290–319 VOLUME 321. Numerical Computer Methods (Part C) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 322. Apoptosis Edited by JOHN C. REED VOLUME 323. Energetics of Biological Macromolecules (Part C) Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 324. Branched-Chain Amino Acids (Part B) Edited by ROBERT A. HARRIS AND JOHN R. SOKATCH VOLUME 325. Regulators and Effectors of Small GTPases (Part D: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 326. Applications of Chimeric Genes and Hybrid Proteins (Part A: Gene Expression and Protein Purification) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 327. Applications of Chimeric Genes and Hybrid Proteins (Part B: Cell Biology and Physiology) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 328. Applications of Chimeric Genes and Hybrid Proteins (Part C: Protein–Protein Interactions and Genomics) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON

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VOLUME 329. Regulators and Effectors of Small GTPases (Part E: GTPases Involved in Vesicular Traffic) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 330. Hyperthermophilic Enzymes (Part A) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 331. Hyperthermophilic Enzymes (Part B) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 332. Regulators and Effectors of Small GTPases (Part F: Ras Family I) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 333. Regulators and Effectors of Small GTPases (Part G: Ras Family II) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 334. Hyperthermophilic Enzymes (Part C) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 335. Flavonoids and Other Polyphenols Edited by LESTER PACKER VOLUME 336. Microbial Growth in Biofilms (Part A: Developmental and Molecular Biological Aspects) Edited by RON J. DOYLE VOLUME 337. Microbial Growth in Biofilms (Part B: Special Environments and Physicochemical Aspects) Edited by RON J. DOYLE VOLUME 338. Nuclear Magnetic Resonance of Biological Macromolecules (Part A) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 339. Nuclear Magnetic Resonance of Biological Macromolecules (Part B) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 340. Drug–Nucleic Acid Interactions Edited by JONATHAN B. CHAIRES AND MICHAEL J. WARING VOLUME 341. Ribonucleases (Part A) Edited by ALLEN W. NICHOLSON VOLUME 342. Ribonucleases (Part B) Edited by ALLEN W. NICHOLSON VOLUME 343. G Protein Pathways (Part A: Receptors) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 344. G Protein Pathways (Part B: G Proteins and Their Regulators) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 345. G Protein Pathways (Part C: Effector Mechanisms) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT

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VOLUME 346. Gene Therapy Methods Edited by M. IAN PHILLIPS VOLUME 347. Protein Sensors and Reactive Oxygen Species (Part A: Selenoproteins and Thioredoxin) Edited by HELMUT SIES AND LESTER PACKER VOLUME 348. Protein Sensors and Reactive Oxygen Species (Part B: Thiol Enzymes and Proteins) Edited by HELMUT SIES AND LESTER PACKER VOLUME 349. Superoxide Dismutase Edited by LESTER PACKER VOLUME 350. Guide to Yeast Genetics and Molecular and Cell Biology (Part B) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 351. Guide to Yeast Genetics and Molecular and Cell Biology (Part C) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 352. Redox Cell Biology and Genetics (Part A) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 353. Redox Cell Biology and Genetics (Part B) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 354. Enzyme Kinetics and Mechanisms (Part F: Detection and Characterization of Enzyme Reaction Intermediates) Edited by DANIEL L. PURICH VOLUME 355. Cumulative Subject Index Volumes 321–354 VOLUME 356. Laser Capture Microscopy and Microdissection Edited by P. MICHAEL CONN VOLUME 357. Cytochrome P450, Part C Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 358. Bacterial Pathogenesis (Part C: Identification, Regulation, and Function of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 359. Nitric Oxide (Part D) Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 360. Biophotonics (Part A) Edited by GERARD MARRIOTT AND IAN PARKER VOLUME 361. Biophotonics (Part B) Edited by GERARD MARRIOTT AND IAN PARKER VOLUME 362. Recognition of Carbohydrates in Biological Systems (Part A) Edited by YUAN C. LEE AND REIKO T. LEE

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

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VOLUME 382. Quinones and Quinone Enzymes (Part B) Edited by HELMUT SIES AND LESTER PACKER VOLUME 383. Numerical Computer Methods (Part D) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 384. Numerical Computer Methods (Part E) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 385. Imaging in Biological Research (Part A) Edited by P. MICHAEL CONN VOLUME 386. Imaging in Biological Research (Part B) Edited by P. MICHAEL CONN VOLUME 387. Liposomes (Part D) Edited by NEJAT DU¨ZGU¨NES, VOLUME 388. Protein Engineering Edited by DAN E. ROBERTSON AND JOSEPH P. NOEL VOLUME 389. Regulators of G-Protein Signaling (Part A) Edited by DAVID P. SIDEROVSKI VOLUME 390. Regulators of G-Protein Signaling (Part B) Edited by DAVID P. SIDEROVSKI VOLUME 391. Liposomes (Part E) Edited by NEJAT DU¨ZGU¨NES, VOLUME 392. RNA Interference Edited by ENGELKE ROSSI VOLUME 393. Circadian Rhythms Edited by MICHAEL W. YOUNG VOLUME 394. Nuclear Magnetic Resonance of Biological Macromolecules (Part C) Edited by THOMAS L. JAMES VOLUME 395. Producing the Biochemical Data (Part B) Edited by ELIZABETH A. ZIMMER AND ERIC H. ROALSON VOLUME 396. Nitric Oxide (Part E) Edited by LESTER PACKER AND ENRIQUE CADENAS VOLUME 397. Environmental Microbiology Edited by JARED R. LEADBETTER VOLUME 398. Ubiquitin and Protein Degradation (Part A) Edited by RAYMOND J. DESHAIES VOLUME 399. Ubiquitin and Protein Degradation (Part B) Edited by RAYMOND J. DESHAIES VOLUME 400. Phase II Conjugation Enzymes and Transport Systems Edited by HELMUT SIES AND LESTER PACKER

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VOLUME 401. Glutathione Transferases and Gamma Glutamyl Transpeptidases Edited by HELMUT SIES AND LESTER PACKER VOLUME 402. Biological Mass Spectrometry Edited by A. L. BURLINGAME VOLUME 403. GTPases Regulating Membrane Targeting and Fusion Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 404. GTPases Regulating Membrane Dynamics Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 405. Mass Spectrometry: Modified Proteins and Glycoconjugates Edited by A. L. BURLINGAME VOLUME 406. Regulators and Effectors of Small GTPases: Rho Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 407. Regulators and Effectors of Small GTPases: Ras Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 408. DNA Repair (Part A) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 409. DNA Repair (Part B) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 410. DNA Microarrays (Part A: Array Platforms and Web-Bench Protocols) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 411. DNA Microarrays (Part B: Databases and Statistics) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 412. Amyloid, Prions, and Other Protein Aggregates (Part B) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 413. Amyloid, Prions, and Other Protein Aggregates (Part C) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 414. Measuring Biological Responses with Automated Microscopy Edited by JAMES INGLESE VOLUME 415. Glycobiology Edited by MINORU FUKUDA VOLUME 416. Glycomics Edited by MINORU FUKUDA VOLUME 417. Functional Glycomics Edited by MINORU FUKUDA VOLUME 418. Embryonic Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA

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Isolation, Identification, and Production of Posttranslationally Modified Chemokines Tamara Loos,*,1 Anneleen Mortier,*,1 and Paul Proost* Contents 1. Introduction 2. Isolation and Stimulation of Peripheral Blood Mononuclear Cells (PBMC) 3. Concentration of Isolated Proteins 4. Affinity Chromatography 5. Specific Sandwich Enzyme-Linked Immunosorbent Assay (ELISA) 6. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 7. Ion Exchange Chromatography 8. Reverse-Phase High-Pressure Liquid Chromatography (RP-HPLC) 9. Ion Trap Mass Spectrometry 10. Edman Degradation 11. Total Protein Quantification Methods 12. Illustration: Isolation and Identification of Natural Posttranslationally Modified CXCL8 Isoforms 13. Solid-Phase Peptide Synthesis 13.1. Synthesis of the peptide chain 13.2. Deprotection of the synthesized peptide chain 13.3. Folding of the raw protein 14. Citrullination of Chemokines 15. Identification of Enzymes Generating the Natural Posttranslationally Modified Chemokines, as Exemplified by Aminopeptidase N(APN)/CD13 16. Comparison of the Heparin-Binding Properties of Chemokine Isoforms Acknowledgments References

* 1

4 5 6 7 7 8 9 10 10 11 12 13 14 15 19 19 21

23 24 25 25

Laboratory of Molecular Immunology, Rega Institute for Medical Research, Leuven, Belgium Both authors contributed equally

Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05401-9

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2009 Elsevier Inc. All rights reserved.

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Abstract Chemokines attract cells during the development of lymphoid tissues, leukocyte homing, and pathologic processes such as cancer and inflammation. Limited posttranslational modification of chemokines may significantly alter the glycosaminoglycan and/or receptor binding properties and signaling potency of these chemotactic proteins. To compare the in vitro and in vivo biologic activities of posttranslationally modified chemokine isoforms, considerable amounts of pure chemokine isoforms are required. This chapter describes a number of chromatographic techniques that are useful for the isolation of natural, posttranslationally modified chemokines from primary human cell cultures. In addition, combination of immunologic assays and biochemical techniques such as automated Edman degradation and mass spectrometry are used for the identification of modifications. Alternate methods for the generation of specific chemokine isoforms are discussed such as modification of chemokines by specific enzymes and total chemical syntheses and folding of chemokine isoforms. In particular, in vitro processing of chemokines by the protease aminopeptidase N/CD13 and citrullination or deamination of chemokines by peptidyl arginine deiminases (PAD) are described as methods for the confirmation or generation of posttranslationally modified chemokine isoforms.

1. Introduction Chemokines are crucial proteins for the directed migration of leukocytes during the development of lymphoid tissue, leukocyte homing, and inflammation (Springer, 1994). The regulation of chemokine activity is a crucial event for the outcome of the immune response. Posttranslational modifications were reported to regulate chemokine activity in addition to regulation of chemokine and chemokine receptor expression levels, the production of ‘‘decoy’’ or ‘‘scavenging’’ chemokine receptors, presentation of chemokines on glycosaminoglycans, and synergistic activity between chemokines and other chemotactic factors (Colditz et al., 2007; Gouwy et al., 2005; Johnson et al., 2005; Mantovani et al., 2006; Mortier et al., 2008). Since the discovery of the first inflammatory chemokines, proteolytic processing or glycosylation was detected on inflammatory chemokines such as CXCL8/interleukin-8 (IL-8), CXCL7/NAP-2 (neutrophil-activating peptide-2), and CCL2/MCP-1 (monocyte chemotactic protein-1) (Brandt et al., 1991; Furutani et al., 1989; Jiang et al., 1990; Robinson et al., 1989; Van Damme et al., 1989b; Walz et al., 1990). Recently, deaminated or citrullinated natural chemokines were also identified (Loos et al., 2008; Proost et al., 2008). Although for some chemokines, such as CXCL7, the importance of posttranslational processing for the regulation of

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chemokine activity was evidenced even before the identification of chemokine receptors, the presence and biologic consequences of other modifications were only recently resolved. For others (e.g., glycosylation of CCL2) the biologic consequences are still to be determined. The limited availability of the posttranslationally modified chemokines was and often remains an obstacle for detailed biochemical and biologic characterization. Gene regulation studies revealed that the chemokine production pattern depends much on the cell type and inducers used (Loos et al., 2006; Proost et al., 2003, 2004b). Moreover, the abundance of different posttranslationally modified isoforms of a particular chemokine varies within cell sorts (Gimbrone et al., 1989; Schro¨der et al., 1990; Van Damme et al., 1989a; Yoshimura et al., 1989). In an attempt to isolate novel CXCL8 and CXCL10/IP-10 (interferon-gamma–inducible protein-10) isoforms, induction experiments were performed on peripheral blood–derived mononuclear cells (PBMC) followed by a four-step purification procedure (Loos et al., 2008; Proost et al., 2008). First, the conditioned medium was concentrated to controlled pore glass or silicic acid, followed by the purification to homogeneity by heparin affinity, ion exchange, and reverse-phase high-pressure liquid chromatography (RP-HPLC). Immunoreactivity, quantity, and purity of the chemokine-containing fractions between each step were analyzed with specific enzyme-linked immunosorbent assays (ELISA) and sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Finally, ion trap mass spectrometry and Edman degradation were applied to identify posttranslational modifications. Protein quantification was performed with the Bradford or bicinchoninic acid (BCA) total protein assays. Alternatively, posttranslationally modified chemokines were chemically synthesized or recombinant chemokines were enzymatically processed in vitro to verify or generate modified isoforms.

2. Isolation and Stimulation of Peripheral Blood Mononuclear Cells (PBMC) Freshly isolated buffy coats (Blood Transfusion Center Red Cross, Leuven, Belgium) are treated with one volume of hydroxyethyl-starch (Plasmasteril, Fresenius, Bad Homburg, Germany) and one volume of Dulbecco’s phosphate-buffered saline (DPBS; 0.0095 M phosphate buffer, pH 7.35, without Ca2þ and Mg2þ) for 30 min at 37  C, resulting in the sedimentation of most erythrocytes (Van Damme et al., 1997, 2000). After centrifugation of the supernatant at 200g for 10 min, the pellet is washed with 50 ml PBS. Next, the resuspended pellet is loaded on top of three volumes of Ficoll-sodium metrizoate (Lymphoprep, Nycomed, Oslo, Norway), and mononuclear cells and granulocytes are segregated by

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gradient centrifugation at 400g for 30 min (without brakes). The PBMC appearing in the transition layer are removed and washed twice with PBS. Finally, the pellet is resuspended in RPMI 1640 enriched with 2% fetal calf serum (FCS) at an appropriate concentration (typically 2.106 to 5.106 cells/ ml) and stimulated with cytokines and/or Toll-like receptor (TLR) ligands. Serum is a very rich and complex source of growth factors and other proteins. This increases the chemokine production yield, although an addition of more then 2% FCS severely complicates purification of chemokines to homogeneity. After 24 to 96 h of culture at 37  C in the presence of 5% CO2, the conditioned medium is harvested, centrifuged at 200g, subsequently at 1000g, and finally stored at 20  C until purification to homogeneity. Because of the minute amounts of chemokines produced (typically ng/ml range), liters of conditioned medium are required if an amount of chemokine needs to be purified sufficient for subsequent biologic characterization.

3. Concentration of Isolated Proteins A tenfold concentration of the proteins present is obtained by adsorption to controlled pore glass (CPG) (Proost et al., 2004a; Struyf et al., 2000; Wuyts et al., 1997). During the production process of the CPG, the borosilicate base is heated until the borate is released from the silicate and pores are formed (particle size: 120 to 200 mesh; pore size: 35 nm; Serva, Heidelberg, Germany). These pores increase the surface of the matrix, the yield, and the purity of the proteins. The rather expensive CPG beads may be regenerated chemically by cleaning with concentrated nitric acid that degrades the proteins that are still present on the CPG beads after the elution procedure. Therefore, the use of silicic acid is often preferred (Matrex silica, particle size: 35 to 70 mm; pore size: 10 mm; Amicon, Beverly, MA). The binding to CPG (30 ml/L conditioned medium) and silicic acid (10 g/L) should be performed at neutral pH and 4  C for 2 h to acquire proper ionic interactions between the negatively charged silica and the positively charged proteins. Next, the CPG or the silicic acid is washed for 30 min with 10 mM glycine, pH 3.5, or DPBS containing 1 M NaCl, pH 7.4, respectively. The proteins are eluted from the CPG beads by adding 300 mM glycine, pH 2.0, to destroy the ionic interactions and hydrogen bonds. The elution from the silicic acid is performed by adding DPBS, pH 7.4, containing 1.4 M NaCl, breaking ionic interactions, and 50% ethylene glycol, splitting hydrogen bonds. An additional step of 300 mM glycine, pH 2.0, can be introduced to remove remaining proteins and thereby improve the yield. If acid-stable proteins need to be purified, this step may be implemented immediately. Finally, the eluate is neutralized and desalted through dialysis with a 3.5-kDa cutoff membrane against 50 mM TRIS and 50 mM NaCl,

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pH 7.4. During the last dialysis step, 15% polyethylene glycol (PEG) (average molecular mass, 20,000) may be added to concentrate the preparation to a smaller volume before subjection to affinity chromatography.

4. Affinity Chromatography Because chemokines are characterized by a high affinity for heparin, bulk impurities are removed with heparin affinity chromatography ( Janson and Ryden, 1989; Wuyts et al., 1997). The dialyzed eluate recovered from CPG or silicic acid adsorption is loaded on a heparin-Sepharose CL 6B column (GE Healthcare, Diegem, Belgium) in equilibration buffer (50 mM TRIS, pH 7.4, containing 50 mM NaCl). Elution from the column is performed at a flow rate of 20 ml/h (60 ml bed volume) by administrating an increasing NaCl gradient starting from 50 mM to 2 M in equilibration buffer, hereby first eluting the low-affinity proteins because of competition with Naþ ions for interaction with heparin. The column is regenerated by rinsing with 0.1 M TRIS, pH 8.5, containing 50 mM NaCl and subsequently with 0.1 M NaAc, pH 5.0, containing 0.5 M NaCl. Alternately, if a certain chemokine is aspired, specific antibody affinity chromatography can be applied (Wuyts et al., 1997). Purified antichemokine antibody is coupled to CNBr-activated Sepharose 4B (GE Healthcare). The amount of Sepharose required (1 g/3.5 ml final gel volume) is weight out and washed with 1 mM HCl on a sintered glass filter resulting in it swelling and forming a gel suspension. The antibody (5 to 10 mg/ml final gel) is dissolved in coupling buffer consisting of 0.1 M NaHCO3, pH 8.3, containing 0.5 M NaCl before mixing end-over-end with the Sepharose gel for 2 h at room temperature or overnight at 4  C. The remaining active groups on the Sepharose beads are blocked by adding 0.2 M glycine, pH 8.0, for 2 h at room temperature or overnight at 4  C. The excess antibody is washed away with coupling buffer alternating with a 0.1 M acetate buffer, pH 4.0, containing 0.5 M NaCl. Finally, the empty column is packed with the antibody-bound Sepharose. In analogy with heparin affinity chromatography, the chemokine preparation is equilibrated in DPBS, pH 7.5, before loading onto the column, and stepwise elution is obtained by administrating a 0.1 M NaCl buffer containing 0.1 M citrate, pH 2.0, at a flow rate of 40 ml/h.

5. Specific Sandwich Enzyme-Linked Immunosorbent Assay (ELISA) Fractions containing immunoreactivity are detected by specific sandwich enzyme-linked immunosorbent assays (ELISA). A 96-well ELISA plate is coated overnight at 4  C with a specific primary antibody reconstituted

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in DPBS. Unbound proteins are washed away with DPBS containing 0.05% Tween-20 (v/v) that decreases the surface tension. Next, empty regions on the plastic are blocked with 0.1% casein (w/v) in DPBS containing 0.05% Tween-20 for 1 h at 37  C. Samples are diluted in the same blocking buffer, and appropriate antigen binds to the primary antibody in the course of 2 h at 37  C. Subsequently, a secondary specific antibody originating from a different organism is added for 1 h at 37  C. Then, the plate is washed and the secondary antibody is targeted by a horseradish peroxidase (HRP)– labeled tertiary antibody for 30 min at 37  C. Alternately, the secondary antibody can be tagged with biotin recognized by HRP-labeled streptavidin. The chromogen 3,30 ,5,50 -tetramethylbenzidine (TMB; 0.42 mM; SigmaAldrich, St. Louis, MO) reconstituted in 0.1 M NaAc and 0.1 M citrate, pH 4.9, supplemented with 0.004% H2O2 (v/v) is added to the wells. The H2O2 is reduced into water by peroxidase, causing the colorless TMB chromogen to oxidize into a blue product. This reaction is stopped by adding an equal volume of 1 M H2SO4, resulting in development of a yellow color. The intensity of the yellow color is proportional to the amount of HRP and hence to the amount of bound chemokine. The optical density is measured at 450 nm with a spectrofluorometer (Titertek, Huntsville, Al). The concentration of bound antigen can be determined by implementing a dilution series of a chemokine standard.

6. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) The purity of the chemokine preparations is evaluated by SDS-PAGE ( Janson and Ryden, 1989; Kinter et al., 2000; Wuyts et al., 1997). To detect 5- to 20-kDa proteins, TRIS/Tricine gels consisting of three layers differing in acrylamide versus bisacrylamide composition (% T and % C) are used according to Eqs. (1.1) and (1.2) (Schagger et al., 1987).

% T ¼ ½acrylamide ðgÞ þ bisacrylamide ðgÞ  100=100ml

ð1:1Þ

% C ¼ bisacrylamide ðgÞ  100=½acrylamide ðgÞ þ bisacrylamide ðgÞ ð1:2Þ The upper layer, also called the ‘‘stacking’’ gel, contains the largest pores and concentrates the sample (5% T and 5% C). The ‘‘spacer’’ gel is the middle layer, separating the small proteins from the bulk (10% T and 3.3% C). The third and lowest layer is the ‘‘separating’’ layer, consisting of smaller pore sizes, resolving proteins in individual bands (13% T and 5% C). All gels are prepared in 3 M TRIS, pH 8.5, containing 0.3% SDS. N,N,N0 ,

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N0 -tetramethyl ethylenediamine (TEMED) (0.066%) and ammonium persulfate (0.66%) are added to initialize polymerization. The separating gel also contains 10% glycerol to avoid mixture with the spacer gel. Isobutanol is deposited on top of the spacer gel to avoid interference of oxygen that oxidizes radicals and, therefore, may prevent polymerization. Samples are loaded in 50 mM TRIS, pH 6.8, containing 4% SDS, 12% glycerol, 2% b-mercaptoethanol, 0.01% Brilliant Blue G (tracking dye; Bio-Rad Laboratories, Hercules, CA), and denaturation is performed by heating at 95  C for 5 min before loading onto the stacking gel. The anode buffer in the electrophoresis system contains 0.2 M TRIS, pH 8.9, whereas the upper cathode buffer consists of 0.1 M TRIS, pH 8.2, 0.1 M Tricine, and 0.1% SDS. Proteins are visualized by silver staining (Guevara Jr et al., 1982). To decrease background noise, the SDS-PAGE components are removed by resting the gel for 1 h in 20% ethanol, 5% acetic acid, and 2.5% sulfosalicylic acid, followed by three washes with 20% ethanol. The shrunk gel is then placed in silver staining solution. A solution of 1g AgNO3 in 10 ml degassed water is prepared and added dropwise to a mixture containing 2.1 ml 14.8 M NH4OH, 0.35 ml NaOH (32% w/v), and 40 ml ethanol in 150 ml degassed water. After 1 h, the gel is washed three times with 20% ethanol and developed in 20% ethanol, 0.01% citric acid, and 0.037% formaldehyde, which oxidizes Ag ions bound to protein clusters. Reduction of these oxidized Ag ions by citric acid results in the deposition of Ag. Development is stopped by treatment with 20% ethanol and 0.5% acetic acid. The gel is then washed for 30 min in water and stored overnight in 50% ethanol. Subsequently, gels and a small volume of 50% ethanol (to prevent the gels from drying) may be packed between sealed plastic sheets and stored for weeks in the dark at 4  C. Evaluation of the relative molecular mass (Mr) and the amount of proteins can be achieved by comparison with a standardized mixture of proteins (100 ng each) containing myosin (200 kDa), b-galactosidase (116.3 kDa), phosphorylase B (97.4 kDa), bovine serum albumin (66.3 kDa), glutamic dehydrogenase (55.4 kDa), lactate dehydrogenase (36.5 kDa), carbonic anhydrase (31 kDa), trypsin inhibitor (21.5 kD), lysozyme (14.4 kDa), aprotinin (6 kDa), and insulin B chain (3.5 kDa) (Bio-Rad Laboratories).

7. Ion Exchange Chromatography Most chemokines have a high pI. Therefore, chemokines may be separated with cation exchange chromatography ( Janson and Ryden, 1989; Wuyts et al., 1997). The cation exchanger applied consists of small, perfectly spherical MonoBeads based on a 10-mm beaded hydrophilic polystyrene/divinyl benzene resin substituted with methyl sulfonate groups

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(Mono S, GE Healthcare). The advantages of MonoBeads are its chemical (pH range from 2 to 12; resistance to organic solvents) and physical stability and high-resolution separating capacity. Affinity chromatography–purified samples are dissolved in 50 mM formate, pH 4.0, and proteins are eluted from the column in a 0 to 1 M NaCl gradient in 50 mM formate, pH 4.0, in 30 min at a flow rate of 1 ml/min.

8. Reverse-Phase High-Pressure Liquid Chromatography (RP-HPLC) In a final step, the difference in hydrophobicity between chemokines is exploited to purify chemokines to homogeneity by reverse-phase highpressure liquid chromatography (RP-HPLC) ( Janson and Ryden, 1989; Wuyts et al., 1997). A silica-based matrix with octyl C8 n-alkyl hydrocarbon derivatives is used (2.1  220-mm Brownlee C8 Aquapore RP-300 column, Perkin-Elmer, Norwalk, CT). This matrix is especially suitable for the purification of hydrophobic molecules and for the separation of closely related proteins. Because certain chemokines only elute from C18 columns at high solvent concentrations, C8 columns are preferred. Proteins that eluted from cation exchange columns in a salt gradient at pH 4.0 are loaded on the column. Columns are washed with 0.1% trifluoroacetic acid (TFA) and eluted by applying an acetonitrile gradient (0 to 80%) in 0.1% TFA at a flow rate of 0.4 ml/min (for columns with an internal diameter of 2.1 mm). Detection is performed by ultraviolet (UV) adsorption at 214 nm and/or electrospray ion trap mass spectrometry after splitting the eluate (1/150) online.

9. Ion Trap Mass Spectrometry Mass spectrometry is a powerful technique to identify the Mr of a compound on the basis of the ratio mass/charge (m/z). Here, electrospray ionization is combined with an ion trap mass analyzer (Esquire LC, Bruker Daltonics, Bremen, Germany). The sample is either diluted in 50% acetonitrile in 0.1% acetic acid and manually injected at a flow rate of 300 ml/h or online analyzed from the split outlet of the HPLC system (at a flow rate of 160 ml/h). A small diameter needle subjected to high voltage then sprays the sample. The protons from the acid render the proteins in the droplets positively charged, causing them to move toward the negatively charged instrument. By applying heat (300  C) and a flow of heated nitrogen, the droplets evaporate during the spray until the gas phase is reached. The major advantage of electrospray ionization in acidic conditions is the compatibility

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with RP-HPLC and its tendency to protonate at basic sites, which results in multiply charged ions with a proton for the NH2-terminus and for several basic residues present in the peptide (M þ nHþ). The use of detergents such as SDS should be avoided, because they render peptides negative, making them less detectable, and modify the surface tension, interfering with the evaporation of the charged droplet (Kinter and Sherman, 2000). Chemokines diluted in 0.1% acetic acid or 0.1% TFA will produce ions carrying typically 5 to 15 positive charges. Although use of acetic or formic acid results in a higher sensitivity on the mass spectrometer, TFA is often used when mass spectrometry is preceded by RP-HPLC, because gradients in TFA result in sharper elution profiles for proteins from RP-HPLC columns. A data analysis program (Bruker Daltonics) deconvolutes the spectrum and calculates the molecular mass of the uncharged protein from the multiple charged ions.

10. Edman Degradation Automated amino acid sequence analysis (Procise 491 cLC protein sequencer, Applied Biosystems, Foster City, CA) is based on the Edman degradation reaction (Edman, 1949; Edman et al., 1967; Hunkapiller et al., 1978; Kinter et al., 2000). The NH2-terminal amino acid is quantitatively coupled to phenylisothiocyanate (PITC), forming a phenylthiocarbamoyl (PTC)-derivative in the presence of N-methyl piperidine at 48  C. After removing the excess PITC with n-heptane and ethyl acetate, 100% TFA is added, resulting in the cleavage of the peptide bond between the first two NH2-terminal amino acids, leaving the second amino acid available for a new Edman degradation cycle. The PTC derivative of the NH2-terminal amino acid is released as an anilinothiazolinone (ATZ) derivative and, after extraction with 1-chlorobutane and transfer to a conversion chamber, is rapidly converted into a more stable phenylthiohydantoin (PTH) derivative in the presence of 25% TFA at 64  C. Next, the PTH derivative is loaded onto a Procise cLC PTH RP-HPLC column (0.8  250 mm) and eluted in a gradient of solvent A (3.5% tetrahydrofuran in water supplemented with 15 ml/L Premix buffer, Applied Biosystems) and solvent B (12.5% v/v isopropanol in acetonitrile) at 55  C, and UV absorbance of the PTH-amino acids is detected at 270 nm. After comparing the observed elution time with known eluting positions of a standardized mixture of PTH amino acids, the released NH2-terminal amino acid can be identified. Some chemokines are obstructed by an NH2-terminal modification, such as pyroglutamic acid for the CC chemokines CCL2, CCL7, and CCL8. PITC is unable to couple to these modified NH2-termini, which renders these proteins resistant to Edman degradation. If these proteins contain Asp-Pro sequences (which is the case for a number of chemokines), formic acid (75% at 37  C for 48 to 72 h)

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cleaves these proteins, specifically between Asp and Pro, making the released COOH-terminal peptide accessible to Edman degradation.

11. Total Protein Quantification Methods Although protein concentration can be estimated by comparison with an internal benchmark with a specific sandwich ELISA, SDS-PAGE, mass spectrometry, or Edman degradation, a chemokine preparation purified to homogeneity can also be quantified with total protein detection assays such as the Bradford or BCA assay. The Bradford protein assay uses the binding properties of the dye Coomassie brilliant blue G-250 (Bio-Rad Laboratories) (Bradford, 1976). When the unbound red/brown dye binds to proteins, a conformational change occurs in its structure resulting in a spectral shift from an absorption maximum of 465 nm (green) to 595 nm (blue). The optimal wavelength to measure the blue color from the Coomassie dye-protein complex is 595 nm, because the difference between the two forms of the dye is then greatest. However, the Coomassie dye is incompatible with surfactants, excluding the technique for samples reconstituted with solubilizing detergent. In practice, sample dilutions and dilution series of bovine serum albumin (BSA) are prepared in a 96-well plate (100 ml/well) and 200 ml of Coomassie dye (1/5 diluted in DPBS) is added. Absorption can be measured immediately at 595 nm. In addition, the bicinchoninic acid (BCA) assay is prevalently used, a method based on the biuret reaction (Smith et al., 1985). In alkaline medium, peptides of 3 amino acids or larger reduce Cu2þ into Cu1þ forming a blue-colored chelated complex. Such complex is formed between one Cu1þ ion and four to six nearby peptides bonds, which is proportional to the intensity of the color. BCA selectively and with high sensitivity recognizes these Cu1þ ions, resulting in a purple color complex. The optimal wavelength is 562 nm, where a strong linearity exists between the absorbance and the protein concentration. Dilution series of BSA, starting at a concentration of 2 mg/ml, and chemokines are prepared in a volume of 25 ml in a 96-well plate. Next, 200 ml of dye (1/50 reagent B/A) (Pierce, Thermo Fischer Scientific) is added to each well. After 30 min of incubation at 37  C, the absorbance is measured at 562 nm. Because colorimetric development in most assays is related to the amino acid composition of the protein, it is advisable to apply multiple quantification methods in parallel to avoid misinterpretation. In the BCA assay, the presence of cysteine, cystine, tyrosine, and tryptophan can influence the outcome of the assay, whereas in the Bradford assay, the relative abundance of the amino acids arginine, lysine, and histidine is important.

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12. Illustration: Isolation and Identification of Natural Posttranslationally Modified CXCL8 Isoforms PBMC from 24 buffy coats were pooled (11.4  109 cells) and induced at 5  106 cells/ml with 10 mg/ml of the TLR3 ligand and double-stranded RNA polyriboinosinic: polyribocytidylic acid (polyrI:rC) and 20 ng/ml interferon-g (IFN-g) in RPMI 1640 containing 2% FCS. CXCL8 was isolated from the conditioned medium by described four-step purification method. After adsorption to silicic acid, the eluate was subjected to heparin Sepharose chromatography (GE Healthcare) (Fig. 1.1A). Besides CXCL8 protein, other chemokines were also detected such as CCL2, CCL7, CCL8, CXCL4, CXCL9, CXCL10, and CXCL11 as evidenced by specific sandwich ELISA (data not shown). Heparin Sepharose column fractions 7 to 9, which contain the most CXCL8 immunoreactivity, were pooled and loaded on a Mono S cation exchange column (GE Healthcare). CXCL8 eluted from the column at approximately 0.75 M NaCl after 63 min, as evidenced by ELISA (Fig. 1.1B). Fractions 64 to 66 (elution time 63 to 66 min) (Fig. 1.1C) and 67 to 70 (elution time 66 to 70 min) (data not shown) recovered from the ion exchange column were loaded on a 2.1  220-mm Brownlee C8 Aquapore RP-300 HPLC column (PerkinElmer) in 0.1% TFA. CXCL8 isoforms eluted from the column at approximately 31% acetonitrile and were collected in 1-min fractions. Online mass spectrometry showed the presence of three major proteins eluting from the RP-HPLC column after 55 to 61 min (Fig. 1.1D). Deconvolution of the spectra resulted in the detection of three CXCL8 isoforms in fraction 56 that were differently processed at their NH2-terminus (i.e. CXCL8[2 to 77], CXCL8[1 to 77], and CXCL8[6 to 77]). Additional analysis of the proteins in fraction 56 by Edman degradation resulted in the detection of a modified Arg in position 5 for CXCL8(1 to 77) (Fig. 1.2). Because the experimentally determined Mr of CXCL8 was not significantly different from the expected Mr, modification of Arg to citrulline (Cit) was considered. Cit is only 1 mass unit larger than Arg, and on the total Mr of CXCL8, this would fall within the accuracy of the mass spectrometer. L-Cit was subjected to Edman degradation, and PTH-Cit was found to elute at exactly the same position as the modified Arg (i.e. in between PTH-Thr and PTH-Gly). Analysis of the other RPHPLC fractions that contained CXCL8 immunoreactivity (fractions 56 to 61) resulted in the identification of several citrullinated and/or NH2terminally truncated CXCL8-forms (Fig. 1.3). Citrullination was only detected on Arg5 and not on other Arg in natural CXCL8 (Proost et al., 2008).

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Tamara Loos et al.

250 200

2500

100 50

0 <2.5

0

5

10

15

20

0

25

[NaCl] (M) (---)

5000

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10,000

CXCL8 (ng/ml) (

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10,000

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<25 0

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Retention time (min)

% acetonitrile (---)

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Intensity ⫻ 106

)

20

D

C

U.V.214 nm(mAU) (

10

8380.9 Mr (A) CXCL8 (6−77)

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1.0

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)

)

2

)

B

>12,500

[NaCl] (M) (---)

A

8800

9200

0.4 0.2 0.0

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Mr

Retention time (min)

Figure 1.1 Purification, isolation, and identification of CXCL8 isoforms. PBMC from 24 buffy coats were pooled (11.4 109 cells) and induced at 5 106 cells/ml with 10 mg/ml of polyrI:rC and 20 ng/ml IFN-g in RPMI 1640 containing 2% FCS. (A) After adsorption to silicicacid,theeluatewassubjectedtoheparinSepharosechromatography(GEHealthcare). Proteins eluted from the column by applying a NaCl gradient. Total protein (mg/ml) was measuredwiththeBradfordassay.CXCL8proteinconcentrationwasevaluatedbyaspecific sandwich ELISA. (B) Fractions 7 to 9 that eluted from the heparin Sepharose column were loaded on a Mono Scation exchange column (GE Healthcare) in 50 mM HCOOH. CXCL8 eluted from the column at approximately 0.75 M NaCl after 63 min, as evidenced by ELISA. (C) Fractions 64 to 66 recovered from the ion exchange chromatographic step were loaded on a RP-HPLC column (2.1  220-mm Brownlee C8 Aquapore RP-300 column, PerkinElmer) in 0.1% TFA. CXCL8 isoforms eluted from the column at approximately 31% acetonitrile. (D) Online mass spectrometry showed the presence of three proteins eluting from the RP-HPLC column after 55 min. The deconvoluted mass spectra corresponded to the Mr of the isoforms CXCL8(2 to 77), CXCL8(1 to 77), and CXCL8 (6 to 77).

13. Solid-Phase Peptide Synthesis To study the characteristics of natural posttranslationally modified chemokines, the availability of sufficient and pure material is an obvious requirement. Because natural sources, in general, do not supply sufficient amounts and because some natural chemokine isoforms cannot be easily separated by conventional chromatography, other approaches were mandatory to obtain sufficient material for biologic assays. One option is solidphase peptide synthesis of chemokines, which provides a good alternative

15

Posttranslational Modification of Chemokines

A E

5 4

Q

D N

3

A

TG

Y

V PM

W

F

IK L

R

H

S

2

U.V.270 nm (mAU)

B

1 (AA) 4 L

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5 4

3 P

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V

KL 3

E

1 A

C 5 4 3 2 1 0

Cit

6

8

10

12 14 16 Retention time (min)

18

20

Figure 1.2 Identification of naturally citrullinated CXCL8 by Edman degradation. (A) RP-HPLC chromatogram detected at 270 nm (mAU, milli absorption units) of the 19 PTH-amino acids (indicated by their one letter code). (B) Natural CXCL8 was subjected to Edman degradation. Overlays demonstrate PTH-derivatives detected after 3, 4, and 5 cycles of Edman degradation, revealing the amino acid sequences -KEL-from CXCL8 (6 to 77), -AVL- from CXCL8 (2 to 77), and -LPX- from CXCL8 (1 to 77) with X assigned to an unidentified amino acid. (C) L-Cit was loaded on the reaction vessel of the protein sequencer and the PTH-derivative was analyzed by RP-HPLC. PTH-Cit eluted in between PTH-Thr and PTH-Gly, at exactly the same position as the unidentified amino acid (X) in natural CXCL8.

for the time-consuming production by recombinant expression. Moreover, recombinant proteins are more likely to be contaminated with other potent proinflammatory molecules of biologic origin such as the TLR ligands. Moreover, incorporation of amino acids that are not encoded by DNA (e.g., citrulline, hydroxyproline,. . .) or chemical groups is impossible.

13.1. Synthesis of the peptide chain During solid-phase peptide synthesis, the peptide is assembled from the COOH-terminus towards the NH2-terminus. The a-carboxyl group of the COOH-terminal amino acid is attached to a stable and solid support

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Tamara Loos et al.

125

Arg5 Cit5

100 75 50

56 57 58 59 60 61

56 57 58 59 60 61

CXCL8 (1−77)

56 57 58 59 60 61

CXCL8 (−2−77)

56 57 58 59 60 61

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25 56 57 58 59 60 61

CXCL8 (pmol)

150

CXCL8 (2−77)

CXCL8 (6−77)

CXCL8 (8−77)

CXCL8 (9−77)

RP-HPLC fraction number

Figure 1.3 Abundance of CXCL8 isoforms in PBMC. Edman degradation on RP-HPLC fractions 56 to 61 (Fig.1.1C) lead to the identification of the CXCL8 isoforms CXCL8(2 to 77), CXCL8(1 to 77), CXCL8(2 to 77), CXCL8(6 to 77), CXCL8(8 to 77), and CXCL(9 to 77). Moreover, posttranslational modification of Arg5 into Cit5 was discovered on part of the CXCL8(2 to 77), CXCL8(1 to 77), and CXCL8(2 to 77) proteins (filled histograms). Histograms indicate the amount of each CXCL8 isoform in the individual RP-HPLC fractions.

(i.e., HMP-resin [4-hydroxymethyl-phenoxy-methyl-polystyrene, crosslinked by 1% divinylbenzene]), and they remain coupled during chain assembly. In this manner, the peptide can be easily separated from used reagents and solvents by simple filtration and washing. One by one the consecutive amino acids are coupled to the growing peptide chain, according to the amino acid sequence of the desired protein. The a-amino group of these amino acids is protected from inappropriate binding by a fluorenylmethoxy carbonyl–(Fmoc) protecting group (Atherton and Sheppard, 1989). Moreover, such a protecting group destroys the amino acid’s zwitterionic character. Instead of the base-labile Fmoc protecting group, an acid-labile tertiary-butoxycarbonyl (tBoc) is also widely used (Clark-Lewis et al., 1997). Although successful chemokine synthesis has been achieved with this strategy, the use on the synthesizer of concentrated TFA for the removal of tBoc groups and strongly corrosive hydrofluoric acid during final cleavage and deprotection reactions make it less attractive (Clark-Lewis et al., 1991). Because the side chains of some amino acids also contain chemically reactive groups, hindering clear-cut peptide bond formation, they need to be blocked by protecting groups as well (Atherton and Sheppard, 1989). The choice of the side chain–protecting group relies on the kind of amino acid and the synthesis strategy. The side chain–protecting groups used in our laboratory are t-butyloxycarbonyl for lysine; tert-butyl for serine, threonine, and tyrosine; 2,2,7,8-pentamethylchroman-6-sulfonyl for arginine; tert-butyl-ester for aspartic acid and glutamic acid; and trityl for histidine, cysteine, asparagine, and glutamine (Proost et al., 1995). During peptide synthesis, a portion of the peptide resin can be taken away and stored, while synthesis proceeds on the remaining peptide-resin complexes. This allows for the generation of different NH2-terminally

Posttranslational Modification of Chemokines

17

modified analogs with a common COOH-terminal sequence. Conversely, analogs with a different COOH-terminus have to be synthesized separately. Detailed synthesis protocol Initially, a threefold molar excess (compared with the amount of active groups on the resin) of the COOH-terminal amino acid of the desired protein is activated by 1 M N,N0 -dicyclohexylcarbodiimide in N-methyl pyrrolidone (DCC/NMP) generating an activated ester and subsequently added to the resin in the reaction vessel together with the basic esterification catalyst dimethyl aminopyridine (DMAP; 0.1 M in dimethylformamide [DMF]). Accordingly, the carboxyl group of the COOH-terminal amino acid is coupled to a hydroxyl group of the HMP resin by a symmetric anhydride binding. To prevent coupling of HBTU-activated amino acids at a later stage to unloaded hydroxyl groups, remaining hydroxyl groups are capped with benzoic anhydride, again with DMAP as a catalyst. Subsequently, the succeeding amino acids are coupled one by one to the growing peptide chain (0.1 mmol to 0.25 mmol of peptide resin) (Fig. 1.4). As a first step of chain assembly, the Fmoc protection group must be removed from the resin-coupled amino acid. Deprotection is carried out by fourfold treatment with piperidine (20%) in NMP, which results in the removal of the Fmoc group on the basis of a b-elimination reaction. After each deprotection step, the resin particles in the reaction vessel are washed with NMP to remove the remaining piperidine. The penultimate Fmocprotected amino acid, which needs to be coupled to the resin-bound amino acid, is activated by 2-1H-benzotriazol-1yl-1,1,3,3-tetramethylureniumhexafluorophosphate (HBTU 0.45 M )/1-hydroxybenzotriazole (HOBt 0.45 M ) in DMF. Subsequently, N,N-diisopropyl ethylamine (DIEA; 2 M in NMP) is added, and the HBTU-activated amino acid (1 mmol; 4- to 10-fold excess) is transferred to the reaction vessel to form a peptide bond with the amino acid–resin complex. Because coupling is never 100% complete, the remaining free a-amino groups of the first amino acid residues are capped with acetic anhydride (0.5 M in NMP containing 0.125 M DIEA and 0.015 M HOBt) to prevent them from being coupled to amino acids added during one of the following cycles, which would result in the synthesis of proteins internally lacking one or more amino acids. Finally, the resin particles are washed four times with NMP. These deprotection and coupling steps are repeated for every single amino acid until the entire peptide chain has been completed. In the end, the resincoupled peptide chain is treated once again with piperidine to remove the final Fmoc group from the NH2-terminal amino acid (Fig. 1.4). As the peptide chain grows, interchain and intrachain interactions are more likely to occur. As a result, the NH2-terminus of the peptide may be buried and less accessible, obviously making it harder to achieve efficient deprotection and coupling. Removal of the Fmoc group can be monitored

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O

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-H2N-CH-C-...-NH-CH-C-NH-CH-C-O - resin Rn

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R1 X TFA + thioanisole, water, 1,2-ethanedithiol and crystalline phenol O O

H2N-CH-C-...-NH-CH-C-NH-CH-C-OH Rn

R2

R1

Figure 1.4 General protocol for solid-phase peptide synthesis. During solid-phase peptide synthesis, the COOH-terminal amino acid is stably attached to the HMP-resin. Both the a-amino group and reactive side chains of amino acids are blocked by protecting groups. After loading the resin with the COOH-terminal amino acid, the Fmoc group is removed from the a-amino group by treatment with piperidine, allowing for coupling of the next amino acid. HBTUsolution is added to a cartridge containing the next amino acid. Activated Fmoc-amino acid is formed almost instantaneously and transferred to the reaction vessel, where a peptide bond is formed with the resin-coupled amino acid. Afterwards, uncoupled free a-amino groups are capped by acetic anhydride.This cycle is repeated for each of the following amino acids until the last amino acid has been coupled. The base-labile Fmoc group of the NH2 -terminal amino acid is removed by a final treatment with piperidine, whereas the acid-labile side chain protecting groups and peptide^ resin bond are cleaved withTFA.Water, ethane dithiol, thioanisole, and crystalline phenol are used as scavengers, limiting modifications of the side chains. Rx represents a random amino acid side chain. X,Y, and Z stand for side chain protecting groups.

Posttranslational Modification of Chemokines

19

by UV or conductivity measurements after every piperidine treatment. Hence, if removal of the Fmoc group does not pass very efficiently, the deprotection is automatically prolonged by conditional deprotection modules (Fig. 1.5). Moreover, these conditional deprotection modules are followed by double, instead of single, coupling. An extra amount of amino acid (1 mmol) is activated and transferred to the reaction vessel for prolonged coupling to increase the coupling yield.

13.2. Deprotection of the synthesized peptide chain After synthesis, the resin and the side chain–protecting groups are to be cleaved from the synthetic peptide (Fig. 1.4). Therefore, the peptide is incubated during 1.5 to 2.5 h in a mixture of 10 ml TFA, 0.5 ml thioanisole, 0.5 ml deionized water, 0.25 ml 1,2-ethanedithiol, and 0.75g crystalline phenol under continuous shaking. TFA cleaves the peptides from the resin and removes the side chain–protecting groups, whereas ethane dithiol, thioanisole, water, and phenol are added to keep the side chains of Trp, Tyr, Met, and Cys from being modified by released protecting groups (King et al., 1990). Afterwards, the resin particles are eliminated by filtering the TFA solution through a Biospin filter (Bio-Rad laboratories). The proteins in the filtrate are precipitated in 30 ml cold diethyl ether. After centrifugation, the protein-containing pellet is washed several times with diethyl ether to remove remaining chemicals. The precipitate is dissolved in H2O, lyophilized, and redissolved in 0.1% TFA and purified by RP-HPLC (Source 5 RPC column; GE Healthcare). Proteins are detected by online UV (l ¼ 220 nm) and ion trap mass spectrometry (Esquire LC, Bruker Daltonics). Fractions containing the intact raw protein were lyophilized before the folding procedure.

13.3. Folding of the raw protein To obtain disulfide bridges, the intact linear protein is incubated in 150 mM TRIS (tris[hydroxymethyl]) aminomethane], pH 8.6, containing 3 mM EDTA (ethylenediamine tetraacetic acid), 0.3 mM reduced glutathione, 3 mM oxidized glutathione, and 1 M guanidinium chloride. After acidification, the folded material is purified by RP-HPLC with a C8-Aquapore RP-300 column (2.1  220 mm; PerkinElmer) combined with online detection by mass spectrometry. Finally, Edman degradation on a 491 Procise cLC protein sequencer (Applied Biosystems) and ion trap mass spectrometry are used to confirm the NH2-terminal sequence and the Mr of the folded chemokine. The concentration of the synthesized chemokine is determined with the bicinchoninic acid assay and the yield of the individual amino acids after Edman degradation.

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Tamara Loos et al.

Piperidine deprotection Deprotection Deprotection Deprotection Deprotection

1 2 3 – U.V./conductivity 4 – U.V./conductivity

Difference > threshold = inefficient deprotection Difference < threshold = efficient deprotection

Inefficient deprotection

Efficient deprotection

Fifth deprotection

Extended deprotection

Activation of amino acid

Transfer

Coupling

Inefficient deprotection Extended coupling

Activation of additional amino acid Efficient deprotection Transfer

Extended coupling

Acetic anhydride capping

Figure 1.5 Incorporation of conditional modules in the general protocol of automated solid-phase peptide synthesis. During peptide chain synthesis, amino acids are coupled one by one to the growing peptide chain, directed from COOH- to NH2 -terminus. Each amino acid (except for the COOH-terminal amino acid) is coupled following the same procedure as described in this scheme. Some modules are conditional, meaning that they are only turned on/off under user-defined conditions.Whether or not the conditional modules become active is determined by UVor conductivity measurements on the deprotection solutions after piperidine treatment. When the UV or conductivity

Posttranslational Modification of Chemokines

21

Special remarks considering undesired side reactions 

Attention needs to be paid to the synthesis of proteins carrying a proline at their COOH terminus. Fmoc-proline can be loaded to HMP resin, but a potential diketopiperazine side reaction occurs during the chain assembly that can drastically reduce the yield of final peptide resin (Gisin et al., 1972; Proost et al., 1995).  Proteins bearing a DG sequence are prone to undergo base-mediated aspartimide formation during the repetitive piperidine treatments (Do¨lling et al., 1994; Lauer et al., 1994; Nicola´s et al., 1989; Yang et al., 1994). On nucleophilic attack by water or piperidine these aspartimides readily undergo ring opening, resulting in the generation of either a- and b-aspartyl peptides or N-aspartyl piperidine. We observed this side reaction during the synthesis of CXCL8 (Fig. 1.6). Verifying the Mr of the synthesized CXCL8(6 to 77) by mass spectrometry revealed a difference of 67 mass units with the theoretical molecular weight. This problem can be overcome by use of the preformed dipeptide Fmoc-Asp(OtBu)(Dmb)Gly-OH (Novabiochem, EMD Chemicals, Gibbstown, NJ), which masks the Asp-Gly amide bond, protecting it against aspartimide formation (Mergler et al., 2003; Packman, 1995).

14. Citrullination of Chemokines Citrullination is an irreversible reaction in which peptidyl arginine is deiminated and subsequently hydrolyzed into peptidyl citrulline, resulting in a mass increase of one mass unit and the loss of one positive charge. The enzymes responsible for this conversion are the calcium-dependent peptidyl arginine deiminases (PAD) (Vossenaar et al., 2003). Calcium ions bind to the enzyme, and a conformational change occurs. Consequently, it is speculated that the central cysteine residue in the catalytic domain becomes accessible and its thiol group attacks the guanidino group of arginine, releasing ammonia and forming a tetraheder intermediate. This intermediate structure is then hydrolyzed by water into citrulline (Arita et al., 2004). Chemokine citrullination was recently discovered on natural CXCL8 and CXCL10 and was shown to severely affect the biologic activities of chemokines (Loos et al., 2008; Proost et al., 2008). Chemokines may be measurement after the third deprotection step differs by more than a threshold value (typically 2.5% or 5%) from the UV or conductivity measurement after the fourth deprotection step, conditional modules are switched on. Thus, in case of inefficient deprotections, the conditional modules (underlined) are activated, aiming at higher deprotection and coupling efficiencies and a higher synthesis yield.

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Tamara Loos et al.

A

Intensity x 106

10+ 846.3 1.50

11+ 769.4

Intensity ⫻ 106

1.25

9+ 940.2

4 3 2 1

1.00

0

12+ 705.5

0.75

8453.3

5

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8+ 1057.8

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7+ 1208.3

13+ 0.25 651.2

0.00 700

800

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1300

1400 m/z

B 11+ 763.4 2.0

8386.0 Intensity ⫻ 106

10+ 839.6

12+ 699.9

Intensity ⫻ 106

1.5 9+ 932.7

8 6 4 2 0

8300 8350

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8450

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1.0 13+ 646.1 8+ 1049.2

0.5

7+ 1198.8

6+ 1398.6

0.0 700

800

900

1000

1100

1200

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1400 m/z

Figure 1.6 N-aspartyl piperidine formation of the DG sequence during synthesis of CXCL8(6 to 77) can be overcome by use of the preformed dipeptide Fmoc-Asp (OtBu)-(Dmb)Gly-OH. CXCL8, a chemokine bearing an internal Asp-Gly sequence, was synthesized on a solid-phase peptide synthesizer with Fmoc chemistry. (A) Ion trap mass spectrum of purified TFA-deprotected CXCL8(6 to 77), synthesized following the conventional method with Fmoc-Gly and Fmoc-Asp(OtBu). The Mr of the synthetic proteins is calculated by deconvolution of the multiply charged ions in the raw spectrum and is shown as an insert. The Mr of the synthesized CXCL8(6 to 77) in this case differs by 67 mass units from the expected theoretical Mr of unfolded CXCL8(6 to 77)

Posttranslational Modification of Chemokines

23

citrullinated in vitro by enzymatic treatment to investigate the biologic implications of chemokine citrullination. Citrullination is performed by incubating proteins with PAD in 40 mM TRIS, pH 7.4, supplemented with 2 mM CaCl2 at 37  C. Deimination is stopped with 0.1% TFA, because PAD activity is pH dependent (pH 6 to 9) (Nakayama-Hamada et al., 2005). Next, the citrullinated proteins are purified by RP-HPLC (1  50-mm Brownlee C8 Aquapore RP-300 column, PerkinElmer). To reveal the appropriate incubation period, kinetic studies can be performed. For example, 100 pmol CXCL8 (PeproTech, Rocky Hill, NJ) was incubated with rabbit PAD (Sigma-Aldrich) or with human PAD2 or PAD4 (ModiQuest Research, Nijmegen, The Netherlands) at an enzyme-substrate molar ratio (E/S) of 1:20 or 1:200 for different time periods. After ending deimination, samples were split and desalted on C4 ZipTip (Millipore) before mass spectrometry or in parallel spotted on PVDF membranes (ProSorb; Applied Biosystems) before Edman degradation.

15. Identification of Enzymes Generating the Natural Posttranslationally Modified Chemokines, as Exemplified by Aminopeptidase N(APN)/CD13 To uncover which enzymes may be involved in the generation of certain naturally purified proteolytically processed chemokines, chemokines can be incubated in vitro with potential candidate enzymes, followed by analysis with mass spectrometry. Indeed, incubation of CD26-processed CXCL11(3 to 73) with APN/CD13 generated the same CXCL11 isoforms as purified from natural sources (i.e. CXCL11[4,5,6 to 73]) (Proost et al., 2001, 2007). APN/CD13 is a metalloprotease that removes NH2-terminal amino acids one by one, except for proline, which is resistant to APN/CD13 cleavage (Ashmun et al., 1990; Breljak et al., 2003; Riemann et al., 1999). To investigate in vitro processing by APN/CD13, CXCL11(3 to 73) was incubated for 2 h at 37  C in phosphate-buffered saline (DPBS) with porcine kidney purified microsomal APN/CD13 (Sigma-Aldrich) at an enzymesubstrate molar ratio of 1:4 or 1:25. To prevent CXCL11 from sticking to the plastic tube, the nonionic detergent octyl-b-glucopyranoside (0.1%) was (Mr ¼ 8386 Da). This is due to aspartimide formation during piperidine deprotection, whereupon a nucleophilic attack by piperidine leads to the generation of an N-aspartyl piperidine. (B) The mass spectrum of TFA-deprotected CXCL8(6 to 77) synthesized with the preformed dipeptide Fmoc-Asp(OtBu)-(Dmb)Gly-OH. This spectrum demonstrates that on use of the preformed DG dipeptide, a protein with the correct Mr is synthesized, and no N-aspartyl piperidine formation is observed.

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Tamara Loos et al.

added. The enzymatic reaction was stopped by the addition of 0.1% TFA. Samples were desalted and concentrated with C4 Zip Tips (Millipore) and analyzed by ion trap mass spectrometry. Mass spectrometric analysis after incubation unexpectedly revealed NH2-terminal and COOH-terminal processing of CXCL11. However, when the serine protease inhibitors phenylmethylsulfonyl fluoride (PMSF; 30 mM) or benzamidine (5 mM) were included, COOH-terminal cleavage was blocked, pointing toward the presence of contaminating serine proteases in the commercially available APN/CD13 (Proost et al., 2007). Therefore, it is recommended to add a serine protease-inhibitor, such as PMSF or benzamidine, to the reaction mixture.

16. Comparison of the Heparin-Binding Properties of Chemokine Isoforms Chemokines play a prominent role in directing selective leukocyte recruitment during inflammation and normal immune surveillance. In addition to chemokine receptors, glycosaminoglycans (GAG) are key players in this process. They may drive transcytosis of chemokines across the endothelial cell layer, immobilize chemokines on the luminal surface of the endothelium, and form an immobilized gradient directing leukocytes to the site of inflammation (Colditz et al., 2007; Johnson et al., 2005; Middleton et al., 2002; Parish, 2006). Cell-surface GAG may also induce polymerization of chemokines, increasing their local concentration and, therefore, enhancing their effects on high-affinity receptors within the local environment (Hoogewerf et al., 1997). Hence, besides binding to and signaling through seven-transmembrane-spanning G protein–coupled receptors, inducing well-known effects such as an increase in the intracellular calcium concentration and chemotaxis, chemokine-GAG interaction is important for the in vivo biologic activity of chemokines. The chemokineGAG interaction has been thought to be based on electrostatic interactions between the highly negatively charged sulfated polysaccharides and the largely basic COOH terminus of the chemokine. However, further investigation revealed that this does not completely explain chemokine-GAG interactions (Handel et al., 2005). Because chemokine-GAG interaction seems to play a significant role in in vivo leukocyte migration, studying the effect of posttranslational modifications on this interaction might be very interesting. A technique for the identification or characterization of heparin-binding proteins has been developed recently (Mahoney et al., 2004); 96-well plates are treated by plasma polymerization with allylamine that leads to a change in the surface of microtiter plates, allowing heparin to be immobilized without being modified

Posttranslational Modification of Chemokines

25

(Heparin binding plates; BD, Franklin Lakes, NY). This constitutes the main advantage of this technique. Because there is no need to modify the GAG molecule, it can fully retain its ability to interact with other biomolecules. The assay for the characterization of binding of molecules to the immobilized heparin is similar to an ELISA. To immobilize heparin molecules on the plasma-polymerized coating, each well is incubated overnight with 100 ml of a low molecular weight heparin solution (25 mg/ml in DPBS; Sigma-Aldrich) at room temperature (protected from light). After three wash steps with standard assay buffer (SAB; 100 mM NaCl, 50 mM NaAc, 0.2% [v/v] Tween-20, pH 7.2) to remove unbound heparin, 250 ml of SAB enriched with 0.2% (w/v) gelatin (blocking solution) is added to each well and the plate is blocked at 37  C for 1 h. Chemokine-solutions (100 ml, diluted in blocking solution) are added and allowed to interact with heparin for 2 h at 37  C. Unbound chemokine is removed by washing the plate 3 times with SAB. Bound chemokine is detected by adding 100 ml of a specific biotinylated antichemokine antibody in blocking solution for 1 h at 37  C. The plate is washed three times with SAB, and 100 ml of HRP-labeled streptavidin ( Jackson ImmunoResearch Laboratories, West Grove, PA) is added. After 30 min, unbound streptavidin is removed by washing the plate 3 times with SAB. To produce a visible signal quantifying the peroxidase activity, 100 ml of a chromogenic HRP-substrate solution is added (cf. ELISA). Used biotinylated antibodies need to be checked for equal recognition of the native chemokine as well as the posttranslationally modified chemokines. Percentage binding was calculated by subtracting the mean OD of the negative control (blocking buffer) from the measured OD of the sample, subsequent division with the average OD of the highest concentration of intact chemokine and finally multiplying by 100. This calculation grants a 100% binding to the highest concentration of intact chemokine and 0% binding to the negative control.

ACKNOWLEDGMENTS This work was supported by the Center of Excellence (Credit no. EF/05/15) of the K. U. Leuven, the Concerted Research Actions (G.O.A./2007/15) of the Regional Government of Flanders, the Fund for Scientific Research of Flanders (F.W.O.-Vlaanderen), the Interuniversity Attraction Poles Program-Belgian Science Policy (I.A.P.), and the European Union 6FP EC contract INNOCHEM (grant LSHB-CT-2005-518167). A. M. is a research assistant of the F.W.O.-Vlaanderen.

REFERENCES Arita, K., Hashimoto, H., Shimizu, T., Nakashima, K., Yamada, M., and Sato, M. (2004). Structural basis for Ca(2þ)-induced activation of human PAD4. Nat. Struct. Mol. Biol. 11, 777–783.

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C H A P T E R

T W O

Homo- and Hetero-Oligomerization of Chemokines Ariane Jansma, Tracy M. Handel, and Damon J. Hamel Contents 1. Introduction 2. Methods to Detect and Quantify Oligomerization 2.1. Analytical ultracentrifugation (AUC); sedimentation equilibrium 2.2. Pulsed-field gradient diffusion by NMR 2.3. Dynamic light scattering 2.4. Fluorescence polarization 2.5. FT-ICR mass spectrometry 2.6. Other methods 3. Methods for Collecting Residue-Specific Information on Chemokine Oligomers 3.1. NMR: Heteronuclear single quantum correlation (HSQC) spectroscopy 3.2. NMR: Detection of nuclei in close proximity by means of the nuclear Overhauser effect (NOE) 4. Conclusions Acknowledgments References

32 35 35 39 40 41 42 42 43 43 44 47 47 47

Abstract Chemokines function in cell migration by binding and activating seven transmembrane G protein–coupled receptors (GPCRs) on leukocytes and many other diverse cell types. The extracellular binding event stabilizes specific conformations of the receptor that trigger cascades of intracellular signaling pathways involved in cell movement and activation (Baggiolini, 1998; Baggiolini et al., 1997; Charo and Ransohoff, 2006; Hartley et al., 2003; Kunkel and Butcher, 2002; Loetscher and Clark-Lewis, 2001). Although the current consensus is that monomeric forms of chemokines are necessary for receptor binding to induce

Skaggs School of Pharmacy and Pharmaceutical Science, University of California, San Diego, La Jolla, California, USA Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05402-0

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cell migration, oligomeric states of chemokines may be associated with other complex functional roles such as regulation, haptotactic gradient formation, protection from proteolysis, and signaling related to processes distinct from migration. Accordingly, diverse biophysical methods have been used to identify and characterize the details of these quaternary interactions. This chapter aims to summarize these methods and to provide guidelines for their application in future studies.

1. Introduction Chemokines are small (
8 to 12 kDa) secreted proteins that have been classified into four subfamilies (CC, CXC, CX3C, and C) on the basis of the relative position of their conserved N-terminal cysteine residues. All chemokines share a highly conserved monomeric structure consisting of a disordered N-terminal region, followed by an irregular ‘‘N-loop’’, three antiparallel b-strands, and a C-terminal a-helix (Fig. 2.1A) (Blain et al., 2007; Czaplewski et al., 1999; Fernandez and Lolis, 2002; Jin et al., 2005). Chemokine quaternary structure is more varied. Two primary structural

Figure 2.1 Examples of chemokine structures. (A) CCL2/MCP-1 monomer (PDB code 1DOL). (B) CCL2 dimer, an example of a ‘‘CC dimer’’ (PDB code 1DOM). The interface is composed of a small antiparallel b-sheet formed from residues at the N-terminus of both subunits. (C) IL-8/CXCL8 dimer, an example of a ‘‘CXC dimer’’ (PDB code 1IL8).The interface is formed by the first beta strand in each subunit, as well as interactions between the C-terminal end of the helix of one subunit and the b-sheet of the opposing subunit. (D) The lymphotactin/XCL1 dimer contains a unique allb-sheet structure that exists in equilibrium with a canonical chemokine monomer (PDB code 2JP1). (E) The CCL2 tetramer has characteristics from both CC and CXC dimer interfaces (adapted from PDB code 1DOL). (F) H-form of IP-10/CXCL10 tetramer associating through the third b-strands forming a 12-stranded antiparallel b-sheet with a sharp kink in the middle (PDB code1O80).

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types of dimers have been observed: CC dimers that interact by means of a two-stranded antiparallel b-sheet near the N-terminus, and CXC dimers formed by the first strand of the b-sheet from each monomer (Fig. 2.1B, C) (Blain et al., 2007; Czaplewski et al., 1999; Fernandez and Lolis, 2002; Jin et al., 2005). More recently, a third type of dimer was identified for the C chemokine, lymphotactin/XCL1, that consists of an all-b-sheet arrangement with no similarity to other known protein structures and that rapidly interconverts with the canonical monomeric chemokine fold (Fig. 2.1D) (Tuinstra et al., 2008 and chapter 3 by Brian Volkman). In addition, whereas the known quaternary structure of dimers are similar among members of each subfamily, some chemokines (e.g., MPC-1/CCL2, PF4/CXCL4, IP-10/CXCL10) have been shown to form tetramers that in some cases have both CC and CXC interfaces (Czaplewski et al., 1999), whereas others adopt completely novel folds (e.g., IP-10/CXCL10) (Fig. 2.1E, F) (Swaminathan et al., 2003). Finally, several chemokines form heterodimers, and it has been shown that heterodimerization can occur within members of a given subfamily, as well as between subfamilies (Crown et al., 2006; Paoletti et al., 2005; von Hundelshausen et al., 2005). Although, the current understanding of the functional relevance of these various oligomeric forms is far from complete, there is ample evidence suggesting that oligomerization is important to the overall mechanism of cell migration. Previous studies have shown that mutant forms of chemokines that are unable to oligomerize are generally still fully functional with respect to receptor binding and cell migration in vitro, suggesting that receptor binding occurs through the monomeric form (Czaplewski et al., 1999; Lowman et al., 1997; Paavola et al., 1998; Proudfoot et al., 2003). Nevertheless, these monomeric mutants are inactive in vivo when tested in an intraperitoneal recruitment assay (Campanella et al., 2006; Proudfoot et al., 2003, and see Chapter 18 by A. Luster). The prevailing explanation for these apparently anomalous results is related to the fact that as part of the mechanism for providing directional cues, chemokines are maintained near sites of production by localization on cell surfaces through interactions with glycosaminoglycans (GAGs). These interactions often involve oligomerization of chemokines on the GAGs. Indeed, biochemical and biophysical studies indicate that some chemokine: GAG interactions are facilitated by chemokine oligomerization and also that chemokine oligomerization can be facilitated by interactions with GAGs (Hoogewerf et al., 1997; Lau et al., 2004; Proudfoot et al., 2003). Furthermore, chemokine variants that are incapable of binding GAGs can also be functional in vitro but not in vivo in much the same way as mutants that are unable to oligomerize (Proudfoot et al., 2003). The functional coupling between GAG binding and oligomerization is perhaps best exemplified by the [44AANA47]-RANTES/CCL5 mutant, which is unable to bind GAGs and blocks the activity of the WT protein through a dominant negative effect by forming nonfunctional

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heterodimers with the WT protein ( Johnson et al., 2004; Lau et al., 2004). Although chemokine: GAG interactions are not discussed further here, many reviews are available on this topic (Allen et al., 2007; Fermas et al., 2008; Handel et al., 2005; Imberty et al., 2007; Johnson et al., 2005; Kuschert et al., 1999; Lau et al., 2004; Witt and Lander, 1994; Yu et al., 2005, and chapter 4 in this volume present various methods for characterizing these interactions). In addition to playing a role in cell migration, oligomerization may also contribute to the functional regulation of chemokines. For example, the disordered N-termini are the key signaling domains in all chemokines, and thus proteolytic processing of the N-terminus represents a natural mechanism for modulating chemokine function (Fox et al., 2006). Most frequently, agonist activity is reduced or abolished completely on N-terminal proteolysis; however, there are examples of increased activity, and even alterations in receptor-binding specificity with N-terminal processing (Fox et al., 2006; Homey et al., 2002). Although interactions with GAGs have clearly been shown to protect chemokines from proteolysis (Ellyard et al., 2007; Vives et al., 2002), in principle, oligomerization alone could also be protective, both directly and indirectly, by facilitating interactions with GAGs. Finally, oligomerization has been shown to influence cellular signaling. In some cases, oligomerization promotes additional signaling pathways not induced by the monomeric chemokine. For example, although a nonaggregating variant of RANTES/CCL5 was able to induce chemotaxis by means of Gi coupling, it was unable to activate T cells, monocytes, and neutrophils through protein tyrosine kinase (TK) pathways; this contrasts with wildtype (WT) CCL5, which forms large oligomers nucleated by a CC-like dimer, and induces TK proinflammatory pathways (Czaplewski et al., 1999). Oligomerization can also inhibit signaling as demonstrated by an obligate SDF-1/CXCL12 dimer, which was engineered by introduction of an intermolecular disulfide. It was shown to flux calcium but, unlike WT CXCL12, could not induce all of the pathways required for chemotaxis; instead, it blocked migration of cells to the WT chemokine (Veldkamp et al., 2008). Recently, several chemokines have been shown to heterodimerize with other chemokines, and in some cases, such as MCP-1/CCL2 and MCP-2/CCL8, their presence could be correlated with altered or amplified functional responses relative to signaling by only one of the chemokines (Crown et al., 2006; Dudek et al., 2003; Nesmelova et al., 2008). Although the relevance of oligomerization is now well established as the preceding examples suggest, there are 50 human chemokines, and little is known about the vast majority. Clearly, determining whether chemokines oligomerize and the structural details of the oligomers are crucial to understanding the mechanisms of chemokine-mediated processes. Furthermore, nonoligomerizing forms can have therapeutic value as demonstrated for a

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monomeric form of MCP-1/CCL2 that had antiinflammatory properties in animal models of experimental autoimmune encephalomyelitis and arthritis (Handel et al., 2008; Shahrara et al., 2008). This chapter will, therefore, discuss several approaches for studying chemokine oligomerization. The first step in this process is to decide what questions need to be addressed. Is residue-specific information needed to map subunit interfaces, solve oligomeric structures, or drive mutagenesis studies to create nonoligomerizing variants? Or, will simple confirmation and quantitation of oligomerization suffice? Often it is necessary to combine a variety of approaches to gain the most comprehensive understanding (Table 2.1). Furthermore, with all of these approaches, it is usually necessary to follow-up with mutagenesis or in vivo experiments to establish functional relevance (Campanella et al., 2006; Proudfoot et al., 2003).

2. Methods to Detect and Quantify Oligomerization Many variables, including protein concentration, salt concentration, and pH, affect the oligomeric equilibrium of chemokines (Veldkamp et al., 2005). Assessing the oligomeric state of a chemokine under varying conditions provides valuable data on which conditions facilitate or inhibit oligomerization. This knowledge is especially useful before structure determination by crystallography or NMR for understanding how to handle and store chemokines to preserve maximal functionality. The methods are also useful in the design and characterization of chemokine variants with reduced or enhanced propensities for oligomerization. The following section provides details and specific examples of methods that focus on determining the oligomerization state of chemokines and soluble proteins in general.

2.1. Analytical ultracentrifugation (AUC); sedimentation equilibrium In AUC experiments, one subjects a protein sample to a high centrifugal force in an analytical ultracentrifuge. One can then optically measure the distribution of the protein along the radius of the sample cell as a function of time or at equilibrium to determine shape and size distributions of proteins, as well as the equilibrium constant of oligomerization (Balbo et al., 2007). As such, this method has many applications in terms of analyzing chemokine oligomerization and the effects of solution composition and additives like GAGs. Although several types of experiments can be done by AUC, the method most commonly applied to chemokines is sedimentation

Table 2.1 Summary of various methods to study chemokine oligomerization. Advantages, disadvantages, and representative publications are listed Method

Advantages

Global detection of oligomerization Determination of Analytical equilibrium ultracentrifugation constant (AUC) PFG NMR (DOSY)

Rapid, easy analysis

Dynamic light scattering (DLS)

Fast and simple

Easy determination of dissociation constants FT-ICR mass spec. Can detect homo and heterodimerization Residue-specific information HSQC analysis Identification of residues at dimer interface NOE analysis Possible identification of residues at dimer interface Filtered NOEs Identification of residues at dimer interface Fluorescence polarization

Disadvantages

Examples

Expensive equipment, long length of experiments, analysis can be tricky

CXCL8 (Lowman et al., 1997), CCL4 ( Jin et al., 2007), CCL5 (Czaplewski et al., 1999) CXCL12 (Veldkamp et al., 2005), CCL27 CXCL12 (Holmes et al., 2001)

Very expensive equipment, limited to concentrations above
50 mM Large oligomers dominate signal, difficult to determine equilibrium constant Observe only small changes in chemokines

CXCL12 (Veldkamp et al., 2005)

Gas phase detection, no Kd determination

CCL2 and CCL8 (Crown et al., 2006)

Chemical shifts can also reflect change in conformation

CCL4 (Laurence et al., 2000), CCL2/CCL8 (Crown et al., 2006) CCL8 (Clore et al., 1990)

Difficult to distinguish intra- from inter-molecular NOEs, difficult data collection and analysis Low sensitivity, difficult data collection and analysis

CCL2 (Handel and Domaille, 1996), CXCL12 (Veldkamp et al., 2005)

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equilibrium, which is done at moderate centrifugal forces compared with the nonequilibrium method called sedimentation velocity. In the sedimentation equilibrium experiment, sedimentation of the protein by centrifugal force is counterbalanced by back diffusion of the protein along its concentration gradient. After equilibrium is achieved between these two forces, one measures the radial concentration gradient of the protein along the cell, which provides information on molecular mass, and for interacting systems, the dissociation constants. Lowman et al. (1997) determined the dimerization constants of several mutants of IL-8/CXCL8 designed to destabilize dimerization. They were able to identify mutants with significantly lower affinities for dimerization, yet showed that the mutants maintained WT potencies in several functional assays, suggesting that the monomeric form is the relevant form to induce migration. They also showed that dimerization was highly sensitive to solution conditions of ionic strength, pH, and temperature, and that high affinity could be achieved by adjustment of these parameters, suggesting that dimerization may have functional consequences despite the fact that only monomers are effective in activating the receptor (Lowman et al., 1997). Similarly, Paavola et al. (1998) identified a nondimerizing mutant of MCP-1/CCL2 that retained WT binding affinity and ability to promote cell migration. Jin et al. (2007) used AUC in conjunction with NMR to confirm both the molecular mass and correct folding of an engineered dimeric form of MIP-1b/CCL4. In contrast to studies with the disulfide-stabilized dimeric SDF-1/CXCL12 (Veldkamp et al., 2008), this study demonstrated that the disulfide-linked CCL4 dimer was not able to bind to its receptor CCR5, again confirming that CCL4 binds and activates CCR5 as a monomer ( Jin et al., 2007). Finally, Czaplewski et al. (1999) used AUC in parallel with mutagenesis and identified residues D26 and E66 in MIP-1a/CCL3 as critical for the formation of high molecular weight aggregates. They went on to demonstrate that homologous residues in CCL4 and CCL5 also inhibited aggregation, yet all three mutant chemokines maintained the ability to activate CCR1 and CCR5 in vitro (Czaplewski et al., 1999). The major advantage of sedimentation equilibrium is that it allows for the calculation of the molecular weight and dimerization constant(s) for a given system. Protein and protein complexes with masses in the range from <103 to >106 Daltons, and associating systems characterized by KD values between 104 and 108 M1 can be studied by AUC (Balbo et al., 2007). However, one caveat to this method is that it is time-consuming, taking 1 to 2 days per run. Furthermore, the data can be difficult to fit, particularly when there are issues with nonequilibrium aggregation, which plagues some chemokines (e.g., CCL5). One must also consider buffer conditions, because most chemokines are highly basic proteins. For example, it is important to avoid extremely low ionic strengths, because electrostatic interactions can become a significant force in addition to diffusion and

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sedimentation, complicating the analysis. Typically addition of 100 mM salt is sufficient to avoid nonideal behavior, but salt also commonly influences the oligomerization behavior. Primarily because of limitations on the sensitivity of the optical detection, protein concentrations are limited to the micromolar to submillimolar range (O.D. <1 for the highest sample concentration, which for chemokines is approximately 0.1 to 0.2 mM when measuring at 280 nm). Nevertheless, for chemokines, these concentration ranges are generally adequate to define the oligomerization state and/or KD for self-association. Figure 2.2 shows an example of the use of AUC A 0.02 0.00 −0.02 0.2

Residuals

B

−0.2

C

0.2

0.0

0.0 −0.2 D

Absorbance

1.2

0.8

0.4

0.0 6.6

6.7

6.8 6.9 7.0 Radius (cm)

7.1

7.2

Figure 2.2 Sedimentation equilibrium of SDF-1a. SDF-1a (200 mM) was centrifuged at 25,000 rpm, 4  C, until equilibrium was attained. Data were fit to various association state models. Residuals are plotted as the deviation from the best-fit line versus radial position for (A) an interacting monomer^dimer, (B) a monomer only, and (C) a dimer only. (D) Representative data are shown with the best-fit line for the interacting monomer^dimer model. Reprinted from ‘‘Protein Expression and Purification, 2001 Apr, 21(3):367^77, Solution studies of recombinant human stromal-cell^derived factor-1, Holmes, C. D.; Consler, T. G.; Dallas, W. S.; Rocque, W. J.; Willard, D. H.’’ with permission from Elsevier.

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to demonstrate that SDF-1/CXCL12 self-associates and that under the solution conditions used is in equilibrium between a monomer and dimer. In general, sedimentation equilibrium is accepted as the method of choice for studying oligomerization because of the highly quantitative information it provides. Many laboratories combine it with sedimentation velocity as a more rapid (hours) initial experiment to obtain information on the number of species in a sample (e.g., monomer, dimer, tetramer), but sedimentation velocity is less accurate in deriving equilibrium constants. For an excellent detailed description of the method, see Balbo et al. (2007).

2.2. Pulsed-field gradient diffusion by NMR Pulsed-field gradient (PFG) diffusion, effectively measures the average distance traveled by solute molecules in an NMR sample during a fixed period of time. The method relies on the use of pulse-field gradients to spatially encode the NMR signals such that their effective position in the NMR tube can be monitored (Altieri, 1995). Measuring the signal attenuation in successive proton spectra, as a function of increasing gradient field strength, enables calculation of the self-diffusion coefficient, Ds, which is directly related to the hydrodynamic radius of the protein in solution by means of the Stokes Einstein relationship (Altieri, 1995; Veldkamp et al., 2005, 2008). A relatively slowly decaying signal corresponds to a smaller value of Ds, which equates to a larger apparent molecular weight and vice versa. By accurately calibrating gradient strengths with solvents of known Ds, one can obtain an accurate measurement of the Ds of the protein of interest. Alternately, one can use known proteins of known molecular weight and oligomerization pattern as standards for Ds values. There are several interesting applications of PFG diffusion to the study of chemokine oligomerization. The Ds values may be compared at different protein concentrations to chart a chemokine oligomerization profile (Fig. 2.3) and can also be used to determine which solvent conditions affect the oligomerization state. Veldkamp et al. (2005) used this approach to study solvent conditions that promote dimerization SDF-1/CXCL12. By determining Ds as a function of concentration, it was possible to determine a value of KD, which was in good agreement with AUC measurements. Once this ‘‘oligomerization profile’’ has been determined for the WT chemokine, it can be compared with variants of the chemokine of interest to identify residues necessary for oligomerization and to generate mutants that remain monomeric at high concentrations. For example, Paavola et al. (1998) used such a method to determine mutations in the chemokine MCP-1/CCL2 that inhibited dimerization. Figure 2.3 shows a theoretical PFG diffusion profile for a 10-kDa chemokine forming a stable dimer. As the concentration increases, one would expect to see the Ds values rapidly decrease to a

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Ds (⫻10−06 cm2s−1)

1.5 1.4 1000 mM 1.3 100 mM

1.2 10 mM 1.1 1

0

0.2

0.4

0.6 0.8 1.0 1.2 1.4 [Chemokine] (mM)

1.6

1.8

2.0

Figure 2.3 Theoretical PFG diffusion curve showing values for Ds and concentration that would be expected for a typical chemokine of 10 kDa forming a stable dimer.These values are based on previous studies performed on chemokines and standard proteins of known molecular weight and oligomerization (Altieri,1995;Veldkamp et al., 2005).

value corresponding to approximately twice the molecular weight of the monomer and then stabilize as the concentration continues to increase. The ability to evaluate oligomerization at high concentration (millimolar range) is an important advantage of NMR, because these concentrations are not amenable to AUC. In addition, gradient diffusion experiments are rapid and easy to analyze. However, one must have access to the relevant NMR instrumentation. We favor the use of the PFG diffusion technique to screen the behavior of chemokines under different concentrations and buffer conditions. These data can then be used to identify a more limited set of conditions to characterize with the more time-consuming AUC method should this be deemed necessary. Relative to other NMR methods, PFG diffusion is fairly straight-forward to implement, and automated procedures, such as diffusion ordered spectroscopy (DOSY), are available for data processing and analysis in NMR vendor software packages such as Bruker Biospin TopSpin.

2.3. Dynamic light scattering Dynamic light scattering (DLS) is a well-established technique for studying protein oligomerization and is often used by crystallographers to determine a protein’s propensity to aggregate in solution. DLS measures intensity fluctuations in scattered, monochromatic light through a sample in solution and ultimately allows the calculation of the diffusion coefficient and from there the hydrodynamic radius. A good example of DLS applied to chemokines is a study done by Holmes et al. (2001) to determine the

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monomer–dimer equilibrium of recombinant SDF-1/CXCL12. They used DLS to calculate the apparent mass of CXCL12 at six different concentrations and determined the monomer–dimer equilibrium constant to be 114 mM. This value was verified by sedimentation equilibrium, which showed a similar equilibrium constant of 180 mM. DLS has the advantage of speed and requires less technical expertise than many of the other methods described. Typically DLS is less accurate for observing the presence of small oligomers and determining equilibrium constants than sedimentation equilibrium. However, it is especially good for detecting broad distributions of species with very different molecular masses such as native proteins and protein aggregates, which could be useful for chemokines that exhibit such behavior (e.g., RANTES/CCL5).

2.4. Fluorescence polarization Fluorescence polarization (FP) has been used to determine the dependence of the dissociation constant for dimerization of SDF-1a/CXCL12 on solution conditions, including pH, salt, and the presence of heparin (Veldkamp et al., 2005). FP is measured after excitation of a fluorophorecontaining molecule with linearly polarized light that causes ‘‘photoselection’’ of fluorophores with their absorption dipoles aligned parallel to the vector of the polarized light. The FP value depends on the reorientation rate of the fluorophore, which in turn depends on the size of the molecule it is attached to. Small molecules rotate rapidly, causing rapid randomization of the originally polarized light and low FP. By contrast, large molecules rotate slowly and have higher FP values compared with lower molecular weight, faster tumbling counterparts. The dependence on size makes FP a viable method for measuring oligomerization. In the study by Veldkamp et al. (2005) the fluorescence was derived from tryptophan residues that are highly conserved in chemokines. On oligomerization of the chemokine, the FP value increases, and when done as a function of chemokine concentration, the curves can be fit to determine dissociation constants for oligomerization. Given the size of chemokines, only small changes in FP are observed for monomer–dimer equilibria, but they are still sufficient for reasonably accurate affinity determinations. The quantum yield of Trp is also sufficient for the concentration ranges needed for chemokine oligomerization. In principle, sensitivity could be increased by covalent attachment of fluorophores with higher quantum yields. The measurements are rapid and easy both in terms of data collection and analysis. Access to the appropriate instrumentation is generally not difficult. Thus FP is certainly a method to consider for studying oligomerization, especially when comparing effects of mutations and/or solution conditions.

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2.5. FT-ICR mass spectrometry Fourier-transform ion cyclotron resonance (FT-ICR) mass spectrometry is a useful method for studying noncovalent chemokine: GAG interactions as well as oligomerization of chemokines (Crown et al., 2006; Yu et al., 2005, 2006). This method is facilitated by electrospray ionization (ESI), which enables transfer of intact noncovalent complexes from solution to gas phase (Crown et al., 2006). FT-ICR has very high mass accuracy and can be used to determine the molecular weight of complexes, stoichiometry, etc., and previous experiments have yielded results that at least qualitatively correspond to AUC and NMR (Yu et al., 2005, 2006). It is particularly useful for examining heterodimerization, which would not be possible by any of the previously described methods. With FT-ICR, Crown et al. analyzed the ability of the MCP-1 ligands (MCP-1/CCL2, MCP-2/CCL8, MCP-3/ CCL7, MCP-4/CCL13) and eotaxin/CCL11 to homo- and heterodimerize (Crown et al., 2006). CCL2 and CCL8 showed a high propensity to heterodimerize, whereas heterodimers between CCL2 and CCL11, CCL2 and CCL13, and CCL8 and CCL13 were weaker, and no ligand heterodimerized with CCL7. In addition, they demonstrated that in the case of CCL2/CCL8, heterodimerization formed at the expense of homodimerization, suggesting functional relevance (Crown et al., 2006). FT-ICR measurements can be done quite rapidly and, therefore, are useful for screens of this nature. However, because the measurements are done in the gas phase, it remains to be seen how quantitative the method is. Nevertheless, use of FT-ICR in combination with NMR for quantitation (see later), can be quite powerful. Chapter 4 in this volume describes the use of FT-ICR for analyzing chemokine: GAG interactions.

2.6. Other methods 2.6.1. Size exclusion chromatography Size exclusion chromatography (SEC) is a classic method for determining molecular size. It works by determining the volume of solvent required to elute a protein sample over a column containing porous beads. The beads have a defined distribution of pore sizes that control the size or ‘‘fractionation range’’ of proteins that can be separated from each other. The elution volume is relative and must be calibrated with known protein standards, and it is important to realize that molecules are separated according to both mass and shape. However, we mention it here as being one of the least desirable of the methods primarily because it is a nonequilibrium measurement, and dissociation of interacting species will occur during the experiment. Although SEC has been used to analyze chemokine oligomerization (Holmes et al., 2001; Hoogewerf et al., 1997), it is only applicable for very

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tightly bound complexes, which is typically not characteristic of chemokine oligomerization, with some exceptions like RANTES/CCL5. 2.6.2. 15N Heteronuclear relaxation by NMR 15N T1, T2, and heteronuclear 1H–15N NOE relaxation measurements from NMR experiments give information on protein dynamics and are generally interpreted with respect to several time scales: picosecond to nanosecond internal motions within the protein, overall global rotational tumbling of the protein (nanosecond), and microsecond to millisecond time scale motions indicative of chemical exchange such as monomer–dimer equilibrium (Ishima and Torchia, 2000). Baryshnikova and Sykes (2006) evaluated the concentration dependence of NMR relaxation parameters of CXCL12 at different concentrations to determine the binding constant for dimerization. They also demonstrated that regions from CXCL12 involved in the dimer interface in structures previously determined by crystallography are involved in slow time scale motions. Thus information on oligomerization is obtainable with these methods; however, they are not as accurate as methods like AUC. Therefore, this approach is not recommended except for the NMR aficionado, because the experimental data collection and analysis are technically quite involved, extremely time-consuming (many days), and the experiments require the production of multimilligram quantities of 15N-labeled protein.

3. Methods for Collecting Residue-Specific Information on Chemokine Oligomers NMR is a powerful technique for studying protein structures in solution. Several different methods have been applied to chemokines to specifically detect sites of interactions, such as the dimer interface and interactions with GAGs and with receptor fragments (Crown et al., 2006; McCornack et al., 2003; Nesmelova et al., 2008; Veldkamp et al., 2008). These methods all require fully assigned proton-nitrogen and/or proton-carbon correlation spectra for the protein under study and thus require considerable data collection and analysis times, as well as isotopically labeled protein.

3.1. NMR: Heteronuclear single quantum correlation (HSQC) spectroscopy Changes in chemical shift observed in heteronuclear single quantum coherence (HSQC) spectra could be used to determine which residues are involved in forming a dimer interface and potentially to calculate a binding constant. NMR signals corresponding to each backbone 15N-1H residue

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may shift frequency on the basis of changes in the local magnetic environment caused by dimerization. These changes or perturbations can be monitored as a function of concentration to study oligomerization. For example, chemical shift perturbations were used in conjunction with AUC to determine that the residues Phe13 and Pro8 are essential for the dimerization of CCL4/MIP-1b (Laurence et al., 2000). Crown et al. (2006) also used chemical shift perturbations to study the heterodimerization of MCP-1/ CCL2 and MCP-2/CCL8. By titrating increasing amounts of unlabeled CCL8 into a sample of 15N-labeled CCL2, they were able to monitor changes in chemical shifts of CCL2 1H-15N resonances, indicating that the chemokines were heterodimerizing and forming CC-like dimer structures (Crown et al., 2006). The near complete loss of signals for the CCL2 dimer on the addition of stoichiometric amounts of CCL8 indicated that heterodimer formation was favored over homodimer formation, similar to results from FT-ICR. The use of the HSQC was particularly important for this study, because most other methods described in the previous section, with the exception of FT-ICR, would not be able to distinguish homodimer formation from heterodimer formation. HSQC analysis is relatively rapid (minute to hours), although it requires the production and prior assignment of isotopically labeled protein. It is important to note, however, that because chemical shift perturbations can be caused by intermolecular interactions or by conformational rearrangements, these analyses do not unambiguously define the oligomeric interface but rather point to a general region on the basis of a consensus of chemical shift changes mapped onto a structure.

3.2. NMR: Detection of nuclei in close proximity by means of the nuclear Overhauser effect (NOE) The nuclear Overhauser effect (NOE) is used to determine nuclei within close proximity to each other and can include through-space signals both within a monomer (intramolecular) and between two subunits of a dimer (intermolecular). The NOE signal is inversely proportional to the sixth power of the distance between two nuclei and, therefore, is only observable ˚ apart. This distance constraint allows these experibetween nuclei <
6 A ments to define residues at the interface of dimers. For example, Clore et al. (1990) used NOEs to solve the dimer structure of the chemokine IL-8/ CXCL8 and determined that the interface is primarily stabilized by an antiparallel b-sheet formed between the first b-strands of each subunit (a classic ‘‘CXC’’ dimer, Fig. 2.1C), as well as several important sidechain interactions. The notable caveat to this method is the difficulty in distinguishing between intramolecular and intermolecular NOE signals. In many instances there is sufficient signal overlap to make this determination

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virtually impossible. This caveat has led to the development of filtered experiments that, in principle, select only for intermolecular NOEs. 3.2.1. Filtered NOE experiments The problem of distinguishing between intramolecular and intermolecular signals may be addressed with a combination of isotope-filtered and isotopeedited NOE experiments on complexes formed from mixtures of asymmetrically labeled chemokine monomers to break the magnetic symmetry and directly determine intersubunit NOEs (Breeze, 2000; Handel and Domaille, 1996; Jin et al., 2007; Veldkamp et al., 2008). For example, Handel and Domaille (1996) generated a sample of a 50:50 mixture of 15N-labeled and 13C-labeled MCP-1/CCL2 and used a filtered NOE experiment that selectively detects intersubunit signals between 15N-1H and 13C-1H pairs across the dimer interface. One advantage of this (HC)NH-NOE experiment is that magnetization is filtered through two different magnetically active nuclei (as opposed to other filters that record 13C-1H to 12C-1H), allowing additional suppression of intramonomer NOE signal (Handel and Domaille, 1996). Figure 2.4 shows an example of such an experiment compared with the assigned 1H-15N HSQC (Handel and Domaille, 1996). With this experiment, it was possible to assign 65 intersubunit NOEs, which characterized the dimer interface as an antiparallel b-sheet between the N-termini of the subunits. These results were used as additional restraints to solve the CCL2 dimer structure, which was determined to be what is now known as a CC dimer. More recently, Veldkamp et al. used a similar filtered NOE experiment on a mixed-label sample of SDF-1a/CXCL12 to study the interaction between an engineered covalent dimer and the sulfated N-terminal fragment of the CXCR4 receptor (Veldkamp et al., 2008). Their results suggest that receptor sulfation may enhance ligand affinity and stabilize the bound chemokine dimer. Filtered NOE experiments have been used successfully in evaluating other chemokine dimer interfaces; however, not all chemokines may be amenable to this approach. The NOE signal is relatively weak to begin with, and this is exacerbated by the fact that only 50% of the sample will correspond to a mixed-label dimer between 15N and 13C-labeled monomers (25% will be 15N-only and 25% will be 13C only). In addition, the NOE depends on dipolar coupling being the dominant relaxation mechanism. Any other competing relaxation mechanism, such as unfavorable dynamics associated with oligomerization, may make it impossible to obtain sufficient signal for analysis. These experiments and their analysis also require a great deal of experimental sophistication, making them less routinely used. The NMR methods just listed are useful tools for studying chemokine dimerization under physiologic conditions. However, larger order oligomers such as tetramers have not been successfully studied by NMR because of a number of physical and spectroscopic issues that are beyond the scope of this chapter. Fortunately, the structural details of chemokine tetramers have

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A

15N- 1H

HSQC

114

15N

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116 118 120 122 124

10

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7

(HC) NH-NOE 14N

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15F 66H 65D 14N

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57Q

23Q 70Q

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9V 68D

11C 40A

30R

122

39E 124

76T 16R 4A

52C 17N 10

19K

7A

38K 47V

9 1H

8 (ppm)

7

Figure 2.4 (A) 2-D 15N HSQC of uniformly labeled CCL2/MCP-1. (B) 2-D 1H-15N plane of the (HC)NH NOE experiment on a 50:50 mixture of (15N þ 13C)^labeled CCL2. The observed cross-peaks in the 2-D 15N HSQC unambiguously establish the backbone and side chain amide residues involved in the dimerization interface and were used by Handel et al. to solve the dimer structure of CCL2 (Handel and Domaille,1996).

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been solved by the complementary technique of X-ray crystallography as demonstrated for CCL2, IP-10/CCL10, and PF-4/CCL4 (Baldwin et al., 1991; Barinka et al., 2008; Blain et al., 2007; Lubkowski et al., 1997; Ryu et al., 2007; Shaw et al., 2004; Swaminathan et al., 2003).

4. Conclusions Chemokines are involved with many different interactions–with their receptors, with GAGs, and with other chemokines through both homoand hetero-oligomerization. In addition, the chemokine network is complex, with many chemokines interacting with multiple receptors and vice versa, yet many chemokines maintain tissue specificity. Understanding the structural characteristics and mechanisms of chemokine oligomerization is a crucial step in determining details of these different interactions. As this review shows, a wide variety of methods can be used to study chemokine oligomerization, both residue-specific and more generalized. It is often necessary to use these methods in combination with each other or in conjunction with other techniques such as mutagenesis to obtain the most complete picture possible. Although we know that many chemokines dimerize, the importance of higher order oligomerization and heterodimerization is less defined. Correlating these oligomeric states with function remains an important endeavor for fully understanding the role that structural plasticity plays in chemokine functional diversity and regulation.

ACKNOWLEDGMENTS This work was funded by the NIH through the Molecular Biophysics Training Grant (GM08326) awarded to A. J., NRSA postdoctoral fellowship F32GM083463 awarded to D. J. H., and by the Lymphoma Research Foundation, the Department of Defense (USAMRAA W81XWH0710446) and NIH (RO1-AI37113) awards to T. M. H.

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oligomerization for chemokine activity in vivo Inhibition of monocyte chemoattractant protein-1 ameliorates rat adjuvant-induced arthritis. J. Leukoc. Biol. 84, 1101–1108. Hartley, O., Dorgham, K., Perez-Bercoff, D., Cerini, F., Heimann, A., Gaertner, H., Offord, R. E., Pancino, G., Debre, P., and Gorochov, G. (2003). Human immunodeficiency virus type 1 entry inhibitors selected on living cells from a library of phage chemokines. J. Virol. 77, 6637–6644. Holmes, W. D., Consler, T. G., Dallas, W. S., Rocque, W. J., and Willard, D. H. (2001). Solution studies of recombinant human stromal-cell-derived factor-1. Protein Expr. Purif. 21, 367–377. Homey, B., Alenius, H., Mu¨ller, A., Soto, H., Bowman, E. P., Yuan, W., McEvoy, L., Lauerma, A. I., Assmann, T., Bu¨nemann, E., Lehto, M., Wolff, H., et al. (2002). CCL27CCR10 interactions regulate T cell-mediated skin inflammation. Nat. Med. 8, 157–165. Hoogewerf, A. J., Kuschert, G. S., Proudfoot, A. E., Borlat, F., Clark-Lewis, I., Power, C. A., and Wells, T. N. (1997). Glycosaminoglycans mediate cell surface oligomerization of chemokines. Biochemistry 36, 13570–13578. Imberty, A., Lortat-Jacob, H., and Perez, S. (2007). Structural view of glycosaminoglycanprotein interactions. Carbohydr. Res. 342, 430–439. Ishima, R., and Torchia, D. A. (2000). Protein dynamics from NMR. Nat. Struct. Biol. 7, 740–743. Jin, H., Hayes, G. L., Darbha, N. S., Meyer, E., and LiWang, P. J. (2005). Investigation of CC and CXC chemokine quaternary state mutants. Biochem. Biophys. Res. Commun. 338, 987–999. Jin, H., Shen, X., Baggett, B. R., Kong, X., and LiWang, P. J. (2007). The human CC chemokine MIP-1beta dimer is not competent to bind to the CCR5 receptor. J. Biol. Chem. 282, 27976–27983. Johnson, Z., Kosco-Vilbois, M. H., Herren, S., Cirillo, R., Muzio, V., Zaratin, P., Carbonatto, M., Mack, M., Smailbegovic, A., Rose, M., Lever, R., Page, C., et al. (2004). Interference with heparin binding and oligomerization creates a novel anti-inflammatory strategy targeting the chemokine system. J. Immunol. 173, 5776–5785. Johnson, Z., Proudfoot, A. E., and Handel, T. M. (2005). Interaction of chemokines and glycosaminoglycans: A new twist in the regulation of chemokine function with opportunities for therapeutic intervention. Cytokine Growth Factor Rev. 16, 625–636. Kunkel, E. J., and Butcher, E. C. (2002). Chemokines and the tissue-specific migration of lymphocytes. Immunity 16, 1–4. Kuschert, G. S., Coulin, F., Power, C. A., Proudfoot, A. E., Hubbard, R. E., Hoogewerf, A. J., and Wells, T. N. (1999). Glycosaminoglycans interact selectively with chemokines and modulate receptor binding and cellular responses. Biochemistry 38, 12959–12968. Lau, E. K., Paavola, C. D., Johnson, Z., Gaudry, J. P., Geretti, E., Borlat, F., Kungl, A. J., Proudfoot, A. E., and Handel, T. M. (2004). Identification of the glycosaminoglycan binding site of the CC chemokine, MCP-1: Implications for structure and function in vivo. J. Biol. Chem. 279, 22294–22305. Laurence, J. S., Blanpain, C., Burgner, J. W., Parmentier, M., and LiWang, P. J. (2000). CC chemokine MIP-1 beta can function as a monomer and depends on Phe13 for receptor binding. Biochemistry 39, 3401–3409. Loetscher, P., and Clark-Lewis, I. (2001). Agonistic and antagonistic activities of chemokines. J. Leukoc. Biol. 69, 881–884. Lowman, H. B., Fairbrother, W. J., Slagle, P. H., Kabakoff, R., Liu, J., Shire, S., and Hebert, C. A. (1997). Monomeric variants of IL-8: Effects of side chain substitutions and solution conditions upon dimer formation. Protein Sci. 6, 598–608. Lubkowski, J., Bujacz, G., Boque, L., Domaille, P. J., Handel, T. M., and Wlodawer, A. (1997). The structure of MCP-1 in two crystal forms provides a rare example of variable quaternary interactions. Nat. Struct. Biol. 4, 64–69. McCornack, M. A., Cassidy, C. K., and LiWang, P. J. (2003). The binding surface and affinity of monomeric and dimeric chemokine macrophage inflammatory protein 1 beta for various glycosaminoglycan disaccharides. J. Biol. Chem. 278, 1946–1956.

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C H A P T E R

T H R E E

Lymphotactin Structural Dynamics Brian F. Volkman, Tina Y. Liu, and Francis C. Peterson Contents 1. 2. 3. 4. 5. 6. 7.

Introduction Production of Biologically Active Lymphotactin Ltn Folds into Two Unrelated Native State Structures Kinetics of Interconversion in the Ltn Conformational Equilibrium Engineering Conformationally Restricted Lymphotactin Variants Functional Analysis of Distinct Ltn Native State Conformations GAG Binding Residues are Linked to the Ltn Conformational Equilibrium 8. Conclusions References

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Abstract Lymphotactin/XCL1, the defining member of the C class of chemokines, undergoes a conformational change that involves the complete restructuring of all stabilizing interactions. Other chemokines are restricted to a single conformation by a pair of conserved disulfide crosslinks, one of which is absent in lymphotactin. This structural interconversion is entirely reversible, and the two-state equilibrium is sensitive to changes in temperature and ionic strength. One species adopts the conserved chemokine fold as a monomer and functions as an agonist for XCR1, the specific G-protein–coupled receptor for lymphotactin. Rearrangement to the other conformation produces a novel four-stranded sheet that dimerizes to form a beta sandwich with high affinity for cell-surface glycosaminoglycans. We developed methods for resolving the two species and investigated the dynamics of human lymphotactin structural interconversion with NMR spectroscopy, heparin affinity chromatography, and time-resolved fluorescence on the wild-type protein and a panel of amino acid–substituted lymphotactin variants. Our results show that the lymphotactin structural rearrangement occurs at a rate of
1/s and that mutation of residues required for glycosaminoglycan binding shifts the conformational equilibrium toward the chemokine-like fold. We speculate that charge repulsion between arginines 23 and 43 destabilizes the chemokine fold and promotes conversion to the novel Department of Biochemistry, Medical College of Wisconsin, Milwaukee, Wisconsin, USA Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05403-2

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2009 Elsevier Inc. All rights reserved.

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lymphotactin dimer, whereas binding of chloride or another anion stabilizes the chemokine fold by neutralizing the repulsive effect.

1. Introduction Lymphotactin (Ltn) is a chemokine that recruits T and NK cells and is produced mainly by activated CD8þ T cells and activated NK cells. Like other chemokines, Ltn induces intracellular calcium mobilization and chemotaxis by binding a specific G-protein–coupled receptor (GPCR) (Yoshida et al., 1998). When it was originally cloned from a mouse progenitor T cell cDNA library in 1994 (Kelner et al., 1994), comparisons with other chemokines highlighted two novel structural features in the Ltn sequence. At 93 residues in length, the mature, secreted form of Ltn contains a C-terminal extension of
25 amino acids relative to the CXC and CC chemokines. Moreover, it lacks the first and third of four conserved cysteine residues found in all other chemokines, and thus possesses only a single disulfide bond. Although the CXC and CC chemokine genes cluster on human chromosomes 4 and 17, respectively (Modi and Chen, 1998; Naruse et al., 1996), human lymphotactin is located on chromosome 1 (Kelner et al., 1994). On the basis of these distinctions, lymphotactin was taken to define a novel type of chemokine, the ‘‘C’’ family (Kelner et al., 1994), and given the systematic designation XCL1. Ltn binds the GPCR XCR1 with low nanomolar affinity to induce CD8þ T cell and NK cell chemotaxis and chemoattracts CD4þ T cells with lower efficiency (Kelner et al., 1994). This may be due in part to the ability of Ltn to costimulate the apoptosis of CD4þ T cells but not CD8þ T cells (Cerdan et al., 2001). Ltn is produced mainly through T cell receptor activation in CD4þ and CD8þ T cells (Kelner et al., 1994; Tikhonov et al., 2001) but also by NK cells (Kelner et al., 1994) and gd T cells (Boismenu et al., 1996). Together, the data suggest that Ltn produced by T cells can regulate or modulate T cell–mediated immune responses. Lymphotactin expression or activity is associated with a number of T cell–mediated disease states. Data from rheumatoid arthritis patients (Blaschke et al., 2003) and animal models for acute allograft rejection (Wang et al., 1998), Crohn’s disease (Scheerens et al., 2001), and glomerulonephritis (Natori et al., 1998) are consistent with an immunomodulatory role for Ltn. Inhibitors or mimics of this chemokine may, therefore, have therapeutic value in the context of autoimmune and inflammatory diseases. In fact, Ltn has already been used in the development of novel cancer immunotherapies. A number of animal studies have shown that Ltn can recruit T cells to the site of a tumor (Dilloo et al., 1996; Huang et al., 2005), and combined expression of Ltn and interleukin-2 in a neuroblastoma

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tumor vaccine induced measurable antitumor immune responses, including complete remission in two patients (Rousseau et al., 2003). Chemokines share a highly conserved tertiary structure consisting of a flexible N-terminus, three-stranded b-sheet, and C-terminal a-helix, stabilized by two disulfide bonds (Clore and Greonenborn, 1995). Despite its small size (<10 kDa), the surface of each chemokine must support specific high-affinity binding to two different targets to function in vivo. In addition to specifically recognizing and activating a GPCR on the target cell, chemokines also bind glycosaminoglycans (GAGs) immobilized in the extracellular matrix (Proudfoot, 2003, 2006). Although the fundamental structural features of the family are now well established, novel aspects of chemokine structure and recognition continue to be revealed. The ability to form homodimers may also be essential for chemokine function (Campanella et al., 2006; Proudfoot et al., 2003), and chemokine heterodimers have recently been characterized (Crown et al., 2006; Nesmelova et al., 2008). No chemokine receptor structures have yet been solved, but sulfotyrosine modifications in the GPCR N-terminus enhance their recognition by chemokine ligands (Bannert et al., 2001; Farzan et al., 2002; Fong et al., 2002; Seibert et al., 2002, 2008), as illustrated recently for a complex between SDF-1/CXCL12 and its receptor CXCR4 (Veldkamp et al., 2006, 2008). Unlike all other chemokines, lymphotactin is constrained by only one disulfide bond, permitting it to access two completely different native state structures. A series of NMR studies showed that Ltn is conformationally heterogeneous in solution (Kuloglu et al., 2001), interconverting between the conserved chemokine fold (Kuloglu et al., 2002) and an unrelated dimeric structure (Tuinstra et al., 2008). This metamorphic folding behavior is unlike any previously described protein conformational rearrangement (Murzin, 2008). The chemokine-like Ltn conformation is a functional XCR1 agonist but has no high-affinity glycosaminoglycan (GAG)-binding site. In contrast, the alternative structure binds GAGs with high affinity but fails to activate XCR1. Because each structural species displays only one of the two functional properties essential for activity in vivo, the conformational equilibrium is likely to be essential for the biologic activity of lymphotactin. Despite its ability to orchestrate T cell–mediated immune responses, a specific biologic role for lymphotactin remains obscure. Some investigators have reported difficulty obtaining reproducible lymphotactin activity, and these challenges may derive from its unusual conformational plasticity. We investigated the dynamics of the native state equilibrium with a number of different methods as described in this chapter, with the ultimate goal of understanding how interconversion between two distinct lymphotactin structures contributes to its functional role in the immune system.

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2. Production of Biologically Active Lymphotactin Despite the challenge associated with large-scale production of disulfide-containing proteins, many biologically active chemokines have been prepared by chemical synthesis or bacterial expression (Clark-Lewis et al., 1997). A key consideration is to mimic the mature protein secreted from mammalian cells after processing of an amino terminal signal peptide, because like most chemokines, lymphotactin can be inactivated by any modifications to the amino terminal sequence (Tuinstra et al., 2007). Amino acid sequences for human lymphotactin and its orthologs are shown in Fig. 3.1A. We have used two different approaches for bacterial expression of Ltn for structural and functional studies. Our original expression system used Factor Xa cleavage of an N-terminal fusion protein to release Ltn with the native N-terminus (Kuloglu et al., 2001). This material was biologically active in assays that measure XCR1 activation by chemotaxis or intracellular calcium flux, but nonspecific proteolysis of the Ltn C-terminus led us to develop a more robust method based on cyanogen bromide (CNBr) cleavage of an N-terminal Hexa histidine tag. CNBr cleaves the peptide bond following methionine residues, but human Ltn contains two nonconserved Met residues (M63 and M73) that we changed to valine and alanine, respectively (Tuinstra et al., 2007). Ltn proteins produced by the two methods are indistinguishable in terms of structure and biologic activity (Tuinstra et al., 2007, 2008). Final purification of chemokines is typically achieved by reverse-phase HPLC after disulfide formation in an oxidation/folding step. Lymphotactin refolding can be achieved either in solution or with an on-column buffer exchange process to catalyze disulfide formation (Veldkamp et al., 2007). Oncolumn refolding is considerably faster and can improve the yield of protein with the correct disulfide pattern for some chemokines. However, because Ltn contains only a single disulfide bond, solid-phase refolding presents no particular advantage, and we typically perform disulfide oxidation in solution as previously described (Peterson et al., 2004). An unusual feature of the lymphotactin structure is glycosylation of the extended C-terminus in a portion of the protein purified from cultured mammalian cells (Dorner et al., 1997). Investigations into the functional significance of glycosylation used Ltn synthesized with carbohydrate modifications (Marcaurelle et al., 2001) or expressed in insect cells with a baculovirus system (Dong et al., 2005). Glycosylation had no detectable effect on XCR1 activation (Marcaurelle et al., 2001), and our own measurements confirm that the Ltn C-terminus (residues 73 to 93) is dispensable for both calcium-flux (Tuinstra et al., 2007) and T cell chemotaxis

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Figure 3.1 Lymphotactin adopts two unrelated folds in equilibrium. (A) Alignment of Ltn sequences from human, mouse, rat, cow, and chicken. (B) 1H-15N HSQC spectra acquired for wild-type human Ltn at 37  C and (C) 10  C in 20 mM sodium phosphate (pH 6.0) containing 200 mM sodium chloride. (D) Structure of wild-type Ltn at 10  C showing that under these solution conditions Ltn adopts the canonical chemokine fold, subsequently referred to as Ltn10. Unstructured residues (1 to 7, 76 to 93) were omitted for clarity. (E) 1H-15N HSQC spectrum of wild-type Ltn at 40  C acquired in 20 mM sodium phosphate (pH 6.0) in the absence of salt. (F) Structure of wild-type Ltn at 40  C (Ltn40) adopts a novel four-stranded antiparallel b-sheet that self-associates as a head-to-tail dimer. Disordered N- and C-termini (1 to 7, 56 to 93) were omitted for clarity.

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(S. Kutlesa, F. C. Peterson, and B. F. Volkman, unpublished results). A purpose for Ltn glycosylation thus remains unknown, and the functional role of the novel C-terminal sequence is a focus of our ongoing studies.

3. Ltn Folds into Two Unrelated Native State Structures NMR spectroscopy provided the first evidence of conformational duality in the lymphotactin structure. A 2-D 1H-15N HSQC spectrum acquired in typical conditions for protein NMR (20 mM sodium phosphate buffer, 200 mM NaCl, pH 6.0, 37  C) contains roughly twice the number of peaks expected for a stably folded 93-residue protein (Fig. 3.1B). Intense signals in a narrow 1H chemical shift range (
7 to 8 ppm) suggested that some of the protein was unfolded, whereas other more dispersed peaks corresponded to one or more folded states, but the spectrum was otherwise uninterpretable. We verified the identity, purity, and disulfide oxidation state by analytical HPLC and electrospray ionization mass spectrometry and reproduced the same 1H-15N HSQC spectrum in multiple preparations. Initially, we varied sample pH, temperature, and buffer composition with no obvious improvement in the HSQC spectrum; however, larger changes in temperature and solution ionic strength had a surprisingly dramatic effect. After systematic NMR screening of both parameters, we obtained a vastly simplified HSQC spectrum at 10  C in 200 mM NaCl (Fig. 3.1C) and solved the NMR structure under those conditions (Kuloglu et al., 2001). We refer to the conformation observed in low-temperature/high-salt conditions as Ltn10 (Fig. 3.1D), a monomeric state that closely resembles other chemokines with the addition of a highly flexible C-terminal extension consisting of residues 70 to 93. A very different HSQC pattern was observed at higher temperature in low ionic strength buffer (Fig. 3.1E) (Kuloglu et al., 2002). Analytical ultracentrifugation revealed that the Ltn structure at 40  C in the absence of salt (Ltn40) is dimeric with a Kd in the low micromolar range, but the structure of the Ltn40 dimer proved difficult to resolve. Our initial NMR studies of Ltn40 revealed a dramatic change from the typical pattern of secondary structure elements observed in the chemokine-like Ltn10 structure. Nuclear Overhauser effect (NOE) cross peaks identified longrange contacts between a new b-strand comprised of residues 11 to 14 (b0) and the b3 strand and showed that the a-helix had become disordered with the rest of the C-terminal domain (residues 53 to 93), but they failed to identify the Ltn40 dimer interface (Kuloglu et al., 2002). In an effort toward selective detection of intermolecular NOEs linking aliphatic protons from one subunit to amide protons of the other subunit, we acquired a F1

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13C-edited/F3 15N-edited

3D NOESY-HSQC spectrum on a equimolar mixture of [U-15N]-Ltn and [U-13C]-Ltn, but it was devoid of signals. Heteronuclear filtering can be technically difficult in protein NMR spectroscopy, particularly the 13C filtering of proton signals exhibiting a wide range of 1H-13C coupling constants. We successfully detected unambiguous intermolecular NOEs only after implementing an F1 13C/15Nfiltered/F3 13C-edited 3D NOESY-HSQC pulse sequence originally described by Palmer and coworkers (Stuart, 1999). Their pulse scheme compensates for the normal variations in 1H-13C scalar couplings in proteins and, most importantly, is both robust and simple to use, requiring no complex pulse calibrations or empirical optimizations. We routinely use this filtered NOESY experiment to solve structures for homodimer (Peterson et al., 2006) and heterodimer (Veldkamp et al., 2008) complexes. A total of 31 intermolecular NOEs were assigned in the filtered NOESY spectrum of Ltn40, each of which gave rise to a pair of symmetry-related intermolecular distance restraints. These restraints enabled us to solve the structure of the Ltn40 dimer (Tuinstra et al., 2008), which adopts a novel b-sandwich fold with four strands in each monomer subunit and no helices (Fig. 3.1F). Although the b1-, b2-, and b3-strands may seem unchanged, the rearrangement from Ltn10 to Ltn40 creates a completely different pattern of b-sheet hydrogen bonding and hydrophobic packing as described previously (Kuloglu et al., 2002; Tuinstra et al., 2008). Thus, Ltn folds into two distinct, unrelated structures, depending on temperature and solution conditions.

4. Kinetics of Interconversion in the Ltn Conformational Equilibrium For both native state Ltn structures to contribute to its chemokine activity in vivo, we postulated that individual molecules must be capable of converting from the Ltn10 state to the Ltn40 state and vice versa. A series of HSQC spectra collected on a single sample from 10 to 40  C confirmed that the Ltn10–Ltn40 transition is fully reversible, and spectra at intermediate temperatures contain distinct signals for each conformational state (Kuloglu et al., 2002; Tuinstra et al., 2007). Thus, changes in the Ltn conformational equilibrium induced by temperature, ionic strength, mutations, or binding interactions can be monitored by 1-D 1H and 2-D 1H-15N NMR spectroscopy. It is also of interest to measure the kinetics of conformational exchange, and the choice of technique depends on the time scale. The presence of distinct HSQC signals for the two species in equilibrium indicates that the frequency of Ltn10–Ltn40 interconversion is significantly lower than the observed chemical shift differences (
50 to 100 Hz).

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We initially tried to assess the rate of structural interconversion by monitoring amide H/D exchange rates. We reasoned that because the conformational rearrangement breaks every hydrogen bond and replaces it with a different one (Kuloglu et al., 2002), the lifetime of the Ltn10 and Ltn40 species would set an upper limit on the protection factor of all amide hydrogens. If the interconversion is very slow (minutes or longer), the observed H/D exchange rates would be dominated by this rate. We acquired 1-D 1H and 2-D 1H-15N HSQC spectra on a sample of lyophilized Ltn dissolved in 100% D2O. Regardless of pH, temperature, or salt concentration, we failed to detect any amide 1H signals in the first spectrum, which was acquired less than 5 min after dissolving the sample. Knowing that the interconversion rate was on the order of seconds (not milliseconds or minutes), we measured 2-D longitudinal 15N zz-exchange spectra (Farrow et al., 1994) to detect exchange peaks that appear at the intersection of 1H and 15N shifts for the same residue in the Ltn10 and Ltn40 conformations. We acquired a series of 2-D exchange spectra with different exchange mixing periods at 37  C in low salt buffer conditions where Ltn40 is more abundant than Ltn10 but both species are present at detectable levels (Fig. 3.2A) (Tuinstra et al., 2008). No exchange cross peaks were detected until the mixing time exceeded 50 msec with maximal intensities observed at
350 msec. Nonlinear fitting to equations accounting for a two-state exchange and 15N T1 relaxation (Tollinger et al., 2001) yielded rate constants for the forward (kforward = 0.4 s1) and reverse (kreverse = 1.2 s1) reactions consistent with an overall exchange rate, kex, of
1.5 s1. The single tryptophan in Ltn, W55, is another useful sensor of the Ltn10–Ltn40 conformational equilibrium (Kuloglu et al., 2002; Tuinstra et al., 2008). Transfer of the W55 side chain from a constrained environment in the Ltn10 hydrophobic core to a mobile, solvent-exposed state in Ltn40 shifts the tryptophan fluorescence emission maximum to a longer wavelength and reduces the intensity as illustrated in Fig. 3.2B (Kuloglu et al., 2002). Thus, changes in the conformational equilibrium can be detected by monitoring the fluorescence intensity at
335 nm. We speculated that time-resolved optical spectroscopy could be used to monitor the response of an Ltn10/Ltn40 mixture to an increase in salt concentration or the addition of a glycosaminoglycan like heparin. To demonstrate the feasibility of this approach, we performed a series of salt-jump experiments with the intrinsic fluorescence emission of W55. Although monitoring the emission at 335 nm, we injected buffer, concentrated NaCl, or low molecular weight heparin into a sample of 15 mM Ltn (Fig. 3.2C). The jump in NaCl concentration from 0 to 250 mM produced an increase in fluorescence intensity consistent with a shift toward Ltn10. In contrast, injection of heparin produced a decrease in intensity that we interpreted as an increase in the relative concentration of Ltn40.

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Figure 3.2 Spectroscopic detection of structural interconversion. (A) Exchange peaks between the Ltn10 and Ltn40 1H-15N resonances of Gly44 were detected with a 2-D 15 N zz-exchange spectrum acquired on 250 mM wild-type Ltn in 20 mM sodium phosphate, pH 6.0, 150 mM NaCl. Exchange spectra were measured with mixing times of 50, 150, 200, 250, 300, 350 (shown), 400, 500, 750, and 900 msec. Intensities of the peaks were fit to a two-state exchange model yielding rates of 0.4 and 1.2 s1 for the forward (Ltn40 ! Ltn10) and reverse (Ltn10 ! Ltn40) reactions, respectively. (B) Tryptophan emission spectrum of wild-type Ltn (15 mM) at 30  C in 0 mM NaCl, where the Ltn10:Ltn40 ratio is roughly 1:1. Arrows indicate how fluorescence intensity increases and the emission maximum shifts to lower wavelengths as the tryptophan is buried in the Ltn10 core, whereas increased solvent exposure and flexibility in Ltn40 decreases the intensity and causes a red shift. (C) Time-dependent fluorescence intensity at 335 nm for Ltn samples (15 mM, 30  C, 0 mM NaCl) injected at time 0 with buffer (), 15 mM low molecular weight heparin (▲), or 200 mM NaCl (e). Equilibration curves were fit to a simple exponential decay with ProFit software yielding Ltn40 ! Ltn10 and Ltn10 ! Ltn40 exchange rates of 0.22 and 0.73 s1, respectively.

As described in the following, this result is consistent with a model in which cell-surface GAGs bind preferentially to the Ltn40 species. Rates for the Ltn40 ! Ltn10 and Ltn10 ! Ltn40 reactions at 30  C (0.22 and 0.73 s1, respectively) obtained from nonlinear fitting are slightly lower than those measured by 2-D exchange NMR at 37  C. Although the fluorescence approach may be improved by use of a stopped flow apparatus with a shorter dead time for sample mixing, the general agreement of the preliminary results from fluorescence titrations and 2-D exchange NMR validates both methods for kinetic analysis of the Ltn conformational change. Future studies of the

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temperature dependence of kex will permit us to define the height of the energetic barrier (activation energy, Ea) separating the Ltn10 and Ltn40 species.

5. Engineering Conformationally Restricted Lymphotactin Variants Conveniently, the Ltn structure is highly sensitive to variations in temperature and salt concentration within the ranges typically used for biophysical measurements on proteins (e.g., 10 to 45  C, 0 to 200 mM NaCl), and this allowed us to resolve the two conformations present at physiologic conditions. However, assays for biologic activity with animals or cell cultures impose constraints on the buffer composition and temperature. These limited our ability to assign specific functional roles to the Ltn10 and Ltn40 species, because both would be present in significant amounts when attempting to measure intracellular calcium flux, chemotaxis, or cell recruitment in vivo. To solve this problem, we generated a panel of Ltn variants designed to favor either the Ltn10 or Ltn40 structure and prevent interconversion. Although the 3-D structure of Ltn40 had yet not been solved, its secondary structure was known (Kuloglu et al., 2002), and we used the structures of Ltn10 and the chemokine HCC-2 to design two different disulfide crosslinks that would prevent the rearrangement to Ltn40 as described previously (Tuinstra et al., 2007). Amino acid substitutions corresponding to the CC1-Ltn and CC3-Ltn variants are shown in Fig. 3.3. We took a different approach in designing an Ltn40-restricted variant: selectively destabilize the Ltn10 fold. Because residues 52 to 93 are disordered in Ltn40, we identified W55 as an essential hydrophobic core

Figure 3.3 Ltn variants stabilize either the Ltn10 or Ltn40 structure. The introduction of a second disulfide bond to stabilize the Ltn10 conformation was based on alignment of Ltn10 with CC chemokines. Construction of CC1-Ltn (T10C þ dipeptide insertion G32-AC-S33) is based on alignment with the first disulfide with RANTES, whereas the positions of the cysteine substitutions in CC3-Ltn (V21C/V59C) mimic the unusual third disulfide in HCC-2. Stabilization of the Ltn40 conformation is accomplished by introducing an aspartate or alanine point mutation at W55. Loss of the tryptophan side chain disrupts the hydrophobic core and destabilizes the Ltn10 conformation. Mutated residues are shown in white or black outline.

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residue in Ltn10 that makes no contribution to the Ltn40 structure and changed it to an alanine or aspartic acid (Fig. 3.3) (Tuinstra et al., 2008). All of the mutants were successfully expressed and purified, and we examined their 2-D 1H-15N HSQC spectra. For an objective measure of their similarity to one structure or the other we plotted the 1H and 15N shifts for each mutant against those of Ltn10 and Ltn40 (Fig. 3.4A). The W55D mutant matches the Ltn40 shifts very closely and shows a poor correlation with Ltn10, and HSQC spectra of the mutant proteins acquired at different temperatures show no evidence of interconversion to the other conformation (Fig. 3.4B), and similar results were obtained for the W55A mutant (not shown). Conversely, the nearly perfect correlation with Ltn10 shifts for the CC1 and CC3 variants show convincingly that the disulfide-locked proteins retain the chemokine-like fold, and we confirmed this by solving the NMR structure of CC3 (Fig. 3.4C) (Tuinstra et al., 2007). We next used the CC3-Ltn and W55D-Ltn proteins in a series of functional studies designed to assign functional roles to the Ltn10 and Ltn40 states.

6. Functional Analysis of Distinct Ltn Native State Conformations Chemokine function depends on dual biochemical activities, GAG binding, and GPCR activation. In the case of lymphotactin, the two functional roles may be segregated by a conformational barrier corresponding to the activation energy for Ltn10 $ Ltn40 interconversion. Treating the CC3 and W55D variants described previously as ‘‘pure’’ versions of Ltn10 and Ltn40, respectively, we compared their ability to activate the Ltn receptor XCR1 stably expressed on the surface of HEK293 cells (generously provided by Joseph Hedrick, Schering-Plough Research Institute) (Shan et al., 2000). As detailed by Tuinstra et al. (2007), changes in intracellular calcium levels in response to each protein at a concentration of 200 nM were monitored with cells loaded with the Fluo-3-AM dye at 37  C with a PTI spectrofluorometer. CC3-Ltn and wild-type Ltn induced similar calcium flux responses (Tuinstra et al., 2007), whereas W55D-Ltn was completely inactive (Tuinstra et al., 2008). To compare relative GAG binding activity for the Ltn10 and Ltn40 species, we eluted CC3-Ltn and W55D-Ltn from a heparin-Sepharose column with a sodium chloride concentration gradient (Tuinstra et al., 2008). Wild-type Ltn elutes in two broad fractions at
450 and 700 mM NaCl (Fig. 3.5 and Table 3.1), suggesting that the two conformations bind heparin with significantly different affinities. CC3-Ltn elutes at a single sharp peak at 450 mM NaCl, whereas the broader W55D-Ltn elution resembles the high-affinity fraction of wild-type Ltn with a peak at 750 mM NaCl.

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CC1 140

A

5.0

CC3 9.0

7.0

5.0

W55D

7.0

9.0

5.0

7.0

9.0 9.0

1HN 15

120

100

140 100 120 15N chemical shift

10 ⬚C

100

120

7.0

140

(ppm)

45 ⬚C

C Native disulfide bond

120 125 125

120

W55D

115 110 15N chemical

shift (ppm)

CC3

115

110

B

140

5.0

100

W55

1H

9.0

140

5.0

100

7.0

120 120

D58 G32

chemical shift (ppm)

Ltn10 Ltn40

H

9.0

8.0 1H

7.0

6.0

9.0

8.0

7.0

Engineered disulfide bond M73

6.0

chemical shift (ppm)

Figure 3.4 CC3-Ltn and W55D-Ltn are structurally indistinguishable from Ltn10 and Ltn40. (A) Amide chemical shifts for CC1-, CC3-, and W55D-Ltn plotted against the corresponding values for wild-type Ltn at 10 and 40  C. The excellent correlation of the Ltn10 chemical shifts with CC1-Ltn and CC3-Ltn compared with the poor correlation with the Ltn40 chemical shifts suggests that CC1-Ltn and CC3-Ltn adopt the chemokine-like fold. The converse is true for W55D-Ltn, demonstrating its similarity to Ltn40. Outliers in each plot are labeled and correspond to residues at or adjacent to the mutated positions in the amino acid sequence. (B) 1H-15N HSQC spectra of CC3-Ltn and W55D-Ltn acquired at 10 and 45  C in 20 mM sodium phosphate (pH 6.0). Comparison of the low- and high-temperature spectra indicates that CC3-Ltn only adopts the Ltn10 conformation and W55D-Ltn only adopts the Ltn40 conformation. (C) Ribbon diagram of CC3-Ltn showing the canonical chemokine fold. The structure was determined at 25  C in 20 mM sodium phosphate, a solution condition that normally contains a mixed population of the two Ltn conformers. The disulfide bond incorporated to restrict the conformation is shown.

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Lymphotactin Structural Dynamics

CC3

1.0

1

WT 2

W55D

R65S

R23A

2 1 1

R43A

R43A

1

2

0.25

2

R23A

0.5

K66A

2 R23A/R43A

0.75

R23A/R43A

Fractional population

K66A R65S

1

0 300

400

500 600 [NaCl] (mM)

700

800

Figure 3.5 Dual Ltn conformations bind heparin with different affinities. Elution of wild-type Ltn from a heparin-Sepharose column yields a biphasic profile with a lowand high-salt peak. In contrast, CC3-Ltn and W55D-Ltn elute as a single fraction corresponding to the low- and high-salt wild-type Ltn peaks, respectively. Alanine substitution of two arginines involved in GAG binding, R23 and R43, shifts the elution of the high-affinity (Ltn40) fraction. The high/low affinity peak ratio for R43A is skewed toward the Ltn10 conformation. This effect was more pronounced in the R23A/R43A double mutant, favoring the Ltn10 conformation by a ratio of 2:1. In contrast, mutation of residues that do not participate in heparin binding (R65A and K66S) had a minimal effect on the elution profiles or peak ratios when compared with wild-type Ltn.

Table 3.1 Elution from heparin-sepharose and relative intensities for lymphotactin variants

Ltn mutant

WT CC3 W55D R23A R43A R23A/ R43A R65S K66A

Peak 1 elution (mM NaCl)

Peak 2 elution (mM NaCl)

Fractional population peak 1

Fractional population peak 2

452 430 N.D. 382 376 309

694 N.D. 753 557 458 344

40 100 0 41 53 71

60 0 100 59 47 29

415 408

658 659

29 38

71 62

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9.8

9.4

9.0

8.6

9.4 9.8 9.0 8.6 1H chemical shift (ppm)

9.8

9.4

9.0

chemical shift (ppm)

1:1 W55D:CC3 250 mM heparin

15N

1:1 W55D:CC3 50 mM heparin

1:1 W55D:CC3 No heparin

128 127 126 125 124 123

Not surprisingly, our results showed that only the chemokine-like Ltn10 structure is capable of XCR1 signaling, and we learned that the Ltn40 dimer encodes tight GAG binding, whereas the Ltn10 species binds heparin with low but detectable affinity. To show that cell surface GAGs interact preferentially with Ltn40, we attempted to use 2-D NMR as the equivalent of a pull-down or coprecipitation assay. Initially, we added low molecular weight heparin to an NMR sample of [U-15N]-wild-type Ltn at conditions where both conformations are present, expecting to see an effect on the Ltn40 peaks only. Instead, HSQC signals for both species diminished and disappeared with increasing amounts of heparin. As illustrated by fluorescence spectroscopy in Fig. 3.2C, the addition of heparin to an equilibrium mixture of Ltn10 and Ltn40 causes a shift toward the Ltn40 state. At NMR concentrations, heparin forms insoluble complexes with Ltn40 and probably shifts the conformational equilibrium until all the protein has precipitated. To allow for selective heparin binding to either the Ltn10 or Ltn40 species without the potential for interconversion, we performed the same titration on a mixture of 15N-labeled CC3-Ltn and W55D-Ltn (Tuinstra et al., 2008). HSQC spectra of wild-type Ltn and the CC3/W55D mixture HSQC are virtually identical, but on addition of a heparin tetrasaccharide to the mixed sample, only the Ltn40 signals were lost (Fig. 3.6). Collectively, these studies support a model for lymphotactin activity in which the native state is coupled to two different binding equilibria, either of which can alter the Ltn10–Ltn40 conformational equilibrium (Fig. 3.7). In this model, cell surface GAGs can inhibit Ltn activity by shifting the equilibrium toward the

8.6

Figure 3.6 Selective heparin binding monitored by 2-D NMR. HSQC spectra recorded on an NMR sample containing 250 mM each of CC3-Ltn and W55D-Ltn at 37  C in 20 mM sodium phosphate (pH 6.0). Before the addition of heparin tetrasaccharide (left panel), 1H-15N HSQC signals are observed for W55D-Ltn (gray, Ltn40 conformation) and CC3-Ltn (black, Ltn10 conformation). Addition of heparin tetrasaccharide results in broadening (center panel) and disappearance of the W55D-Ltn resonances (right panel), whereas the CC3-Ltn resonances are unaffected.

65

Keq < 1 ΔG ⬚ = 0

XCR1 Ltn40

Lymphocyte

Ltn10

Ltn40

Keq < 1 ΔG ⬚ < 0

Ltn10

Keq < 1 ΔG ⬚ > 0 Ltn40

Cell-surface GAGs

Lymphotactin Structural Dynamics

Ltn10

Figure 3.7 Functional relevance of the Ltn conformational equilibrium. Under physiologic conditions, Ltn partitions equally between the Ltn10 and Ltn40 conformations. The conformational equilibrium may shift in response to interactions with cell-surface GAGs or XCR1. Cell-surface GAGs stabilize Ltn40 relative to Ltn10, whereas XCR1 binding may shift the equilibrium toward the Ltn10 conformer.

Ltn40 state and reducing the concentration of Ltn10 available for XCR1 activation. Presumably, there must be a mechanism for shifting the equilibrium toward the Ltn10 state when conditions require XCR1-mediated signaling and T cell chemotaxis. Whether the XCR1 receptor can shift the conformational equilibrium directly has not been investigated, and other factors that alter the Ltn10–Ltn40 balance may be identified in future in vivo studies of lymphotactin function.

7. GAG Binding Residues are Linked to the Ltn Conformational Equilibrium In a previous study, we substituted most of the lysine and arginine residues in Ltn with alanine and used surface plasmon resonance (SPR) to measure heparin binding affinities. We identified R23 and R43 as the residues that contributed most to GAG binding, because each alanine substitution reduced the affinity for heparin by
30-fold (Peterson et al., 2004). R23 and R43 are adjacent in the Ltn10 structure (Fig. 3.1C), but as described earlier, heparin-Sepharose binding studies showed that Ltn40 supplies the GAG binding function. Because SPR measurements provide no information on the relative affinities of the two Ltn structural species, we ran the panel of arginine and lysine mutants over the heparin column and noted the elution of the high- and low-affinity fractions. The results are illustrated schematically in Fig. 3.5 and summarized in Table 3.1. For substitutions that have little or no effect on GAG binding (e.g., R65S and K66A), the position and relative intensities of the two peaks are similar to wild type. As expected, the high-affinity fractions of R23A and R43A elute at significantly lower salt concentrations (557 and 458 mM NaCl, respectively) than wild-type Ltn40 (694 mM NaCl). Interestingly, the peak

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ratio for R43A was skewed toward the low affinity fraction compared with wild-type Ltn (Table 3.1), and the effect was even more pronounced for the R23A/R43A double mutant, which favored the low-affinity fraction by a ratio of more than 2:1. Elimination of key residues for high-affinity GAG binding seems to shift the conformational equilibrium toward Ltn10, but this could simply reflect a diminished capacity for heparin to stabilize Ltn40 and shift the equilibrium in the opposite direction. To determine whether substitutions at R23 and R43 alter the relative stabilities of Ltn10 and Ltn40 independent of heparin, we compared the HSQC spectra of wild-type Ltn and the R23A, R43A, and R23A/R43A mutants. Except for signals from the substituted residues, the HSQC spectrum of each mutant is superimposable with the spectrum of wild-type Ltn, thus the tertiary structure is unchanged. As shown in Fig. 3.8, wild-type Ltn readily converts from Ltn10 to Ltn40 as the temperature is increased from 10 to 45  C. However, substitution of either R23 or R43 with alanine seems to stabilize the Ltn10 species and shift the Ltn40 transition to higher temperatures (data not shown). The effect is more

A

B

45 ⬚C

9.0

8.0

chemical shift (ppm)

7.0

6.0

0.2 0.0 1.0

LTN40 0

10

20

30

40

50

30

40

50

LTN10

0.8 0.6

120

Fractional population

115 120 125 1H

6.0

chemical shift (ppm)

110 7.0

0.4

0.4

125

115 8.0

LTN10

0.6

15N

WT R23A/R43A 9.0

1.0 0.8

110

10 ⬚C

0.2 0.0

LTN40 0

10

20

Temperature (⬚C)

Figure 3.8 Mutation of GAG binding residues alters the Ltn10–Ltn40 equilibrium. Alanine substitution of R23 and R43 stabilize the Ltn10 conformer. (A) 1H-15N HSQC spectra of wild-type and R23A/R43A-Ltn acquired at 10 and 45  C in 20 mM sodium phosphate (pH 6.0). Comparison of the low- and high-temperature HSQCs indicates that the R23A/R43A double mutant stabilizes the Ltn10 conformation in the absence of salt. (B) Relative intensities of the W55 1He1 peak in the Ltn10 and Ltn40 conformations is plotted as a function of temperature. The plots indicate the equilibrium midpoint for wild-type Ltn is
25  C. In contrast, the equilibrium midpoint for R23A/R43A is shifted to
50  C under conditions that typically favor the Ltn40 conformation.

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Lymphotactin Structural Dynamics

+ R23 R23 +

R23 +

Cl−

R43

+

R43

+

R43

+

Figure 3.9 Ionic strength dependence in the Ltn structural equilibrium. Electrostatic repulsion in Ltn10 is relieved by salt or conversion to Ltn40 conformer. In the Ltn10 ˚ ) for their guanidinyl groups to conformation R23 and R43 are close enough (<8A coordinate a chloride ion (left panel). In the absence of salt, electrostatic repulsion between the positively charged side chains destabilizes the Ltn10 conformation (center panel). Alternately, the R23-R43 repulsion can be eliminated by conversion to the Ltn40 conformer (right panel).

pronounced for the R23A/R43A double mutant that remains predominantly in the Ltn10 form even at 45  C (Fig. 3.8). How does alanine substitution of one or two solvent-exposed loop residues affect the conformational equilibrium so dramatically? A recent molecular dynamics study suggests the close proximity of R23 and R43 in the Ltn10 conformation may be a destabilizing factor in low ionic strength solution. In a 14-nsec MD simulation performed at 45  C in the presence of 200 mM NaCl, Cui and coworkers found that the positively charged R23 and R43 side chains coordinate a chloride ion that also forms hydrogen bonds with the T41 hydroxyl and backbone amides from the 40’s loop (Formaneck et al., 2006). Chloride association with R23, R43, and the 40’s loop persisted for the last 5 nsec of the simulation, illustrating a specific protein–ion interaction that would favor the Ltn10 conformation. In the absence of a stabilizing anion, electrostatic repulsion between the nearby R23 and R43 side chains is relieved by the Ltn10 ! Ltn40 rearrangement ˚ which separates the two positively charged residues by a distance of
20 A (Fig. 3.9). Replacement of either R23 or R43 reduces the repulsive effect, diminishing the need for anion stabilization and favoring the Ltn10 structure relative to Ltn40.

8. Conclusions Unique features of the lymphotactin sequence distinguish it from other chemokines. Structural analysis revealed an unprecedented nativestate interconversion between two unrelated structures, each of which contributes to the biologic activity of Ltn. In this work, we showed that

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the Ltn10 and Ltn40 structures interconvert with a frequency of
1 s1 with fluorescence and NMR experiments. Data from mutagenesis, heparin affinity chromatography, and NMR interpreted in the context of published MD simulations suggested that electrostatic repulsion in the Ltn10 structure between key GAG-binding residues could be stabilized by anion binding in solutions of high ionic strength. Our long-term goals are to understand each step in the process of structural interconversion between the Ltn10 and Ltn40 states and to relate this unusual conformational equilibrium to lymphotactin activity in vivo. The experimental approaches and results described here serve as a foundation for more detailed studies of lymphotactin dynamics with a combination of mutagenesis, optical and NMR spectroscopy, and MD simulations.

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and oligomerization are essential for the in vivo activity of certain chemokines. Proc. Natl. Acad. Sci. USA 100, 1885–1890. Rousseau, R. F., Haight, A. E., Hirschmann-Jax, C., Yvon, E. S., Rill, D. R., Mei, Z., Smith, S. C., Inman, S., Cooper, K., Alcoser, P., Grilley, B., Gee, A., et al. (2003). Local and systemic effects of an allogeneic tumor cell vaccine combining transgenic human lymphotactin with interleukin-2 in patients with advanced or refractory neuroblastoma. Blood 101, 1718–1726. Scheerens, H., Hessel, E., de Waal-Malefyt, R., Leach, M. W., and Rennick, D. (2001). Characterization of chemokines and chemokine receptors in two murine models of inflammatory bowel disease: IL-10/ mice and Rag-2/ mice reconstituted with CD4þCD45RBhigh T cells. Eur. J. Immunol. 31, 1465–1474. Seibert, C., Cadene, M., Sanfiz, A., Chait, B. T., and Sakmar, T. P. (2002). Tyrosine sulfation of CCR5 N-terminal peptide by tyrosylprotein sulfotransferases 1 and 2 follows a discrete pattern and temporal sequence. Proc. Natl. Acad. Sci. USA 99, 11031–11036. Seibert, C., Veldkamp, C. T., Peterson, F. C., Chait, B. T., Volkman, B. F., and Sakmar, T. P. (2008). Sequential tyrosine sulfation of CXCR4 by tyrosylprotein sulfotransferases. Biochemistry 47, 11251–11262. Shan, L., Qiao, X., Oldham, E., Catron, D., Kaminski, H., Lundell, D., Zlotnik, A., Gustafson, E., and Hedrick, J. A. (2000). Identification of viral macrophage inflammatory protein (vMIP)-II as a ligand for GPR5/XCR1. Biochem. Biophys. Res. Commun. 268, 938–941. Stuart, A. C., Borzilleri, K. A., Withka, J. M., and Palmer, A. G., 3rd., (1999). Compensating for variations in 1H-13C scalar coupling constants in isotope-filtered NMR experiments. J. Am. Chem. Soc. 121, 5346–5347. Tikhonov, I., Kitabwalla, M., Wallace, M., Malkovsky, M., Volkman, B., and Pauza, C. D. (2001). Staphylococcal superantigens induce lymphotactin production by human CD4þ and CD8þ T cells. Cytokine 16, 73–78. Tollinger, M., Skrynnikov, N. R., Mulder, F. A., Forman-Kay, J. D., and Kay, L. E. (2001). Slow dynamics in folded and unfolded states of an SH3 domain. J. Am. Chem. Soc. 123, 11341–11352. Tuinstra, R. L., Peterson, F. C., Elgin, E. S., Pelzek, A. J., and Volkman, B. F. (2007). An engineered second disulfide bond restricts lymphotactin/XCL1 to a chemokine-like conformation with XCR1 agonist activity. Biochemistry 46, 2564–2573. Tuinstra, R. L., Peterson, F. C., Kutlesa, S., Elgin, E. S., Kron, M. A., and Volkman, B. F. (2008). Interconversion between two unrelated protein folds in the lymphotactin native state. Proc. Natl. Acad. Sci. USA 105, 5057–5062. Veldkamp, C. T., Peterson, F. C., Hayes, P. L., Mattmiller, J. E., Haugner, J. C., 3rd., de la Cruz, N., and Volkman, B. F. (2007). On-column refolding of recombinant chemokines for NMR studies and biological assays. Protein Expr. Purif. 52, 202–209. Veldkamp, C. T., Seibert, C., Peterson, F. C., De la Cruz, N. B., Haugner, J. C., 3rd., Basnet, H., Sakmar, T. P., and Volkman, B. F. (2008). Structural basis of CXCR4 sulfotyrosine recognition by the chemokine SDF-1/CXCL12. Sci. Signal. 1, ra4. Veldkamp, C. T., Seibert, C., Peterson, F. C., Sakmar, T. P., and Volkman, B. F. (2006). Recognition of a CXCR4 sulfotyrosine by the chemokine stromal cell-derived factor1alpha (SDF-1alpha/CXCL12). J. Mol. Biol. 359, 1400–1409. Wang, J. D., Nonomura, N., Takahara, S., Li, B. S., Azuma, H., Ichimaru, N., Kokado, Y., Matsumiya, K., Miki, T., Suzuki, S., and Okuyama, A. (1998). Lymphotactin: A key regulator of lymphocyte trafficking during acute graft rejection. Immunology 95, 56–61. Yoshida, T., Imai, T., Kakizaki, M., Nishimura, M., Takagi, S., and Yoshie, O. (1998). Identification of single C motif-1/lymphotactin receptor XCR1. J. Biol. Chem. 273, 16551–16554.

C H A P T E R

F O U R

Interactions of Chemokines with Glycosaminoglycans Damon J. Hamel,* India Sielaff,† Amanda E. I. Proudfoot,† and Tracy M. Handel* Contents 1. Introduction 2. Methods to Detect, Quantify, and Characterize Chemokine: Gag Interactions 2.1. Biochemical and in vivo methods 2.2. In vivo cellular recruitment 3. Biophysical Methods 3.1. Isothermal fluorescence titration 3.2. Surface plasmon resonance (SPR) 3.3. Sedimentation equilibrium analytical ultracentrifugation 3.4. NMR: Heteronuclear single quantum correlation (HSQC) spectroscopy 3.5. Fourier-transform ion cyclotron resonance mass spectrometry (FT-ICR MS) 3.6. Mass spectrometry methods for characterizing GAG-binding epitopes on chemokines: Proteolytic footprinting 3.7. Emerging mass spectrometry methods for characterizing protein:GAG interactions: Hydrogen deuterium exchange and radiolytic oxidation 4. Summary Acknowledgments References

72 80 80 85 87 87 88 90 91 93 95

96 97 98 98

Abstract Many proteins require interactions with cell surface glycosaminoglycans (GAGs) to exert their biologic activity. The effect of GAG binding on protein function ranges from essential roles in development, organogenesis, cell growth,

* {

Skaggs School of Pharmacy and Pharmaceutical Science, University of California, San Diego, La Jolla, California, USA Merck Serono Geneva Research Centre, Geneva, Switzerland

Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05404-4

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2009 Elsevier Inc. All rights reserved.

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cell adhesion, inflammation, tumorigenesis, and interactions with pathogens. A classic example is the role of GAGs in the interaction of fibroblast growth factors with their receptors, where GAGs play a role in specificity determination and control of receptor-ligand engagement. The other well-studied example involves the binding of antithrombin to heparin/heparan sulfate, which results in the inactivation of the coagulation cascade. In view of their specialized activity in cellular recruitment, chemokines interact with GAGs, minimally as a mechanism for localization of chemokines to specific anatomical spaces enabling them to act as directional signals for migrating cells. The biological relevance of these interactions has been recently demonstrated by functional characterization of mutants that are deficient in GAG binding. These mutants bind receptor normally in vitro but are unable to recruit cells in vivo. Observations like this have motivated investigations to identify GAG-binding epitopes on chemokines, the specificity and affinity of chemokines for different GAGs, the oligomerization of chemokines on GAGs, and the efficacy of GAG-binding mutants in the context of in vivo cell recruitment and animal models of disease. To this end, several techniques have been developed to measure the interactions of chemokines with GAGs. In this chapter we describe these various assays with particular reference to those that have been used to assess the binding of chemokines to GAGs and to define their epitopes. In the end, we believe both in vitro and in vivo characterization are absolutely necessary for understanding these interactions and their biologic relevance in the context of the whole organism.

1. Introduction In the past several years, it has become increasingly apparent that to function properly, chemokines require interaction not only with G protein– coupled receptors (GPCRs) but also with the carbohydrate moieties (glycosaminoglycans or GAGs) of proteoglycans on endothelial cells and the extracellular matrix (Handel et al., 2005; Johnson et al., 2005). The biological relevance of GAG interactions was first convincingly demonstrated with engineered variants of chemokines (RANTES/CCL5, MCP-1/CCL2, and MIP1b/CCL4) that had mutations which inhibited their ability to bind to heparin, yet minimally perturbed their ability to bind to their respective receptors and promote cell migration in vitro. Nevertheless, these mutants were ineffective in inducing cell migration in vivo, in contrast to their WT counterparts, demonstrating the importance of the GAG interaction (Proudfoot et al., 2003). Since these studies, related experiments with GAG-binding mutants of MCP-3/CCL7 (Ali et al., 2005) and Lymphotactin/XCL1 (Peterson et al., 2004) have also demonstrated the requirement of GAG interactions for chemokine-induced cell migration in vivo. Minimally, these interactions are thought to provide a mechanism for the localization

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and concentration of chemokines on cell surfaces, which in turn provides guidance cues for migrating cells; in the absence of such interactions, rapid diffusion of chemokines would occur, especially in the presence of blood flow, dissipating directional gradients and resulting in chemokine concentrations below the threshold required to activate their receptors. The importance of these interactions has been further underscored by the ability of GAG-binding mutants of chemokines to block the action of wild type chemokines both in normal and pathological states. For example, [44AANA47]-RANTES, a GAG-binding mutant of CCL5/RANTES, was shown to be a dominant negative inhibitor of cell recruitment by the WT protein, both by preventing higher order oligomerization and by forming nonfunctional heterodimers with WT RANTES wherein only some of the subunits in the hetero-oligomer have intact GAG epitopes ( Johnson et al., 2004). This same mutant also attenuated atherosclerotic plaque formation in LDLr
/
mice by inhibiting leukocyte recruitment into plaques, producing a less inflammatory and more stable plaque phenotype (Braunersreuther et al., 2008). A GAG-binding mutant of MCP-3/CCL7 similarly has been shown to inhibit chemokine-mediated recruitment in murine air pouches and to inhibit the chemotactic activity of synovial fluid from patients with rheumatoid arthritis (Ali et al., 2005). Studies of viral mechanisms of immunomodulation also underscore the importance of GAG interactions. Many viruses produce chemokine-binding proteins (CBPs) that inhibit the activity of chemokines; for example, the murine poxvirus pathogen, ectromelia virus E163, secretes a 31-kDa glycoprotein that selectively binds a number of CC and CXC chemokines with high affinity, and in the case of SDF-1a/CXCL12 and IP-10/CXCL10, functions by blocking their GAG-binding epitopes (Ruiz-Arguello et al., 2008). Advantageously, E163 also binds directly to GAGs with high affinity, allowing it to remain in the vicinity of sites of viral infection, ready to interfere with the function of locally secreted chemokines. Other promiscuous CBPs include T7 from the myxoma rabbit poxvirus, which blocks GAG interactions of chemokines from the CXC, CC, and C families (Lalani et al., 1997), whereas M3 is a murine herpesvirus CBP that binds and inactivates chemokines from all four families (van Berkel et al., 2000). M3 is particularly effective in neutralizing chemokine function; in the case of the CC chemokine, MCP-1/CCL2, it blocks the GAG-binding epitopes and the receptorbinding epitopes of the chemokine and interferes with the ability of CCL2 to oligomerize (Alexander et al., 2002) (another functionally important interaction of chemokines described later and in Chapter 2). More details of virally encoded chemokine-binding proteins and methods for their study are given in Chapters 9 and 10 in volume 460. The interaction of chemokines with GAGs is unsurprising from a structural point of view, because almost all chemokines are highly basic proteins with many Arg, Lys, and His residues on their surfaces, whereas

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GAGs are acidic, sulfated linear polysaccharides that typically bind proteins rich in these amino acids. Interestingly, the two chemokines that are acidic, MIP-1a/CCL3 and MIP-1b/CCL4, both bind GAGs, albeit less strongly than basic chemokines. Linear epitopes for GAG binding have been defined on chemokine surfaces such as XBBXBX and XBBBXXBX motifs (where B represents a basic amino acid), suggesting common features of proteinGAG recognition; however, chemokines are known to oligomerize on GAGs, thereby spatially diversifying the nature of the binding epitopes (Handel et al., 2005). Figure 4.1 shows the distribution and diversity of GAG-binding epitopes on the surfaces of dimeric structures of several chemokines. Adding to the complexity, Fig. 4.2 shows the distribution of GAG-binding epitopes on three different tetrameric forms of a single chemokine, IP-10/CXCL10, which may allow this chemokine to Chemokine

GAG-binding

Residues

MCP-1/CCL2

R18, K19, R24, K49, K58, H66

MIP-1a /CCL3

R18, K45, R46, K48

Rantes/CCL5

R44, K45, R47

SDF-1a/CXCL12

K24, H25, K27, R41, K43

IL-8/CXCL8

K20, R60, K64, K67, R68

Lymphotactin/XCL1

R23, R43

Figure 4.1 Illustration of GAG-binding epitopes of chemokines defined by mutagenesis coupled with many of the assays outlined in this chapter. This figure shows surface topology models of chemokine homodimers. The GAG-binding epitopes, as defined by mutagenesis, are shown in black and listed on the right. They are taken from the following references: MCP-1/CCL2) (Hemmerich et al., 1999; Jarnagin et al., 1999), MIP-1a/CCL3 (Koopmann and Krangel, 1997), RANTES/CCL5 (Proudfoot et al., 2001), SDF-1/CXCL12 (Amara et al., 1999; Sadir et al., 2001), 1L-8/CXCL8 (Kuschert et al., 1998), and Lymphotactin/XCL1 (Peterson et al., 2004). Images were generated with PyMol.

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Figure 4.2 Different tetrameric structures of IP-10/CXCL10 have been solved and are suggested to be associated with differential GAG binding. Left, Human IP-10 M form with MCP-1/PF4 like quaternary structure. Middle, Human IP-10 H form. Right, Murine IP-10. The top row contains backbone traces plus the secondary structure of each form. The middle row contains a surface representation in the same orientation as the top row, with GAG-binding epitopes in black. The bottom row has been rotated 180 about the X-axis. PDB entries are listed below the name of each tetrameric form. Images were generated with PyMol.

selectively interact with different types of GAGs. This variability in the nature of the binding sites suggests that in addition to providing mechanisms for localization of chemokines, the GAG interaction may add to the specificity of chemokine function (Handel et al., 2005; Hoogewerf et al., 1997; Johnson et al., 2005; Webb et al., 1993). For example, although many chemokines bind and activate the same receptor, GAG interactions may selectively recruit specific chemokines to cell surfaces and tissues to fine tune the immune response. Indeed, cells can alter their GAG coats in the context of pathological states like cancer, possibly to selectively recruit certain chemokines, cytokines, and growth factors to modulate phenotype (Sasisekharan et al., 2002). The biologic importance of chemokine-GAG interactions and the discovery that GAG-deficient chemokines can have therapeutic potential has motivated many studies to define the GAG-binding epitopes on chemokines and to characterize the structure, affinity, and specificity of these interactions. However, these studies are technically challenging for many reasons. Unlike proteins or nucleic acids, which are fairly homogeneous, GAGs are exceptionally heterogeneous polymers with respect to both length and composition. Considering composition alone, it has been

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suggested that more than 12 billion sequences are possible for an oligosaccharide of six disaccharide units, representing 100 times the variation of a hexapeptide and 2 million times that of a nucleic acid hexamer (Shriver et al., 2002). This diversity derives from the nontemplate nature of GAGs and the fact that the GAG biosynthetic reactions do not always proceed to completion. These same characteristics also make them difficult to produce recombinantly and to characterize them analytically and structurally compared with nucleic acids and proteins. The six major classes of GAGs include heparin, heparan sulfate (HS), chondroitin sulfate (CS), dermatan sulfate (DS), keratin sulfate (KS), and hyaluronic acid (HA). However, most studies use heparin as a model system (and to a lesser extent heparan sulfate), because it is more easily obtained and cheaper than other GAGs. As an interesting side note, heparin is more highly sulfated than heparan sulfate, the most abundant GAG expressed on virtually every cell in the body. Heparin (and HA) are also soluble GAGs, whereas HS, DS, KS, and CS are usually covalently attached to a protein core, forming an overall structure referred to as proteoglycan. Traditional dogma suggests that the GAG chains are the functional ligand-binding units of proteoglycans. However, there is emerging evidence that the core proteins can also impact ligand binding through interactions that are both direct and indirect (i.e., the spacing of the GAG chains on the protein) (Herndon et al., 1999). Thus the use of ‘‘model reagents’’ is a major caveat of studies of chemokine-GAG interactions and may be limiting the ability of investigators to truly get at issues of specificity and binding affinity for example. Nevertheless, the use of heparin as a representative GAG has been quite successful in defining GAG-binding epitopes on chemokines (Ali et al., 2005; Amara et al., 1999; Campanella et al., 2003; Koopmann et al., 1999; Koopmann and Krangel, 1997; Lau et al., 2004b; Peterson et al., 2004; Proudfoot et al., 2001). Bearing in mind that the use of heparin is not always optimal, in this chapter, we describe many of the assays and techniques that have been used to characterize chemokine-GAG interactions, along with their advantages and disadvantages (Table 4.1). These include both biochemical and biophysical methods. With respect to the biochemical assays, in our experience, it seems that some assays are better than others for rank ordering GAG-binding hotspots and determining chemokine:GAG affinities, and where appropriate we comment on this and offer plausible explanations for discrepancies. In general, we have found that the use of several assays in combination is best for obtaining consensus results, with the ultimate test of biologic relevance being the use of in vivo cell migration experiments to evaluate carefully designed GAG-binding–deficient mutants compared with WT chemokines. The biophysical assays, described in the second section, are generally more demanding and require sophisticated equipment and expertise. However, these methods, and extensions not included here,

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Table 4.1 Summary of various methods described in this chapter to study chemokine: GAG interactions, listing advantages and disadvantagesa Method

Advantages

Disadvantages

Heparin affinity chromatography

Rapid and easy; little protein required

Equilibrium competition binding

Fast and very reliable; IC50 and Kd determinations are feasible; can be used to determine binding epitopes on chemokines; competitors can be chemokines or GAGs; one can observe GAGinduced oligomerization Fast and simple; not biased toward charged amino acids for identification of GAG-binding epitopes on chemokines in contrast to the heparin affinity assay Easy; little protein required; no specialized equipment needed; can customize binding surface with GAG of interest so the assay is not limited to heparin

True affinities are not measured; results are biased toward identifying charged amino acids as binding epitopes Requires radioactively labeled chemokine

Tritiated heparin binding assay

Enzyme-linked immunosorbent saturation binding assay

Requires radioactively labeled GAGs; less reliable than other assays like equilibrium competition

Provides relative affinities rather than true affinities

(continued)

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Table 4.1 (continued) Method

Advantages

Disadvantages

In vivo cellular recruitment

Provides the only assay of biologic relevance in vivo; can be used to prioritize the importance of GAGbinding epitopes; can be used to test chemokine or GAGbased inhibitors in competition studies Easy and no labeled material is required; good for relative ranking of chemokine mutants

Minimal quantitative data

Isothermal fluorescence titration

Surface plasmon resonance (SPR)

Sedimentation equilibrium analytical ultracentrifugation

Can get kinetic rates of association and disassociation in addition to equilibrium constants; can get information on oligomerization; in principle, one can customize the sensor chip with the GAG of interest Provides information on stoichiometries, equilibrium constants; can discriminate quite well between different models of chemokine:GAG interaction

Consumes a fair amount of protein; Kd’s are apparent because of oligomerization; chemokine:GAG aggregation can be problematic Experimental execution and data analysis can be quite complicated; expertise required

Experimental execution and especially data analysis can be complicated; a common problem with chemokine: GAG complexes is time-dependent aggregation

x

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Table 4.1 (continued) Method

Advantages

Disadvantages

Heteronuclear single quantum correlation (HSQC) spectroscopy

Provides amino acid resolution information on GAG-binding epitopes on chemokines.

Fourier transform ion cyclotron resonance mass spectrometry (FT-ICR MS)

A very powerful and emerging technique for studies of chemokine:GAG complexes; it is possible to detect many different complexes in terms of stoichiometries and compositions caused by the mass accuracy of the technique; can be used to select highaffinity GAGs and determine their composition A powerful method to relatively quickly identify GAG binding epitopes on chemokines; little protein is required

Limited to studies with size and compositionally defined GAGs caused by NMR issues with heterogeneity; with longer GAGs, solubility can be problematic Not quantitative; requires very expensive equipment and significant expertise

Proteolytic footprinting coupled with mass spectrometry

a

Low resolution unless effort is taken to use multiple proteases; expensive equipment and expertise is required

All methods suffer from reagent limitations resulting in the fact that most studies use heparin. Radiolytic oxidation and HDMS are not included in the table because they have not yet been used to characterize chemokine: GAG interactions.

particularly involving mass spectrometry, represent important future directions for the chemokine:GAG field. Indeed, a burning issue is to understand how much specificity is really encoded by chemokine:GAG interactions. For this we must be able to identify and isolate or synthesize high-affinity ligands from complex GAG mixtures so that we can move beyond studies with heparin.

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2. Methods to Detect, Quantify, and Characterize Chemokine: Gag Interactions 2.1. Biochemical and in vivo methods 2.1.1. Heparin affinity chromatography A commonly used method to determine the relative ‘‘affinity’’ of chemokines for GAGs is to determine the amount of salt needed for elution of the chemokine from a heparin sepharose column. For example, a comparison of several chemokines showed the following rank order of affinities for heparin Sepharose: RANTES/CCL5 > I-TAC/CXCL11 > SDF-1/CXCL12 > IL-8/CXCL8 > MCP-1/CCL2 > MIP-1a/CCL3 > MIP-1b/CCL4 (Handel et al., 2005). To determine the contribution of specific residues of a given chemokine to its GAG-binding epitope, the salt concentration required for elution of alanine point mutants are compared with that of the WT protein, yielding per residue values for D[NaCl]H. Often, these measurements are done in parallel with determining the amount of salt required to elute mutants from a nonspecific S-sepharose column (D[NaCl]S) and compared with the elution from the heparin sepharose column. This provides a measure of the specificity of the protein-heparin interaction by accounting for nonspecific electrostatics. The specificity index is related to DDNaCl as calculated from the formula that follows, where the superscripts H and S refer to elution from heparin sepharose or S-sepharose column, respectively (Kuschert et al., 1998; Lau et al., 2004a,b).

DD½NaCl ¼ D½NaClH
D½NaClS where D[NaCl]H ¼ D[NaCl]H WT
D [NaCl]H mutant and D[NaCl]S ¼ DNaCl]S WT
D[NaCl]S mutant For example, in the preceding rank order for binding to heparin sepharose, although RANTES required the highest concentration of NaCl for elution, the order when bound to S-sepharose was I-TAC/CXCL11 > SDF-1/CXCL12 > RANTES/CCL5 > IL-8/CXCL8 > MCP-1/CCL2 > MIP-1a/CCL3 > MIP-1b/CCL4, placing RANTES in third place. However, when converted to the specificity index, the results indicate the highest specificity of RANTES for this GAG (Handel et al., 2005). An advantage of the heparin affinity chromatography method is that it is fast, inexpensive and uses relatively small amounts of protein. However, it is not a true measure of affinity. Because salt is used for elution, the method is primarily useful for identifying amino acids involved in electrostatic interactions (although the electrostatic component is accounted for, at least in part, by determining the specificity index for a given amino acid). Nevertheless, this approach has been successfully used to characterize the

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GAG-binding epitopes of several chemokines including CXCL8 (Kuschert et al., 1998), CCL2, (Lau et al., 2004b), CCL5 (Proudfoot et al., 2001), and MIP-1a (Koopmann and Krangel, 1997), and when compared with other methods such as heparin competition binding (described later), it seems to give consistent results (Lau et al., 2004b). This method could be adapted to characterizing interactions with other GAGs besides heparin, although such columns are not commercially available at the present time. 2.1.2.1. Materials and methods This assay only requires a chromatography system, a conductivity meter, heparin sepharose and cation exchange columns, recombinant chemokine, and standard laboratory buffers. The amount of chemokine depends on the detection limit of UV absorbance at 280 nm, but reliable results are obtained with 50 to 100 mg on a standard FPLC system. The amounts of protein can be reduced if detection is conducted at 214 nm. The protein is applied to the heparin sepharose column, previously equilibrated in the buffer of choice (typically 50 mM TRIS/HCl, pH 7.5), and eluted with a linear gradient of 0 to 2 M NaCl in the same buffer. The salt concentration at which the protein elutes is measured with a conductivity meter. An equivalent amount of chemokine is treated under the same conditions with a cation exchange column. The amount of salt needed for elution of protein from both columns is compared, and the difference is the heparin specificity index as defined in the previous equations.

2.1.3. Equilibrium competition binding To determine more accurate affinities of chemokines for GAGs compared with the heparin affinity chromatography assay, classical equilibrium competition–binding assays can be used. These assays also use an immobilized heparin format. Heparin sepharose beads are incubated with the radioactively labeled chemokine of interest and are competed off with increasing concentrations of the GAG (Kuschert et al., 1999). Alternately, it is also possible to use unlabeled chemokine instead of GAG as the competitor. In both cases, one would expect to observe that addition of competitor would simply compete off the radiolabeled chemokine, and this is exactly what is observed with GAG as competitor. However, such assays with chemokines as competitors often cause the recruitment of additional radiolabeled chemokine onto the immobilized heparin because of oligomerization of chemokines on GAGs (Hoogewerf et al., 1997) as shown in Fig. 4.3 (Handel et al., 2008). This assay, therefore, serves not only as a good method for determining binding affinities of different chemokines for heparin and the relative ability of different types of GAGs to compete for chemokine binding to heparin, but it is also an excellent qualitative screen of GAG-induced chemokine oligomerization.

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% Hot ligand bound

200

150

100

50

0 −12

−11

−10

−9

−8

−7

−6

−5

−4

Log [cold ligand]

Figure 4.3 Equilibrium competition binding of MCP-1/CCL2 (open circles) and a nonoligomerizing variant [P8A]-MCP-1 (closed circles) on immobilized heparin. 125Ilabeded MCP-1 was incubated with heparin Sepharose beads, and unlabeled MCP-1 or [P8A]-MCP-1 was added. The increase in bound 125I- labeled MCP-1 on addition of WT MCP-1 is diagnostic of oligomerization, whereas only competitive inhibition is observed with [P8A]-MCP-1. Reprinted with permission from: ‘‘Journal of Leukocyte Biology, 2008 84:1101–8; An engineered monomer of CCL2 has anti-inflammatory properties emphasizing the importance of oligomerization for chemokine activity in vivo; Handel, T. M., Johnson, Z., Rodrigues, D. H., Dos Santos, A. C., Cirillo, R., Muzio, V., Riva, S., Mack, M., De´ruaz, M., Borlat, F., Vitte, P. A., Wells, T. N., Teixeira, M. M., Proudfoot, A. E.’’

A disadvantage of this method is the use of radioactively labeled material, which is expensive and needs a specifically designated working space. However, the method is fast and quite reliable and is, therefore, a frequently used strategy. For example, it was used to define the GAG binding site of RANTES (Proudfoot et al., 2001) and then to demonstrate that interference with heparin binding and with oligomerization presents a novel anti-inflammatory strategy ( Johnson et al., 2004). This method has also been used to demonstrate the presence of an unusually high affinity GAG-binding site for I-TAC/CXCL11 (Sielaff et al., 2009). Overall, we find that the equilibrium competition–binding assay is a straightforward and useful assay for characterizing chemokine:GAG interactions for many applications. 2.1.3.1. Materials and methods This assay requires laboratory equipment routinely used for equilibrium competition–binding assays with iodinated tracers such as a shaker to accommodate 96-well plates, a vacuum filtration apparatus for 96-well plates, and a Microbeta Scintillation Counter for 96-well plates. Required reagents include 125I-labeled chemokine, heparin-sepharose beads, 96-well filter plates, nonprotein adsorbing 96-well plates, the desired competitors such as soluble GAGs or unlabeled chemokine, and standard laboratory buffers.

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A dilution series of competitor (unlabeled chemokine or GAG) is prepared in binding buffer (50 mM TRIS, pH 7.5, 1 mM CaCl2, 5 mM MgCl2, 0.5% BSA), ideally in threefold dilutions spanning 4 to 5 orders of magnitude to achieve 12 data measurements in a nonprotein binding 96-well plate; 25 ml of the dilutions is transferred to a filter plate in triplicate (NB: if RANTES/CCL5 is used in the assay, the binding buffer must be supplemented with 0.15 M NaCl to prevent oligomerization). To achieve a final assay volume of 100 ml, 25 ml of binding buffer is added to each well. The iodinated chemokines are generally obtained with a specific activity of 2000 Ci/mmol. We recommend reconstituting them in 500 ml of water to obtain a 23 nM stock, which is diluted in binding buffer to 0.4 nM so that the addition of 25 ml of the solution to each well of the filter plate results in a final concentration of 0.1 nM radiolabeled chemokine. 1 g of heparin sepharose beads is rehydrated in 10 ml of water and left at room temperature for 60 min. The beads are then diluted 400-fold with binding buffer and 25 ml is added to each well. The plates are then incubated for 4 h at room temperature. To remove unbound ligand, the plates are washed three times with 200 ml wash buffer (binding buffer supplemented with 0.15 M NaCl or 0.5 M NaCl in the case of RANTES/CCL5) in a 96-well plate filter device. After the addition of 50 ml of scintillation liquid per well, the radioactivity is determined with a calibrated Microbeta Scintillation Counter (Perkin Elmer). 2.1.4. Tritiated heparin binding assay The tritiated heparin assay is a commonly used biochemical assay that measures the binding of the protein of interest to soluble tritiated heparin. The radioactive heparin is incubated with increasing concentrations of protein in 96-well plates fitted with protein binding cellulose phosphate paper. The wells are washed to remove unbound GAG and the amount of radioactivity retained in the filter is determined. Compared with the heparin affinity chromatography assay, which only accounts for the electrostatic interactions, this assay has the advantage that it reflects the overall binding capacity for heparin of a protein or protein mutant. This method is often used in conjunction with alternative methods to obtain a more complete profile of protein-GAG interactions, for example for the identification of the GAG binding site of MCP-1/CCL2 (Lau et al., 2004b) or for the determination of GAG binding to monomeric chemokine variants (Proudfoot et al., 2003). However, large amounts of protein are required for the assay, and the results can be somewhat variable. Furthermore, inconsistencies compared with the heparin sepharose affinity assay and an isothermal fluorescence titration assay (described later) have been observed (Lau et al., 2004b) that may reflect the varying efficiencies of different protein mutants to bind to the filter.

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2.1.4.1. Materials and methods The assay requires a Microbeta Scintillation Counter (Perkin Elmer), a shaker, and a vacuum filtration system to accommodate 96-well plates. The required reagents include the sodium salt of 3H-heparin, recombinant chemokines, nonprotein adsorbing 96-well plates, protein-adsorbing P-81 Whatman 96-well filter plates, scintillation liquid, and phosphate-buffered saline (PBS). A dilution series of the unlabeled chemokines is prepared in PBS in a nonprotein adsorbing 96-well plate in triplicate, spanning 4 to 5 orders of magnitude. Each well should contain the same volume, ideally 50 ml; 50 ml of 3H-heparin solution in PBS is then added to each well. 3H-heparin sodium salt is supplied by Perkin Elmer at a specific activity of 0.2 to 1.0 mCi/mg in aliquots of 1 mCi. This amount is dissolved in 500 ml H2O and subsequently diluted 1:600 in PBS for use in the assay. The plate is then incubated at 37  C for 1 h. After incubation, 20 ml from each well is transferred into the corresponding wells of a P-81 Whatman filter plate. The filter plate is washed three times with PBS with a vacuum system, and 50 ml of scintillation fluid is added per well. The radioactivity is then determined with a calibrated Microbeta Scintillation Counter.

2.1.5. Enzyme-linked immunosorbent saturation binding assay A more recently developed assay is the saturation binding assay with EpranEx plates. The plates have a special coating that binds heparin or other GAGs such as heparin sulfate (HS). Because of the inability of heparin to adhere to the surface of conventional polystyrene microplates, the development of high-throughput ELISA-like assays that use surface immobilized heparin had previously been hampered. The Plasso EpranExTM plate provides a surface onto which heparin can be immobilized, and the immobilized heparin is then capable of capturing heparin-binding proteins. The principle of this assay is similar to an ELISA. The GAG solution is incubated overnight in the plate and, after a wash step, the protein of interest is added. Detection of binding is achieved with a primary antibody to the protein and a secondary antibody labeled with horseradish peroxidase (HRP), or with a biotinylated primary antibody, followed by ExtrAvidin detection. The assay uses relatively small amounts of protein and GAGs, it is easy to perform, and no specialized equipment is required. It has been increasingly reported in the recent literature, for example, to demonstrate the selectivity of Eotaxin/CCL11 for heparin over other GAGs (Ellyard et al., 2007) and the reduced affinity toward GAGs of citrullinated IL-8/CXCL8 (Proost et al., 2008). A significant advantage of this assay is the ability to investigate interactions with different types of GAGs because of the ability to prepare custom-coated plates. Furthermore, it is amenable to inhibition studies with soluble GAGs as competitor. The assay allows determinations of relative affinities, for example, to study chemokine mutants to identify the GAG

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binding site. Although KD determinations are possible in principle, the exact amount of immobilized GAG, presented in the correct active orientation, cannot be determined. 2.1.5.1. Materials and methods This assay requires standard laboratory equipment, namely a microplate reader and a shaker to accommodate 96-well plates. The necessary reagents are: EpranEx plates (Plasso, Sheffield, UK), GAG (e.g., heparin, recombinant chemokines, a biotinylated primary antibody, ExtrAvidin solution [Sigma]), and NaCl, NaOAc, Tween 20, gelatin, and nonprotein binding 96-well plates. For the coating of the EpranEx plate with GAG, 200 ml of a 25 mg/ml heparin (or other GAG) solution in PBS is added to each well, and the plate is then incubated overnight at room temperature in the dark. The liquid is then discarded and the plate washed three times with standard assay buffer (SAB, 100 mM NaCl, 50 mM NaOAc, 0.2 % (v/v) Tween 20; pH7.2); 250 ml of blocking solution (0.2 % [w/v] gelatin in SAB) is then added to each well, the plate is incubated for 1 h at room temperature and then washed three times with SAB as before. A dilution series of chemokine in SAB, spanning 4 to 5 orders of magnitude, is prepared in a nonprotein binding 96-well plate; 100 ml of the dilution series is transferred in triplicate to the EpranEx plate, and the plate is incubated for 2 h at room temperature. The wells are then washed three times with SAB. To detect the amount of bound chemokine, 200 ml of an antibody in blocking solution is added to each well (diluted according to instructions for ELISA). The plate is incubated for 1 h and washed three times with SAB; 200 ml of ExtrAvidin-AP in blocking solution (1:10,000) is added to each well, the plate is incubated for 30 min at room temperature, and washed three times with SAB; 200 ml of developing reagent is added per well, the plate is incubated for 40 min at room temperature, and the absorbance is measured at 405 nm in a microplate reader. As an alternative to the ExtrAvidin solution, streptavidin-conjugated horseradish peroxidase may be used for detection (Kadi et al., 2006). Furthermore, it is possible to use this assay for competition experiments using a dilution series of competitor, such as heparin or other GAGs, and a constant amount of chemokine.

2.2. In vivo cellular recruitment Use of the preceding biochemical assays with mutants of chemokines has allowed the identification of potential GAG-binding epitopes on the chemokines. However, none of these assays prove whether GAG binding or the identified GAG-binding epitopes are biologically relevant. Thus a common strategy has been to couple the in vitro assays with in vivo assays.

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Chemokines have been used to induce cellular recruitment into different cavities in vivo: an artificially created air pouch (Ramos et al., 2003), the pleural cavity (Pinho et al., 2007), or the joint (Deruaz et al., 2008). We have used a fairly simple approach involving a chemokine-induced peritoneal cellular recruitment assay (Proudfoot et al., 2003). Although the details may vary with chemokine, a typical procedure involves the following: (1) inject chemokine or saline control into the peritoneal cavity of mice, (2) sacrifice mice after a few to several hours, and (3) lavage the cavity and count the number of recruited cells. Parameters to optimize include the duration between injecting and sacrificing the mice, the amount of chemokine, and whether the mice are presensitized (Sielaff et al., 2009). One can also coinject GAG-mutants with wild-type chemokine; this approach has revealed that a GAG mutant of RANTES acts as a dominant negative inhibitor of cell recruitment of the WT protein ( Johnson et al., 2004). In addition, GAGs themselves can be used as competitors, and such experiments provide additional information on GAG specificity (Ellyard et al., 2007). These experiments have the advantage over in vitro experiments in that they provide a readout of biological relevance and can be used to rank order the importance of different chemokine epitopes on GAG binding, as long as mutation of the epitopes do not, or minimally affect, receptor binding. Otherwise, impaired cell migration in response to GAG-binding mutants may be due to effects on GAG binding, receptor binding, or both. A significant disadvantage of in vivo assays is the requirement for large amounts of chemokine, and thus the ability to produce recombinant protein is advised over use of commercially available material. Other in vivo assays have been used to address the relevance of GAG binding such as the murine air pouch assay (Ali et al., 2005; Das et al., 1998; Ellyard et al., 2007). For example, Ali and coworkers used this assay to demonstrate that a GAGbinding mutant of MCP-3/CCL7 could inhibit migration not only to WT CCL3, but also to a receptor sharing chemokine, RANTES/CCL5, and the nonreceptor sharing chemokine, SDF-1/CXCL12 (Ali et al., 2005). 2.2.1. Materials and methods We describe here the method for chemokine-induced peritoneal recruitment. For this assay recombinant chemokines are required, and we have used 8- to 12-week-old female Balb/c mice. 0.1 to 100 mg of chemokine or mutant diluted in 0.2 ml of sterile, lipopolysaccharide-free 0.9% NaCl solution is injected i. p. If used in the experiment, antagonists are administered either i.v. or s.c. The mice are sacrificed after 4 to 18 h by CO2 asphyxiation, depending on the choice of chemokine stimulant. The peritoneal cavity is washed three times with 5 ml PBS, and the washes are pooled. The resulting solution of peritoneal cells is centrifuged at 1500 rpm for 5 min and the cells are resuspended in 1 ml of PBS and counted.

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3. Biophysical Methods This section describes biophysical approaches for characterizing chemokine: GAG interactions. Because several of these methods are technically involved and require more optimization and expertise than the biochemical assays described previously, protocols are not provided for some of these techniques. However, references are provided as guides.

3.1. Isothermal fluorescence titration The simplest of the biophysical methods for characterizing chemokine: GAG interactions is the isothermal fluorescence titration assay (Goger et al., 2002). In this assay, one monitors the change in fluorescence intensity from a Trp residue, which is highly conserved in most chemokines, on addition of GAG. If the protein of interest does not possess a Trp, one can generally replace a Phe or Tyr with a Trp, provided that the mutation does not affect biological activity (Sielaff et al., 2009). The form of the data is a binding isotherm from which an apparent binding constant can be determined as illustrated in Fig. 4.4. Comparison of different mutants in this assay allows one to define residues that contribute to the GAG-binding epitopes, the role of oligomerization on GAG-binding affinity, and the effect of different GAGs on binding affinity (Lau et al., 2004b). The assay is very straightforward, relatively rapid and reproducible, and access to the requisite fluorescence equipment is generally not an issue. This is a recommended approach for affinity measurements because of its simplicity in both execution and interpretation, with the proviso that the measurements are deemed ‘‘apparent,’’ because they can be complicated by changes in avidity caused by concentration-dependent oligomerization of the chemokine. 3.1.1. Materials and methods This assay requires a fluorescence spectrophotometer, heparin, or other GAG, recombinant chemokine, and standard laboratory buffers. The sensitivity is limited by the fluorescence emission at 340 nm, which is quenched on addition of GAG, but dissociation constants between 20 nM and 750 mM have been reported (Goger et al., 2002; Lau et al., 2004b). In previous studies of IL-8/CXCL8, different concentrations of chemokine were used to bias the chemokine to a monomeric (50 nM ) versus dimeric (700 nM) form (Goger et al., 2002), whereas in a study of MCP-1/CCL2, a 1 mM solution of chemokine in PBS was used (Lau et al., 2004b). High-concentration GAG stocks are required to minimize dilution when added to the chemokine solution, and the concentration is calculated from the average molecular weight for the GAG under study (another less

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Figure 4.4 Isothermal fluorescence titration of heparin with WT MCP-1/CCL2 compared with the monomeric variant P8A-MCP-1 and the GAG-binding deficient mutant K18A/R19A MCP-1. The data show the effect of oligomerization on GAGbinding affinity and identify a double mutant with significantly impaired ability to bind heparin. Reprinted from: ‘‘Journal of Biological Chemistry, 2004 279:22294–22305; Identification of the glycosaminoglycan binding site of the CC chemokine, MCP-1: implications for structure and function in vivo; Lau, E. K., Paavola, C. D., Johnson, Z., Gaudry, J. P., Geretti, E., Borlat, F., Kungl, A. J., Proudfoot, A. E., Handel, T. M.’’ with permission from ASBMB.

accurate parameter). The GAG solution is titrated into a fixed concentration of chemokine, and the fluorescence emission is recorded with excitation and emission wavelengths of 282 and 340 nm, respectively, and a 290-nm cutoff filter. The resulting binding isotherms are then fit by nonlinear regression to an equation describing a bimolecular (or other) association reaction, as described in Goger et al. (2002). Again, we refer to the resulting number as an apparent affinity because of the assumptions made in curve fitting to a bimolecular reaction when changes in aggregation state may occur.

3.2. Surface plasmon resonance (SPR) A powerful tool for the study of protein-protein and protein-ligand interactions is an evanescent biosensor technology referred to as surface plasmon resonance (SPR). The technology is based on an optical phenomenon that enables detection of unlabeled binding partners in real time (Schuck, 1997). Effectively, the technique depends on the refractive index of the sample within the evanescent field above the sensor surface. Adsorption or

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desorption of macromolecules at the sensor surface change the local refractive index and produce a shift in the measured resonance angle that, to a good approximation, is proportional to the surface concentration of macromolecules. Operationally, one binding partner is immobilized on a chip. A solution containing the putative binding partner is then flowed over the chip at a constant rate, and the association kinetics of the two binding partners is detected by the change in mass as a function of time. Then in a dissociation phase, buffer is used instead, and the time course of complex dissociation is monitored. In this way it is possible to determine the association/dissociation kinetics of the complex, and from these, an equilibrium dissociation constant can be calculated. This method has been used in several studies for the characterization of chemokine-GAG interactions (Alexander-Brett and Fremont, 2007; Amara et al., 1999; Kawashima et al., 2003; Vives et al., 2002). In the study by Vives et al., SPR was used to analyze the interaction between an N-terminally truncated form of CCL5/RANTES ([9 to 68]-RANTES) and heparin sulfate immobilized on the chip surface (Vives et al., 2002). Although a monomer in solution, the results demonstrated a complex binding model involving dimerization of [9 to 68]-RANTES on heparin sulfate, with positive cooperativity such that the first CCL5 molecule associated with HS with an affinity of 398 nM followed by a second CCL5 molecule with an affinity of 84 nM, even though [9 to 68]-RANTES is a monomer in solution. These results suggest that the chemokine:GAG interaction is facilitated by oligomerization and vice versa, which seems to be a characteristic of many chemokines. The immobilzation of a protein on a surface bears the risk of abolishing partly or completely its binding ability, and the immobilization chemistry must, therefore, be carefully considered. GAGs, however, are readily immobilzed on streptavidin SA sensor chips surfaces (Biacore, GE Healthcare) by biotinylation (Osmond et al., 2002). The immobilzation of GAGs on the surface should not alter their binding characteristics, because they are often attached to cell surfaces by means of proteoglycans in vivo. An advantage of the method is that small amounts of material are needed, and labeling is not required for the protein whose binding is to be detected. Furthermore, it is possible to detect the association or competition with a third binding partner. For example, SPR has been used to examine the competition of the viral chemokine decoy receptor M3 with chemokines binding to heparin (Alexander-Brett and Fremont, 2007). However, one must have access to the requisite expensive Biacore (GE Healthcare) equipment. 3.2.1. Materials and methods For this assay a Biacore instrument is needed. The reagents used are recombinant chemokines, GAGs, biotinylation reagent, streptavidin sensor chips (SA chips), HEPES-buffered saline (HBS), and glycine buffers at

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pH 2 (both from Biacore, GE healthcare) and NaCl. Note: other buffers may be recommended for certain Biacore instruments. The GAGs are biotinylated as previously described (Osmond et al., 2002), dissolved in HBS buffer at 1 to 50 mg/ml and immobilized on the streptavidin chip by flowing the solution over the chip at 10 ml/min to achieve at least 500 to 1000 RU units. The chip is then ready for binding experiments. The chemokines are dissolved at 1 mg/ml and flowed over the chip at 30 ml/ml. Regeneration of the chip is achieved by flowing glycine buffer, pH2, containing 0.5 to 1.0 M NaCl at 30 ml/min over the chip.

3.3. Sedimentation equilibrium analytical ultracentrifugation Analytical ultracentrifugation (AUC), specifically the sedimentation equilibrium technique, is a powerful method for determining the mass of a protein or protein complex in solution and can be used for characterizing associating systems, including subunit stoichiometries and equilibrium constants for the assembly process. AUC is extensively described in Chapter 2 on methods for characterizing chemokine oligomerization, and, therefore, the reader is referred to that chapter for a more detailed summary of the physical basis of the technique. A sample is subject to high centrifugal force in an analytical ultracentrifuge. One then optically measures the distribution of the protein along the radius of the sample cell at equilibrium, from which one can extract molecular mass, and for interacting systems, the dissociation constant (Balbo et al., 2007b). Protein and protein complexes with masses in the range from <103 to >106 Daltons, and associating systems characterized by KD values between 10
4 to 10
8 M
1 can be studied by AUC (Balbo et al., 2007a). Although this method has most frequently been used to characterize chemokine oligomerization and the effect of mutations (Laurence et al., 2000; Paavola et al., 1998), it can also be used to characterize the interaction of chemokines with GAGs. In the case of MCP-1/ CCL2, AUC and NMR studies have shown that this chemokine exists as a dimer in solution (Handel and Domaille, 1996). However, the addition of heparin octasaccharide very clearly induces the chemokine to form tetramers, consistent with a crystal structure form (Lau et al., 2004b; Lubkowski et al., 1997). By contrast, an engineered monomeric mutant, [P8A]-MCP-1 did not oligomerize in the presence of the GAG, even though it still bound GAG (Lau et al., 2004b). These structural properties were correlated with the ability of the WT chemokine but not the monomeric variant to induce cell migration in vivo because of the requirement of both oligomerization and GAG-binding for in vivo function (Proudfoot et al., 2003). One caveat to AUC is that it is time consuming, taking 1 to 2 days per run. Furthermore, the data can be difficult to fit, particularly when there are issues with nonequilibrium aggregation or precipitation, which often occurs in the presence of larger GAGs (hexasaccharides and larger). For example,

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the addition of heparin octasaccharide to Eotaxin/CCL11 causes the timedependent formation of large aggregates that gradually fall out of solution (Handel, unpublished data). When solubility is favorable, as for MCP-1, one must still be careful to calculate the partial specific volume of the complex, and the dissociation constants must be such that one can assume 100% binding, or a concentration series must be collected and fit. Nevertheless, for many chemokines, the relevant concentration ranges (high nanomolar to submicromolar) are generally adequate for these purposes. Fig. 4.5 shows an example of the use of AUC that revealed that CCL2 forms tetramers in the presence of an octasaccharide. It also illustrates how the goodness of the fit, reflected by the residuals, can be used to discriminate between different models of assembly. Access to the appropriate equipment, such as a Beckman Optima XL-A ultracentrifuge and associated curvefitting software, is the basic requirement for conducting these studies.

3.4. NMR: Heteronuclear single quantum correlation (HSQC) spectroscopy Heteronuclear single quantum coherence (HSQC) spectroscopy is an NMR method in which one detects 1H nuclei and their attached 13C or 15N heteronuclei. A 2-dimensional HSQC spectrum thus provides a ‘‘fingerprint’’ of the protein in the way of chemical shifts or cross peaks from 1H-15N atoms from the backbone amides and side chain NH groups 2 (in a 1H-15N HSCQ) or of the 1H-13C groups from both the backbone and side chains (1H-13C HSCQ) in a protein. Fig. 2.4A in Chapter 2 shows an example of a 1H-15N HSCQ, where each cross peak corresponds to an NH (or NH2) group from a different amino acid in the protein. The power of the use of HSQC spectra to monitor interactions of chemokines and GAGs is that the position of the cross peaks (i.e., the chemical shifts) are exquisitely sensitive to the magnetic environment, and perturbations caused by GAG binding can be readily identified by changes in chemical shift. Such changes can, therefore, provide residue specific information on the region of the protein surface where the GAG is binding. This method has been used quite frequently to monitor interactions of chemokines with small saccharides to identify binding sites on the chemokine and to identify features of the GAG, like the presence and position of sulfate groups, which are important for the interaction. For example, titrating different GAGs into IL-8/CXCL8 revealed a heparin-binding site that includes the C-terminal a-helix and a proximal loop encompassing residues 18 to 23 (Kuschert et al., 1998). Similarly, with differentially sulfated disaccharides, it was possible to show the dependence of the interaction on the composition of the GAG on binding and amino acids involved in the binding surface (McCornack et al., 2003), which agreed well with prior studies based on mutagenesis (Laurence et al., 2001).

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A limitation of this method is that one must use GAGs that have defined composition and size, and because these are generally only available for small saccharides, studies have been limited to the use of small GAGs. Compositional and size heterogeneity usually causes exchange broadening and makes the spectra uninterpretable. Longer GAGs can also cause precipitation at the concentrations (50 mM and above) used for NMR. However, because chemokines tend to prefer binding to larger GAGs as indicated by experiments with isothermal calorimetry ([Kuschert et al., 1999], another technique only mentioned here), the use of small GAGs will not capture a picture of the entire binding surface and, instead, may identify nonspecific interactions. Nevertheless, these methods are useful. Isotopically labeled protein is also required, but making labeled protein has been straightforward for chemokines. The experiments are quite easy to collect and interpret, although they require prior assignment of the spectra, which may be complicated for the nonexpert. However, many chemokine structures have already been solved by NMR, and assignments are available in the PDB that can serve as a starting point for such studies. Looking toward the future, as methods for making compositionally defined GAGs improve, these types of NMR experiments and others (Zhuang et al., 2006) will become increasingly more powerful for characterizing chemokine: GAG interactions.

3.5. Fourier-transform ion cyclotron resonance mass spectrometry (FT-ICR MS) Fourier-transform ion cyclotron resonance (FT-ICR) mass spectrometry is a useful method for studying noncovalent chemokine:GAG interactions as well as oligomerization of chemokines (Crown et al., 2006; Yu et al., 2005). As described in Chapter 2, which is focused on the oligomerization of chemokines, this method is facilitated by electrospray ionization (ESI), which enables transfer of intact noncovalent complexes from solution to the gas phase. FT-ICR has very high mass accuracy and can be used to Figure 4.5 Sedimentation equilibrium of MCP-1/CCL2 in the absence (top left) and presence (top right) of a heparin octasaccharide at a 1:1 stoichiometry. Below the data sets, a comparison is shown of residual plots for different models generated from the þoctasaccharide data set. The best fit is determined by the randomness of the distribution of residuals and by minimization of the variance. Here, the monomer-tetramer equilibrium seems to be the best model to describe the data, because the other models exhibit residuals with systematic nonrandom deviations, indicative that they are inaccurate. Reprinted from: ‘‘Journal of Biological Chemistry, 2004 279:22294–22305; Identification of the glycosaminoglycan binding site of the CC chemokine, MCP-1: implications for structure and function in vivo; Lau, E. K., Paavola, C. D., Johnson, Z., Gaudry, J. P., Geretti, E., Borlat, F., Kungl, A. J., Proudfoot, A. E., Handel, T. M.’’ with permission from ASBMB.

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determine the molecular weight of complexes, stoichiometry, etc. With FT-ICR, Crown et al. analyzed the ability of the MCP-1 ligands (MCP-1/ CCL2, MCP-2/CCL8, MCP-3/CCL7, MCP-4/CCL13) and eotaxin/ CCL11 to homo- and hetero-dimerize and the influence of GAGs on this process (Crown et al., 2006). Figure 4.6 shows an example of a spectrum of a sample containing MCP-1/CCL2 þ MCP-2/CCL8 þ the pentasaccharide Arixtra. The high mass accuracy allows easy identification of all of the species, and although exact quantitation is an issue because measurements are done in the gas phase, the intensities of the various species usually reflect the relative population in solution. In these studies, it was possible to show that interaction with GAGs typically promoted hetero-oligomerization. For example, MCP-3/CCL11 does not interact with MCP-2/CCL8 in the absence of GAG, whereas a strong heterodimer complex with one bound pentasaccharide (Arixtra) was observed when the GAG was added. In related studies it was also shown that FT-ICR MS could be used in combination with affinity purification to select and identify chemokine-binding octasaccharides from complex mixtures (Schenauer et al., 2007). Thus this method has great potential for identifying determinants of chemokine: GAG specificity and affinity focused on the composition of the GAG. FT-ICR measurements can be done quite rapidly, and the method requires very little protein and GAG. However, in addition to the previously mentioned issues with quantitation, access to the appropriate and very [CCL2 + CCL2 + arixtra]8+ [CCL2 + CCL8 +arixtra]9+ [M(CCL2)]5+ M4+/D8+(CCL2) [M(CCL8)]5+ [CCL2+ CCL8]8+ [M(CCL2)]6+ [CCL2 + CCL8]9+ [M(CCL8)]6+

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Figure 4.6 ESI-FTICR mass spectra of 10 mM MCP-1/CCL2 plus 10 mM MCP-2/ CCL8 plus 10 mM of the defined pentasaccharide, Arixtra, in 100 mM NH4OAc (pH 6.8). M refers to the monomer species and D to the dimer species. For example, [M(CCL8)]6þ is the þ6 ion of the MCP-2/CCL8 monomer, [CCL2þCCL8]9þ is the þ9 ion of the CCL2/CCL8 heterodimer and [CCL2 þ CCL8 þ Arixtra]8þ is the þ8 ion of the CCL2/CCL8 heterodimer in complex with Arixtra pentasaccharide. Reprinted from: ‘‘Journal of Biological Chemistry, 2006 281:25438–46; Heterodimerization of CCR2 chemokines and regulation by glycosaminoglycan binding; Crown, S. E., Yu, Y., Sweeney, M. D., Leary, J. A., Handel, T. M.’’ with permission from ASBMB.

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expensive instrumentation and the need for mass spectrometry expertise are barriers to routine use of this technique. However, mass spectrometry is emerging as the most powerful method for analytical characterization of carbohydrates that bind with high affinity to chemokines.

3.6. Mass spectrometry methods for characterizing GAGbinding epitopes on chemokines: Proteolytic footprinting Apart from understanding whether proteins interact with GAGs and defining their corresponding affinities, it is often useful to know which regions of a protein form the binding site(s). This information is crucial for understanding, and potentially modulating, protein:GAG interactions. Most methods described previously can be used for this purpose by analyzing point mutants of chemokines; however, although powerful, mutagenesis is a labor-intensive approach. Proteolytic footprinting is a sophisticated method that allows for the identification of regions of a protein that bind to GAGs without the absolute requirement for mutagenesis. The protein of interest, complexed to a GAG of interest, is subjected to digestion with trypsin. The resulting peptides are subsequently analyzed by mass spectrometry and compared with those obtained after a digest of the free protein (Falsone et al., 2007). Regions that are complexed with the GAG are protected from digestion and are thus identified compared with digests in the absence of heparin (or other GAG). This method was used to confirm the GAG binding site of I-TAC/CXCL11 as being principally located in a region encompassing the 50’s loop (53CLNPKSKQAR62), because this region was digested in the absence of heparin but remained intact in the presence of heparin (Sielaff et al., 2009). Although this technique enables the rapid identification of general GAG-binding regions on the chemokine, it does not provide amino acid resolution. The use of additional proteases in addition to trypsin, however, can improve the resolution. 3.6.1. Materials and methods For this assay a nanoHPLC-MS/MS instrument is required. It also requires recombinant chemokine, heparin, or other GAGs, trypsin, NH4HCO3 buffer, NaCl, and formic acid. For the GAG-footprinting experiments, chemokines are proteolytically digested with trypsin for 1 h at 20  C, in an enzyme to substrate ratio of 1:10 (w/w). The digestion is performed in the presence and in the absence of a 10 molar excess of heparin or another GAG in 50 mM NH4HCO3, pH 8.0, containing 150 mM NaCl to suppress nonspecific interactions. Proteolytic cleavage is carried out at 37  C for 18 h. The digest is then stopped by adjusting the pH to 2.5 with formic acid. The resulting peptide fragments are subsequently identified by nanoHPLC-MS/MS as described previously

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(Falsone et al., 2007). Potential GAG-binding epitopes on the chemokine are identified as undigested (protected) peptides in the presence of heparin that are digested in the absence of heparin.

3.7. Emerging mass spectrometry methods for characterizing protein:GAG interactions: Hydrogen deuterium exchange and radiolytic oxidation Footprinting methods related to the proteolysis approach described above include hydrogen deuterium (H/D) exchange and radiolytic oxidation mapping (Takamoto and Chance, 2006; Xu and Chance, 2007). These methods have not yet been applied to chemokine:GAG interactions let alone many other protein:GAG interactions. However, they are emerging methods and, therefore, worth mentioning. In hydrogen/deuterium amide exchange mass spectrometry (HDMS or DXMS), protein is buffer-exchanged from a medium containing H2O to the same buffer containing D2O (Mandell et al., 2005). The labile hydrogen atoms of the backbone amides exchange with solvent at a rate dependent on several parameters, including solvent accessibility and involvement in H-bonds, and the time-dependent rate of exchange can be measured by a shift in mass of 1 Dalton per amide group with mass spectrometry. Regions of higher stability or burial will be characterized by more slowly exchanging amides compared with regions of low stability and exposure. Therefore, if a region of a protein is protected by a GAG, the reduced solvent accessibility should result in protection from exchange compared with the free protein. The experiment is conducted as follows: protein alone or protein and GAG is rapidly diluted into a D2O buffer for various amounts of time. At the end of a given exchange period, the sample is then ‘‘quenched’’ to low pH to terminate the reaction and prevent further exchange during subsequent steps. The samples are then subject to proteolysis with enzymes that work at low pH, like pepsin and fungal protease. One then determines the rate of incorporation of the deuterium into the peptides as a function of time with mass spectrometry. The resolution of the method is, therefore, defined by the peptide coverage, and the use of different proteases, therefore, increases resolution. This method was used to characterize the Link module from human tumor necrosis factor stimulated gene-6 (Link_TSG6) and hyaluronan (HA) oligosaccharides (Seyfried et al., 2007). Although promising, and widely used for protein-protein interactions, this study points out a major caveat. In this study, amides distal to the GAG-binding site were protected because of allosteric effects; thus one must be able to distinguish protection caused by conformational changes versus those caused by ligand binding. A general technical issue with the use of H/D exchange is to avoid back exchange of the deuterium off the protein during sample processing.

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Radiolytic oxidation, another relatively new mass spectrometry technique, does not suffer from this complication (Takamoto and Chance, 2006). This technique is also based on the principle of increased amino acid protection in the presence of ligand/protein/GAG, but in this case, the protection is from hydroxyl radicals that oxidize amino acid side chains. Although we know of only one report of the application to characterizing a carbohydrate binding protein (Charvatova et al., 2008), the method seems to have several advantages over HDMS. First, the modifications are covalent, which facilitates analysis without the worry of back exchange. Second, the oxidation seems to be directly related to solvent accessibility and thus unless major conformational arrangements occur, the results should readily reflect ligand binding epitopes without complications from allostery. All side chains are subject to oxidation, and the reactivities are reasonably well characterized; thus the method in principle has amino acid resolution. Finally, side chain interactions seem most important for binding GAGs, and these are probed in the radiolytic approach, whereas only backbone atoms are probed by HDMS, because the labile side chain atoms exchange much too fast for detection. At this point, the method has been applied to characterize the dimer interface of the carbohydrate-binding protein, galectin-1, but not yet with GAG bound. However, one can anticipate future applications to mapping protein-carbohydrate surfaces as the technique matures. One of the main issues will be the availability of software for the identification of the modifications that occur on hydroxyl radical oxidation, access to the appropriate mass spectrometry equipment, and further understanding of the underlying chemistry of oxidation.

4. Summary Interactions between chemokines and glycosaminoglycans have been proven to be critical for their biological function. Moreover, GAG-binding deficient mutants, engineered to test the relevance of GAG binding in vivo, have been shown to interfere with cell migration induced by the wild-type proteins and to be effective in animal models of disease, suggesting that such mutants could be effective protein therapeutics. These findings represent major advances in the chemokine field, yet there is much to be discovered. What really is the extent of the specificity encoded by interactions of these proteins with GAGs? For chemokines, these interactions could significantly influence their spatial localization on cells and thus function. To answer these questions, further development of methods beyond those outlined here will be necessary, and the chemokine field will likely benefit from emerging technologies involving glycan arrays, glycomics, sequencing by mass spectrometry, and methods for routine synthesis of carbohydrates and

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carbohydrate analogs. Beyond characterizing the interactions between chemokines and GAGs, which is largely the focus of the methods covered in this chapter, we also need to obtain a better understanding of the full significance of their functional roles in vivo.

ACKNOWLEDGMENTS This work was funded by an NRSA postdoctoral fellowship F32GM083463 awarded to D. J. H., and by the Lymphoma Research Foundation, the Department of Defense (USAMRAA W81XWH0710446), and NIH (RO1-AI37113 and R21AI076961) awards to T. M. H. and a European Union FP6 INNOCHEM award (LSHB-CT-2005-518167) to A. E. I. Proudfoot.

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C H A P T E R

F I V E

Multiple Approaches to the Study of Chemokine Receptor Homo- and Heterodimerization Jose´ Miguel Rodrı´guez-Frade, Laura Martinez Mun˜oz, and Mario Mellado Contents 1. Introduction 2. Biochemical Techniques to Measure Chemokine Receptor Oligomerization 2.1. Western blot and immunoprecipitation 2.2. Colocalization assays 2.3. Fluorescence labeling of antibodies 2.4. Construction of fluorescence-labeled receptors 3. Resonance Energy Transfer (RET) Techniques 3.1. Bioluminiscence resonance energy transfer (BRET) techniques 3.2. Fluorescent resonance energy transfer (FRET) techniques 4. Sequential BRET-FRET (SRET) Technology 5. Conclusion Acknowledgments References

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Abstract Chemokines belong to a family of structurally related chemoattractant proteins that bind to specific seven-transmembrane receptors linked to G proteins. They are implicated in a variety of biologic responses ranging from cell polarization, movement, immune and inflammatory responses, as well as prevention of HIV-1 infection and cancer metastasis. Recent evidence indicates that chemokine receptors can adopt several conformations at the cell membrane. Chemokine receptor homo- and heterodimers preexist on the cell surface, even in the absence of ligands. Chemokine binding stabilizes specific receptor conformations and activates distinct signaling cascades. Analysis of the conformations

Department of Immunology and Oncology, Centro Nacional de Biotecnologı´a/CSIC, Madrid, Spain Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05405-6

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adopted by the receptors at the membrane and their dynamics is crucial for a complete understanding of the function of these inflammatory mediators. We focus here on conventional biochemical and genetic methods, as well as on new imaging techniques such as those based on resonance energy transfer, discussing their advantages, disadvantages, and possible complementarity in the analysis of chemokine receptor dimerization.

1. Introduction The family of low molecular weight proinflammatory cytokines termed chemokines were originally described as specific mediators of leukocyte directional movement (Mackay, 2001). Current views nonetheless implicate these molecules in the movement of several cell types, because they participate in functions such as lymphocyte trafficking (Baggiolini, 1998), regulation of T cell differentiation (Sallusto et al., 1998), HIV-1 infection (Berger et al., 1999), angiogenesis (Belpario et al., 2000), development (Raz, 2003), and tumor metastasis (Mu¨ller et al., 2001). Today, scientists refer to the nearly 50 known chemokines as either constitutive chemokines, which are usually regulated during development, or as inducible chemokines, whose expression is regulated mainly by inflammatory mediators (Proudfoot, 2002). In addition, certain viruses encode highly selective chemokine receptor ligands that can serve as agonists or antagonists and may, thus, have a role in viral dissemination or evasion of host immune response (Alcami, 2003). On the basis of their broad range of functions, it is easy to deduce that chemokines must be central to a variety of diseases that are characterized by inflammation and cell infiltration. They have become a major focus of interest as therapeutic targets, because there is a clear correlation between the expression of specific chemokines and the orchestrated recruitment of cell populations during the course of some disease processes (Proudfoot, 2002). The chemokines act by binding to class A rhodopsin–like, seventransmembrane G-protein–coupled receptors (GPCR) (Horuk, 2001). The 20 receptors characterized to date are classified as CCR, CXCR, CX3CR, and XCR on the basis of their ligand specificity (Rossi and Zlotnik, 2000). Another group of receptors (D6, DARC, and CCXCKR) that can interact with several chemokines were recently denominated ‘‘silent’’ receptors, because they are unable to activate signal transduction events that lead to cell chemoattraction (Borroni et al., 2008). Most chemokine receptors are able to interact with more than one chemokine (shared receptors), although there are some examples of specific chemokine-receptor pairs (specific receptors) (Horuk, 2001). Expression of these receptors is finely regulated by factors that include cytokines,

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growth factors, and cell cycle status (Loetscher et al., 1996; Papadopoulus et al., 1999; Parks et al., 1998). It is, therefore, not surprising that cells respond differently to a chemokine, depending on the microenvironment in which they are found. As the chemokine receptors integrate numerous signaling pathways (Soriano et al., 2003; Thelen and Stein, 2008), the chemokine-mediated signaling cascade is more complex than was originally thought. Although initially considered a cytokine habit (Thelen and Baggiolini, 2001), various studies have demonstrated the existence of chemokine receptor homo- and heterodimers and have speculated on the functional relevance of these conformations (Mellado et al., 2001a,b; Percherancier et al., 2005; Vila-Coro et al., 2000; Wang et al., 2006). The difficulties in detecting these complexes with immunoprecipitation methods suggest conformational instability in the absence of ligand, but resonance energy techniques clearly show that chemokine receptor dimers form spontaneously in the absence of ligand (Hernanz-Falco´n et al., 2004; Wilson et al., 2005). A number of questions remain to be answered, however, including the dynamic nature of these receptor complexes and the role of distinct ligands in promoting the conformational changes that trigger function. This is particularly important in the case of chemokines, because there is a relative lack of selectivity in ligand binding, with many receptors showing high affinity for more than one chemokine (Tian et al., 2004) and simultaneous expression of several receptors on the same cell. Although biochemical technologies were classically used to analyze protein-protein interactions, our current knowledge has been supplemented by new approaches on the basis of energy transfer between fluorochromes followed by confocal microscopy. These methods exploit important technologic advances such as laser light sources, fluorescent probes, and improvements in computer science that allow digital imaging and image analysis. Independently of the technique used, the cell system being used must be characterized in detail before attempting analysis of chemokine receptor dimerization. This includes routine testing such as analysis of cell cycle status, cell surface receptor expression, and determination of receptor number and affinity constants, especially when cells are transfected with mutant or fluorescently labeled receptors.

2. Biochemical Techniques to Measure Chemokine Receptor Oligomerization Until recently, most assays used to demonstrate receptor oligomerization were based on biochemical approaches such as immunoprecipitation or crosslinking. Alternately, dominant negative receptor mutants were used to

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abrogate wild-type receptor functions by forming nonfunctional complexes (Rodriguez-Frade et al., 1999). Because any single method alone has intrinsic limitations, most should be used in concert with others to obtain meaningful results.

2.1. Western blot and immunoprecipitation Although the general protocol for immunoprecipitation assays is very similar, lysis buffer compositions vary, and cell number should be adjusted for each case. Cells (1
107 cells/ml), unstimulated or stimulated with the appropriate chemokine, are diluted immediately with 1 ml cold PBS to terminate stimulation. Cells are centrifuged (800g, 5 min, 4  C), washed with cold PBS, resuspended in 200 ml lysis buffer (10 mM triethanolamine, pH 8, 150 mM NaCl, 1 nM EDTA, 10% glycerol, 2% digitonin), and incubated (30 min, 4  C, with continuous rocking). Other lysis buffers can be also used; however, detergents can alter receptor interactions and must be evaluated carefully. A preclearing step is essential to reduce nonspecific binding to the immunoprecipitating antibody (Ab). For this step, incubate the supernatant containing solubilized proteins with antiimmunoglobulin Ab (antibody against immunoglobulin of the animal species from which the immunoprecipitating Ab is derived) coupled to agarose and incubate (30 min, 4  C). Then add the immunoprecipitating Ab (90 min, 4  C), followed by anti-Ig coupled to agarose (without washing; 60 min, 4  C). After extensive washing and centrifugation, resolve the pellets by SDS-PAGE. To adjust the percentage of the acrylamide solution, remember that the predicted molecular weight of chemokine receptors is in the 30- to 50-kDa range. Transfer the gel to nitrocellulose membranes and develop Western blot as described elsewhere (Rodriguez-Frade et al., 1999). The immunoprecipitation technique is based on the use of chemokine receptor-specific Ab and depends greatly on their characteristics (specificity, affinity). This method is useful for evaluation of receptor heterodimers, in which case one receptor should appear in the immunoprecipitates of the other receptor. Alternately, it can be applied to analyze homodimerization, although in this case the receptors must be tagged appropriately. Receptor labeling bypasses the need to raise antibodies specific for target receptors and has been used successfully to demonstrate homodimerization of CCR2 and in the case of other GPCR, such as b2Ars, GABAB, mGluR5, d-opioid, m3-muscarinic, and Ca2þ receptors (Angers et al., 2002; Rodriguez-Frade et al., 1999). Selection of an appropriate tag and its location in the receptor are both important factors. Amino acids added to the extracellular region might alter ligand binding, whereas modification of intracellular domains can disturb the coupling of signaling molecules. Both ligand affinity and receptor distribution should first be evaluated to ensure that the behavior of the tagged receptor is similar to that of the wild-type receptor. FLAG, myc,

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HA, GST, His, GFP, and GAL are among the tags for which there are commercially available antibodies that immunoprecipitate and recognize the tagged receptor in Western blot. Homemade epitopes can also be designed and included into the receptor cDNA sequence with standard PCR techniques (Rodriguez-Frade et al., 1999). Immunoprecipitation methods can be complemented with crosslinking assays with bifunctional reagents such as disuccinimidyl suberate (DSS). Before lysis, cells should be resuspended in 1 ml cold PBS with 10 ml 100 mM DSS and incubated (10 min, 4  C). Care must be taken in cell handling before the lysis step, because the presence of nonintact cells increases the background of nonspecific protein crosslinking. Prepare the DSS reagent just before use. Continue with lysis, immunoprecipitation, and Western blot analysis as described previously. In this case, Western blot analysis with specific Ab will develop the band corresponding to the monomeric receptor, as well as the high molecular weight dimeric, trimeric and oligomeric species.

2.2. Colocalization assays Modern optical microscopy allows us not only to visualize organelles and molecules but also to study their function. In living cells, we can analyze how a molecule moves, changes location, or associates with other molecules. Such phenomena were originally evaluated with colocalization assays, which detect light from two different fluorophores and evaluate a digital image for the presence of the same pixel in two distinct channels. Signal colocalization indicates adjacency of fluorophores, and thus of the molecules they label (Fig. 5.1). A high-numerical aperture microscope lens permits resolution near 300 nm, sufficient to locate molecules in different cell compartments but not to demonstrate molecular association. The technique requires fluorescence-labeled antibodies or receptors coupled to fluorescent proteins. To determine colocalization between chemokine receptors, plate cells on coverslips coated with poly-L-lysine (20 mg/ml, 1 h, 37  C) and culture them (24 h, 37  C, 5% CO2). After washing, fix the cells with 4% paraformaldehyde (10 min, room temperature [RT]). To avoid nonspecific binding, treat the cells with PBS supplemented with 1% BSA, 0.1% goat serum and 50 mM NaCl (1 h, 37  C). Add the receptor-specific Ab (30 min, RT). To facilitate the procedure, use antibodies of distinct species origin (i.e., mouse, rabbit, rat, hamster) to stain the two receptors. If prelabeled Abs are available, their use precludes the need for secondary antibodies. Otherwise, add the mixture of prelabeled secondary Ab (20 min, RT). If both primary Abs are of the same origin and isotype, add one of these Abs, followed by its fluorochrome-labeled secondary antibody. Wash and incubate the cells with undiluted serum from the same species as the primary Ab (30 min, RT). Now stain the cells with the specific Ab for the second receptor as

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A Cy

3

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Figure 5.1 Chemokine receptors colocalize at the cell membrane.(A) Scheme of the colocalization experiment. Chemokine receptors can be fused to fluorescent proteins (left) or immunostained with fluorochrome-labeled specific antibodies (right). (B) In a representative experiment, CCR2/CCR5 stably transfected L1.2 cells were fixed and costained with anti-CCR2 (anti-CCR2-Cy3) and -CCR5 mAb (anti-CCR5-Cy2).The merged image is also shown; arrows indicate colocalization areas (yellow). All images are overlaid on the DIC (differential interference contrast) image.

previously, followed by its secondary antibody labeled with a different fluorochrome (20 min, RT). Evaluate fluorescence on a confocal microscope with filters appropriate for the fluorochromes used.

2.3. Fluorescence labeling of antibodies Although commercially available Ab can be used for these purposes, antibodies can also be labeled in the laboratory. Given their intense fluorescence and low hydrophobicity, the Cy dyes are efficient tags for fluorescence labeling. In the standard labeling procedure, the contents of a commercial vial (‘‘to label 1 mg of protein’’) of Cy2, Cy3, or Cy5 are dissolved in 50 ml dimethylsulfoxide (DMSO). The antibody is dissolved to 1 mg/ml in buffer (100 mM NaCl and 35 mM H3BO3, pH 8.3). Mix 10 ml of dye/DMSO mixture with 200 ml antibody solution and incubate (30 min, 25  C, in the dark). Separate unbound dye by adding 300 ml of 100 mM NaH2PO4, incubate (30 min, 25  C), and load the sample on a PD-10 column preequilibrated with 100 mM NaCl, 50 mM NaH2PO4, 1 mM EDTA

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(pH 7.5). Elute the labeled protein with 2 ml of distilled H2O. The labeled antibody should be titrated against the chemokine receptor before use.

2.4. Construction of fluorescence-labeled receptors For rapid, easy visualization of chemokine receptors, fluorescent proteinbased constructs and microscopy techniques are very helpful. The constructs are typically used in colocalization analysis and energy transfer techniques. Receptors are cloned with standard molecular biology methods into commercially available vectors bearing fluorescent proteins. Insertion of the fluorescent probe in the C-terminal region of the receptor involves elimination of the receptor stop codon, whereas insertion in the N-terminal region requires elimination of the fluorescent protein stop codon. Transfected cells should be analyzed for receptor expression and function.

3. Resonance Energy Transfer (RET) Techniques Newer methods to determine chemokine receptor oligomerization are based on resonance energy transfer (RET). These techniques are also useful for determining conformation dynamics, the role of ligand and receptor levels, and for defining the dimerization site within the cell (Harrison and van der Graaf, 2006). There are two main types of RET, bioluminescence resonance energy transfer (BRET) and fluorescence resonance energy transfer (FRET). In the former, the donor molecule is luminescent (Pfleger and Eidne, 2006); in the latter, the donor fluorochrome transfers energy to an acceptor fluorochromes (Cardullo, 2007). Both techniques require generation of fusion proteins between the receptor and the fluorescent/luminescent donor and acceptor proteins, as well as the use of transfected cells (Boute et al., 2002). Controls must, therefore, be included to rule out alterations in receptor distribution between the cells and to avoid differences in receptor-mediated cell function. Although BRET has been used at the single cell level (Coulon et al., 2008), it is, in fact, an approach for cell suspensions (Pfleger et al., 2006). It allows measurement of energy transfer between receptors independently of their expression pattern and permits quantitation. In contrast, FRET imaging techniques use confocal or wide-field microscopy, allowing measurements in single cells and identification of cell locations at which FRET is detected.

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3.1. Bioluminiscence resonance energy transfer (BRET) techniques BRET makes use of nonradiative energy transfer between a light donor and a fluorescent acceptor. The bioluminescent energy resulting from the catalytic degradation of a substrate by luciferase is transferred to an acceptor fluorophore, which in turn emits a fluorescent signal (McVey et al., 2001). This transfer takes place when there is an effective range of 10 nm between donor and acceptor, which allows at least 50% of the energy to excite the acceptor molecule. For maximum spectral overlap, the acceptor fluorophore (YFP or GFP2) varies depending on the substrate oxidized (coelenterazine or its derivative DeepBlueC, respectively). Because of their small size and their hydrophobicity, coelenterazines cross the cell membrane easily. For BRET measurements, at 48 h posttransfection, wash cells once with PBS. Add coelenterazine H (Nanolight Technology) to a final concentration of 5 mM in PBS, and take readings with a multidetector plate reader that allows the sequential integration of signals detected in the 480  20 nm and 530  20 nm windows for luciferase and YFP light emission, respectively. The BRET signal is determined by calculating the ratio of the light intensity emitted by receptor-YFP to the light intensity emitted by receptor-RLuc. The values are corrected by subtracting the background BRET signal as measured in the same cells expressing the receptor-RLuc or the receptorYFP construct alone. For acquisition of full BRET spectra, cells are transfected with different amounts of receptor-YFP for a given quantity of receptor-RLuc. Cells are detached and resuspended in HBSS containing 0.1% (w/v) glucose. Cells (2
105) expressing different acceptor/donor (YFP/RLuc) ratios are seeded in 100 ml HBSS in a clear-bottom 96-well plate, and a BRET scan is performed by reading luminescence between 400 and 600 nm, immediately after coelenterazine addition. YFP fluorescence is determined in the same cells with a black 96-well plate. For BRET titration experiments, net BRET ratios are expressed as a function of the acceptor/donor ratio. These BRET saturation curves provide an idea of the maximum BRET signal. They also allow evaluation of BRET50, which is proposed to indicate the ability of two partners to interact (Audet et al., 2008); nonetheless, this would only be the case if the association between the receptors is reversible, which has not yet been demonstrated. Total fluorescence and luminescence are used as relative measures of total acceptor and donor protein expression, respectively. Total fluorescence is determined with an excitation filter at 485 nm and an emission filter at 535 nm. Total luminescence is measured 5 to 10 min after coelenterazine addition, in the absence of the emission filter. Because BRET-based experiments do not permit subcellular analysis, results can be altered, for example, by random collisions because of

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accumulation of chemokine receptors in intracellular membranes. To define receptor oligomers, experiments must be performed with several acceptor/donor ratios at a fixed surface density and a number of expression levels at a defined acceptor/donor ratio. Under these conditions, nonspecific interactions are independent of BRET efficiency and of acceptor/ donor ratios. Finally, it is very useful to include positive controls such as the donor genetically fused to the acceptor and, when possible, a known interacting receptor pair whose characteristics resemble those studied. Although the ideal negative control is a noninteracting protein similar to that analyzed, the acceptor protein is normally used alone.

3.2. Fluorescent resonance energy transfer (FRET) techniques In FRET, donor excitation energy is transferred to the acceptor by means of an induced dipole-dipole interaction; efficiency depends on the distance between and orientation of donor and acceptor fluorophores (Sekar and Periasami, 2003). Ideal dyes are photostable, have little intensity fluctuation, and are relatively small in size to minimize perturbation of the chemokine receptor. For all FRET methods, donor/acceptor choice is critical. Donor emission spectra should ideally have maximum overlap with acceptor absorption spectra, although acceptor and donor emissions should be clearly separable to minimize background interference. Fluorochrome incorporation into the protein must also be considered, because FRET sometimes requires the generation of chimeric constructs in which each chemokine receptor is fused to a different, modified form of green fluorescent protein (GFP). Some donor/acceptor combinations are blue (BFP)/GFP, CFP/YFP, green (GFP)/dsREd, GFP2/YFP, and YFP/dsRed. The most frequently used is, nonetheless, the CFP/YFP combination, because both are extremely bright, and this combination offers few technical problems (Pollok and Heim, 1999). BFP is a poor donor, because it is not especially bright, making FRET between BFP and GFP difficult to detect. dsRed is a poor acceptor, because it has a broad absorption spectrum and excites the same wavelength as the donor (GFP or YFP). FRET is sometimes evaluated on intact receptors with specific Ab conjugated to appropriately selected fluorescent dyes. Some common dyes are Alexa488, FITC, Cy3 or Cy2 as donor, and rhodamine-2, Alexa555, Cy3, or Cy5 as acceptor. Although Cy3 (donor) and Cy5 (acceptor) form a suitable fluorochrome pair, the Cy2 donor/Cy3 acceptor pair is often more convenient, because it allows use of the widely available 488-nm argon laser line. Secondary antibodies are occasionally needed, although the increased distance between fluorophores complicates FRET detection; this can be

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resolved with dye-labeled F(ab0 ) fragments. Because FRET depends on fluorochrome distance and orientation, low FRET efficiency values do not necessarily correlate with lack of dimerization. Several methods are used to determine and quantify FRET. The first is sensitized acceptor fluorescence, in which the donor fluorescent dye is excited and the acceptor signal is measured (Fig. 5.2). Another possibility is acceptor photobleaching, a method based on quenching donor fluorescence. Some donor photons are used to excite the acceptor, decreasing the emission energy detected. Photobleaching of the acceptor abolishes FRET, increasing donor light emission (Fig. 5.3). This method cannot be used for living cells, however, because exposure to extended laser energy is thought to damage the cell. Finally, fluorescence lifetime imaging microscopy (FLIM) measures a chromophore’s fluorescence lifetime, allowing spatial resolution of biochemical processes. The fluorescence lifetime of a donor dye decreases under FRET conditions, independently of fluorophore concentration (Periasami et al., 2002). To develop FRET assays by photobleaching with the CFP/YFP pair, plate HEK-293T cells (3.5
104 cells/ml) on poly-L-lysine–coated coverslips (20 mg/ml, 1 h, 37  C) and incubate (24 h, 37  C, 5% CO2). Cotransfect receptor combinations at a 1:1 ratio (one YFP- and one CFP-labeled receptor) and incubate (48 h, 37  C, 5% CO2); confirm equal levels of each receptor at the cell surface with standard flow cytometry analysis. Wash cells in PBS and fix with 4% formaldehyde (4 min, RT). Wash in A YFP emission

YFP

CFP

CFP emission B CFP

YFP

FRET

Figure 5.2 Analysis of chemokine receptor dimerization by the sensitized acceptor fluorescence method for FRET. (A) Scheme illustrating the sensitized acceptor fluorescence method. CFP is excited with a 405-nm laser line and YFP emission detected at 530 nm. Alternately, a decrease in CFP emission can be detected at 460 to 500 nm. (B) Unstimulated HEK-293Tcells were transiently cotransfected with CCR2-CFP and CCR5-YFP, fixed, and FRETdetermined by detection of YFP emission. Images show CFP staining (left),YFP staining (middle), and FRET (right).

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Figure 5.3 Analysis of chemokine receptor dimerization by the photobleaching method in FRET. (A) Schematic representation of the photobleaching method. Some donor energy is used to excite the acceptor, decreasing detectable donor emission (left). Photobleaching of the acceptor abolishes FRET, increasing donor emission (right). (B) FRET evaluation of chemokine receptor heterodimerization with the acceptor photobleaching method. Unstimulated HEK293T cells were transiently cotransfected with a 1:1 combination of CCR2-CFP and CCR5-YFP, fixed, and FRET was determined. The image shows CFP staining before (CFP-pre) and after (CFP-post) photobleaching, as well as a false color merged image (FRET) and a zoom image of FRETat the photobleached area (insets).The DIC image is also included (left).

PBS and mount coverslips onto slides with PBS (pH 7.0) containing 80% glycerol. Evaluate fluorescence on a confocal microscope with appropriate filters for the fluorochrome. In a FRET assay, an image of the cell region of interest is taken with standard spectroscopic settings. CFP and YFP are excited by separate sweeps of the 405-nm (laser diodo [25 mW]) and 515-nm lines (three-line argon laser [45 mW]), respectively, and directed to the cell by means of a 405- to 440/515-nm dual dichroic mirror. The emitted fluorescence is split by a 510-nm dichroic mirror for CFP and directed to a spectral detector adjusted to the 460- to 500-nm range. For YFP, fluorescence is directed to a spectral detector adjusted to the 530- to 570-nm range. Confocal fluorescence intensity data (ICFPpre and IYFPpre) are recorded, with a pinhole of 100, as the average of four line scans per pixel and digitized at 12 bits. Repeated scans with 515 nm maximum light intensity are used to photobleach YFP, which requires 5 to 30 sec at maximal scan rates and a 100-pinhole aperture. After 60 to 90% of YFP bleaching, fluorescence intensity (ICFPpost and IYFPpost) is measured with identical parameters. With ImageJ 1.40g software (NIH), FRET efficiency is determined on a pixel-by-pixel basis (E) and calculated in percent as E = (1  FDpre/FDpost)
100%, where FDpre and FDpost are the background-corrected CFP fluorescence intensities before

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and after YFP photobleaching, respectively (Kenworthy, 2001; Zimmermann, 2002). As a negative control, FRET should be determined in transiently transfected HEK-293T cells with a chemokine receptor-CFP and in a region of the cotransfected cells without photobleaching. FRET efficiency should be calculated from several independent determinations, with at least 50 images from each. Recall that FRET provides meaningful data when determined with cells that have been evaluated individually for similar CFP and YFP fluorescence intensities. An alternative based on the sensitized acceptor fluorescence method allows FRET evaluation in cell populations (Carriba et al., 2008). HEK-293T cells are transiently transfected with vectors encoding the chemokine receptors of interest, coupled to CFP or GFP2 (donor) and YFP (acceptor). At 48 h posttransfection, distribute cell suspensions (20 mg protein/well) into 96-well microplates and read them in a fluorimeter equipped with a highenergy xenon flash lamp, with a 10-nm bandwidth excitation filter at 430 nm (CFP) or 400 nm (GFP2) and 10-nm bandwidth emission filters for 495 nm (CFP), 510 nm (GFP2), and 530 to 535 nm (YFP). To avoid spectral mixing, use identical gain settings for all experiments to maintain a constant relative contribution of the fluorophores to the detection channels. It is critical to measure the contribution of donor and acceptor alone to each detection channel in experiments with cells expressing only one of the proteins; these values are normalized to the sum of the signal obtained in the two detection channels. To exclude an effect on FRET efficiency because of the acceptor/ donor ratio and to be able to compare dimerization efficiency between receptor pairs, generate a FRET curve with constant expression levels of the receptor coupled to the donor and increasing amounts of the receptor coupled to the acceptor. This curve yields the FRETmax, which is not informative of interaction specificity, as well as the FRET50 (the acceptor/donor value at half-maximal FRET), which indicates the propensity of the interacting partners to associate and thus reflects differences in the relative affinity of these two partners. FRET efficiency is determined as previously (Zimmermann, 2002). FRET-based experiments can also be designed to determine the dynamics of receptor conformation at the cell membrane (Pello et al., 2008). If the role of the ligand is being studied, cells should be stimulated before they are fixed. The influence of a given receptor, R1, on R2:R2 homodimers can be determined by measuring photobleaching FRET in HEK-293T cells transiently cotransfected with CFP-R2 and YFP-R2 and comparing the result with FRET in HEK-293T cells transiently cotransfected with these two receptors plus R1. To facilitate analysis, R1 is expressed in a pIRES2AcGFP1-Nuc vector (Clontech). Only R1-expressing cells will be GFP-labeled in the nucleus and will be used to determine FRET. These experiments can also be modified to measure FRET between receptor heterodimers. In all cases, R1 expression in GFPþ cells should be controlled by flow cytometry and the CFP/YFP ratio determined by separate

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measurement of fluorescence levels. Controls should also include evaluation of the effect of R1 on R2 expression. For a complete analysis, the effect of the unlabeled receptor on FRET saturation curves assays should be measured; determine sensitized acceptor FRET in cells coexpressing a constant donor amount and increasing acceptor levels alone or in the presence of the unlabeled specific or unspecific receptors. FLIM is the most powerful FRET technique, because it is independent of transfection levels, although it requires complex equipment. In a typical FLIM determination, cells are plated on a chamber coverglass and transfected as described previously. Avoid fixing to allow in vivo measurements. FLIM is measured with a confocal microscope with a High Speed Lifetime Module and a 60
PlanApo 1.4 objective, or equivalent equipment. Fluorescence lifetime is determined after excitation with a pulsed laser (picosecond pulses) and a bandpass emission filter appropriate for the fluorochrome used and is quantitated with LIMO (Nikon) or similar software. Avoid the use of mounting solutions, which increase autofluorescence. The fluorescence lifetime of a donor is a constant parameter in specific experimental conditions (Fig. 5.4). A reduction in donor lifetime is due to a A

Pulsed laser

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Figure 5.4 Analysis of chemokine receptor dimerization by FLIM. (A) Scheme showing the FLIM method. The fluorescence lifetime of CFP after pulsed laser excitation (440 nm) (left) decreases under FRETconditions (right). (B) CFP fluorescence lifetime images (calculated from the phase shift) of HEK-293 cells expressing CCR5-CFP (left) or CCR5-CFP/CCR2-YFP (right). The pseudocolor scale ranges from 0 (black) to 4.0 nanosec (white).

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quenching effect triggered by the acceptor and is, therefore, indicative of chemokine receptor dimerization. Positive and negative controls as in BRET or in other FRET methods should also be included. Pay careful attention to variations in pH, temperature, and ionic strength in the medium, because these parameters cause alterations in FLIM measurements. Mycoplasma infection of the cells will also cause artifacts.

4. Sequential BRET-FRET (SRET) Technology Although BRET and FRET techniques are widely used to demonstrate homo- and heterodimers in living cells, they are inadequate for evaluating high-order complexes; that is, complexes involving more than two molecules. A new BRET-FRET–based technique called sequential BRET-FRET (SRET) was recently described (Carriba et al., 2008). SRET uses cells expressing a protein fused to RLuc, a protein fused to a BRET acceptor (GFP2 or YFP), and a protein fused to a FRET acceptor (YFP or DsRed). Addition of a RLuc substrate promotes acceptor excitation by BRET and subsequent energy transfer to the FRET acceptor (Fig. 5.5). We use suspensions of transiently cotransfected HEK-293T cells (20 mg protein/well) distributed in 96-well microplates, read in a fluorimeter equipped with a high-energy xenon flash lamp and a series of 10-nm bandwidth excitation and emission filters appropriate for the receptor-fused protein; this allows detection of BRET and FRET acceptor emission. These experiments require quantitation of receptor-fluorochrome expression, separation of the relative contribution of each fluorophore to the detection channels, quantitation of receptor-RLuc expression by determining luminescence, and, finally, SRET determination after addition of the RLuc substrate. The exact proportion of each fluorophore

RLuc 400 nm BRET

dsRED

YFP 530 nm FRET

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0n

m

Colenterazine

Figure 5.5 Sequential resonance energy transfer technique (SRET). Cell coexpressing chemokine receptors, each fused to RLuc, to YFP, or to dsRED. Addition of the RLuc substrate (coelenterazine) triggers luciferase light emission at 485 nm. In consequence, the excited donor FRET (YFP) emits at 530 nm, which excites the FRET acceptor (dsRED). dsRED emission is then detected at 590 nm. This technique is used to study oligomerization.

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in the cell population is thus established, which indicates the receptor ratio used. These experiments measure FRET in cell populations, as for BRET; analyses, therefore, cannot be restricted to a specific cell area. All the general considerations and controls described for BRET and FRET are also applicable for these assays.

5. Conclusion Chemokines are the principal chemotactic factors implicated in the regulation of leukocyte traffic, both to inflammation sites and for establishing lymphoid organ architecture. Chemokines mediate their function by interacting with specific members of the seven-transmembrane, G-protein–coupled receptor family, which are expressed on the cell surface. Much information is available on the biochemical pathways activated by this large receptor family. Recent experiments, including those based on the application of resonance energy transfer (RET) technology, have, nonetheless, revealed an unexpected degree of complexity in chemokine receptor dynamics at the plasma membrane. In addition to the known promiscuity between ligands and receptors, the chemokine receptors can adopt a variety of conformations at the cell surface. These homo- and heterodimeric, and possibly oligomeric conformations, might be modulated by the levels of chemokine receptors or of other GPCR, as well as by chemokine expression. To explain the precise role of ligands and receptors in these dynamics, the classical biochemical techniques such as crosslinking, immunoprecipitation, and Western blot must be complemented by new technologies such as those based on RET and microscopy analysis. For these procedures, methodologic questions will need to be clarified, including use of different cell types, protein overexpression, and use of chemical inhibitors that can alter in vitro distribution, availability, and/or function of chemokine receptors compared with their in vivo behavior. Correctly used, these techniques will clearly be of value in unraveling the complexities of chemokines, their receptors, and their signals. Improved understanding of receptor homoand heterodimerization is changing our view of chemokine receptor structure and activation, which is likely to have substantial influence on drug development and screening.

ACKNOWLEDGMENTS We thank the members of the DIO chemokine group, who contributed to some of the work described in this review. We also thank C. Bastos and C. Mark for secretarial support and helpful editorial assistance, respectively. This work was partially funded by grants from the EU (Innochem LSHB-CT-2005-518167 and Molecular Imaging LSHG-CT-2003-503259),

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the Spanish Ministry of Science and Innovation (SAF2005-03388), and the Madrid Regional Government. The Department of Immunology and Oncology was founded and is supported by the Spanish National Research Council (CSIC) and by Pfizer.

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C H A P T E R

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Plasmon Resonance Methods in Membrane Protein Biology: Applications to GPCR Signaling Zdzislaw Salamon,* Gordon Tollin,*,† Isabel Alves,‡ and Victor Hruby*,† Contents 124 125 125 126 129 131 132 136 138 141 144 144 144

1. Introduction 2. Plasmon Spectroscopy 2.1. Description of surface plasmons 2.2. Varieties of surface plasmon resonances 3. Sensor Construction and Sample Deposition 4. Lipid Bilayer Deposition 5. Spectral Data Analysis 6. Membrane Protein Insertion 7. Ligand and G-Protein Binding by GPCRs 8. Conclusions Conflicts of Interests Acknowledgments References

Abstract Plasmon waveguide resonance (PWR) spectroscopy, a variant of surface plasmon resonance (SPR) spectrometry, allows one to examine changes in conformation of anisotropic structures such as membranes and membrane-associated proteins such as G-protein–coupled receptors (GPCRs). The binding and resulting structural changes that accompany interactions of membrane protein with ligands (agonists, antagonists, inverse agonist, etc.), G-proteins, and other effectors and modulators of signaling can be directly examined with this technique. In this chapter we outline the instrumentation used for these studies, the experimental methods that allow determination of the structural changes, and thermodynamic and kinetic parameters that can be obtained from these studies.

* { {

Department of Biochemistry and Molecular Biophysics, University of Arizona, Tucson, Arizona, USA Department of Chemistry, University of Arizona, Tucson, Arizona, USA Department of Chemistry, Universite Pierre et Marie Curie, Paris, France

Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05406-8

#

2009 Elsevier Inc. All rights reserved.

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1. Introduction Examination of the structures of integral membrane proteins and of the structural changes that accompany their interactions with biologically important ligands such as agonists, antagonists, inverse agonists, partial agonists, allosteric modulators, and other integral membrane proteins has been very difficult. This is due to their relatively low abundance in membranes, the highly anisotropic properties of cellular membranes, the relative lack of biophysical and analytical methods that can directly probe integral membrane protein structure, and the difficulty of purifying and crystallizing integral membrane proteins in their biologically active forms. The use of radioactive and fluorescent probes on the ligands for integral membrane protein, fluorescent, and other physical probes on the integral membrane proteins and on the membranes has allowed the development of binding assays (thermodynamic and kinetic information can be obtained) and the evaluation of certain intermolecular interactions at the membrane surface in response to ligands, but there often are difficulties of interpretation and of evaluating artifacts because of the structural modifications that have occurred by addition of the probes and reporter groups, fluorescent proteins, dyes, etc. Clearly there is a need for better biophysical methods that can directly probe these biologic systems. In this chapter, we will discuss a relatively new biophysical method, plasmon-waveguide resonance (PWR) spectroscopy (Salamon and Tollin, 1999a,b; Salamon et al., 1997a,b,c, 1999a) that allows one to directly examine structural changes of integral membrane proteins that result from their interactions with ligands or other proteins on the membrane surface. Anisotropic properties can be directly probed, as can thermodynamic and kinetic properties that result from these interactions, without the need for any chemical modifications. In this chapter, we will illustrate the uses and power of this new method with G-protein–coupled receptors (GPCRs) as an example of an integral membrane protein. GPCRs are the largest class of integral membrane proteins, in fact the largest class of proteins in the human genome with more than 1000 different proteins now known. They are absolutely essential for most aspects of intercellular communication in complex biologic multicellular systems such as human beings and are the targets of most hormones and neurotransmitters that modulate behavior and metabolism. Furthermore, they are directly involved in many diseases. In fact, they are the targets of approximately 50% of all current drugs. In this chapter, we will examine the use of PWR to monitor and provide thermodynamic and kinetic information in regard to ligand binding, information transduction by G-proteins, GTP-GDP exchange occurring as a consequence of receptor activation, effects of membrane composition, and other aspects of GPCR action that can be directly examined by PWR spectroscopy.

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2. Plasmon Spectroscopy 2.1. Description of surface plasmons The concept of surface plasmons (Salamon and Tollin, 1999a; Salamon et al., 1997b) originates from the plasma formulation of Maxwell’s theory of electromagnetism, where the free electrons of a metal are treated as a high-density liquid (plasma). Plasma oscillations in metals are collective longitudinal excitations of the electrons, and plasmons are the quanta representing these charge-density oscillations. The spreading electron density fluctuations generate a surface-localized electromagnetic wave that propagates along the plane interface between the metal and an adjacent dielectric medium (e.g., air or water), with the electric field normal to this interface and decreasing exponentially with distance from both sides of the interface. These characteristics of the electromagnetic field also describe the guided surface waves (also known as evanescent waves) generated optically under total internal reflection conditions when all of the incident light is reflected at the boundary of the incident and emerging media. Surface plasmon excitation is a resonance phenomenon that occurs when energy and momentum conditions between incident light photons and surface plasmons are matched according to the following equation (Salamon et al., 1999a): 1=2

kSP ¼ kph ¼ ðo=cÞe0 sin a0 ;

ð6:1aÞ

kSP ¼ ðo=cÞðe1 e2 =e1 þ e2 Þ1=2 :

ð6:1bÞ

where

kSP is the longitudinal component of the surface plasmon wave vector, kph is the component of the exciting light wave vector parallel to the active (metal) medium surface, o is the frequency of the surface plasmon excitation wavelength (l), c is the velocity of light in vacuo, e0, e1, and e2 are the complex dielectric constants for the incident, surface active and dielectric (or emerging) media, respectively, and a0 is the incident coupling (resonance) angle. To satisfy this relationship in a conventional resonator assembly (Salamon and Tollin, 1999b; Salamon et al., 1997c), plasmon excitation must occur through an evanescent wave generated by p-polarized incident light (electric vector perpendicular to the surface). This plasmon excitation geometry turns out to be ideal for application as a biomedical sensor, because the excitation light is totally separated from the emergent medium

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(e.g., a biologic fluid to be analyzed). This allows the assay of such fluids regardless of their optical transparency (including, for example, blood, urine, cell suspensions, etc.). The resonance conditions mentioned previously can be fulfilled by changing the incident light angle, a, at a constant value of photon energy (i.e., wavelength) (Salamon and Tollin, 1999a; Salamon et al., 1997b). At resonance, the surface plasmons are generated at the expense of the energy of the excitation light leading to significant alterations of the totally reflected light intensity. Thus, a plot of the reflected light intensity versus incident angle is a quantitative measure of the resonance and constitutes a surface plasmon resonance spectrum (Salamon and Tollin, 1999b, 2000; Salamon et al., 1997c, 1999a ). Furthermore, any alteration in the optical properties of the metal/dielectric medium interface (such as, for example, immobilization of material) will affect the surface plasmon wave vector and, therefore, change the resonance characteristics, leading to changes in both the position and shape of the resonance curve. As can be seen from Eq. (6.1), these optical properties are fully described by the complex dielectric constant, e, which contains the refractive index, n, and the extinction coefficient, k (i.e., e ¼ nik) as well as the thickness, t, of a layer of material deposited at the interface. The sensitivity (Salamon and Tollin, 1999b, 2000), S, of measurements of such alterations can be defined as the change in reflectance (dR), measured at a specific angle, a1, within the range of the resonance curve, divided by the change in one of the three optical parameters (dn, dk, and dt), i.e.,

Sa1 ¼ ½dRðaÞ =dn dk dta1

ð6:2Þ

In general, the magnitude of changes in the experimental value of R are controlled by two factors: the shift of the position and the shape of the resonance spectrum. Both of these factors are related to the sharpness of the spectrum (i.e., its half-width); this defines the optical resolution (i.e., the smallest changes that can still be resolved as two different readings). Therefore, the overall sensitivity of a plasmon resonator will depend on both the magnitude of the electromagnetic field at the resonator surface and the optical resolution.

2.2. Varieties of surface plasmon resonances 2.2.1. Conventional surface plasmon resonance (SPR) In the most straightforward case, for which the hypotenuse of a right angle prism is coated with a single high-performance metal layer (usually Ag or Au, although gold is often used in biosensors because of its resistance to corrosion), one can generate surface plasmons on the outer surface of the

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Distance from the prism surface, nm

Normalized electric field amplitude

0

200

400

600

3 s

PWR (Ag, SiO2)

2 p

1

SPR (Au)

0 SiO2 layer Metal layer (Ag,Au)

Figure 6.1 Calculated electric field amplitudes for SPR and PWR resonators with excitation light wavelength 632.8 nm.The sensitivity of the resonator is proportional to the amplitude of the field at the external surface (gold layer in the case of SPR and silica layer in the case of PWR).

metal with visible light (typically 500 to 700 nm), as indicated in Fig. 6.1 for either a 55-nm Ag or a 48-nm Au layer (which are the optimal thicknesses for these metals) (Salamon and Tollin, 1999a,b; Salamon et al., 1997b,c). The electromagnetic wave created as a result of surface plasmon excitation is characterized by several important properties, which are very relevant in biosensor applications. First, there is an enormous increase in the intensity of the electromagnetic field generated by surface plasmons compared with that at the incident surface. It is also important to note that the energy of the field is proportional to the square of the electromagnetic wave intensity. This property further increases the sensitivity of the measurement. Second, the magnitude of the increase in the electromagnetic field intensity depends on the optical properties of the metal layer used to generate plasmons. Thus, as the calculation presented in Fig. 6.1 demonstrates, silver produces approximately a twofold higher intensity, which results in a fourfold higher overall sensitivity than gold.

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2.2.2. Plasmon-waveguide resonance (PWR) In 1997 we developed (Salamon and Tollin, 2001a, 2002; Salamon et al., 1997, 1999b) a variant of SPR that involves more complex assemblies in which surface plasmon resonances in a thin metal film are coupled with guided waves in a dielectric overcoating, resulting in excitation of both plasmon and waveguide resonances. We termed this phenomenon coupled plasmon-waveguide resonance (CPWR [Salamon and Tollin, 1999a,b; Salamon et al., 1999a]; later shortened to PWR). We constructed a spectrometer based on this technology, which we have applied to the study of biologic membrane systems (Hruby and Tollin, 2007; Salamon and Tollin, 2001b; Salamon et al., 1999a). We also licensed this technology to Proterion Corp., who developed and patented a PWR instrument (Anafi et al., 2004), which we have also used successfully in the study of biomembrane systems. A plasmon-waveguide resonator contains a metallic layer (as in a conventional SPR assembly), which is deposited on either a prism or a grating and is overcoated with either a single dielectric layer or a system of dielectric layers; these layers are characterized by appropriate optical parameters so that the assembly is able to generate surface resonances on excitation by both p- and s-polarized light components (Fig. 6.1). The addition of such a dielectric layer (or layers) to a conventional SPR assembly plays several important roles. First, it enhances the spectroscopic capabilities (because of excitation of resonances with both p- and s-polarized light components), which results in the ability to directly measure anisotropies in refractive index and optical absorption coefficient in a thin film immobilized onto the surface of the overcoating (Salamon and Tollin, 2001c). This allows one to obtain information regarding structural changes in the analyte, providing the material is oriented uniaxially at the interface. Second, it functions as an optical amplifier that significantly increases electromagnetic field intensities at the dielectric surface compared with conventional SPR, as illustrated by Fig. 6.1. This results in an increased sensitivity and spectral resolution (the latter caused by decreased resonance line widths, as also shown in Fig. 6.2). Usually these two resonances ( p- and s-) have different sensitivities resulting from different evanescent electrical field intensity distributions (Fig. 6.1). In the simplest PWR sensor comprised of silver covered with silica, the s-polarization is significantly more sensitive (fivefold to 10-fold) than that of p-polarization. It is important to note that both of these resonances are much more sensitive than conventional SPR. The latter resonators are usually based on a gold metal film. As noted previously, such a sensor is approximately four- to fivefold less sensitive than a silver-based one. The simplest PWR sensor described previously further increases the sensitivity by a factor of 4 (for p-polarization) or 10 (for s-polarization), resulting in an overall 20- to 50-fold increase in sensitivity compared with a conventional gold-based SPR sensor. It should also be noted that the PWR sensitivity could be

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Normalized reflectance

1.0

SPR (Au) 0.5

p

s

PWR (Ag, SiO2) 0.0 65

70 Incident angle, deg

75

80

Figure 6.2 Calculated resonance curves for SPR and PWR resonators shown in Fig. 6.1.

further amplified by modifying the simplest two-layered (silver and silica) sensor with more complex arrangements of dielectric layers. Such systems have been developed in our laboratory to be able to distinguish between different microdomains in lipid bilayer membranes (Salamon et al., 2005). These latter systems are characterized by an overall sensitivity at least two orders of magnitude higher that that obtained with conventional gold-based sensors. A third important characteristic of PWR is that the dielectric overcoating also serves as a mechanical and chemical shield for the thin metal layer. This allows reactive metals such as silver to be used in an aqueous environment, the latter being crucial for biosensor applications. It also provides a surface that can take advantage of the wide range of chemistries developed for the covalent immobilization of materials.

3. Sensor Construction and Sample Deposition Figure 6.3 shows a view of the sample compartment and sensor of a Proterion PWR instrument (Anafi et al., 2004). The sensor prism is pressed against a Teflon block containing a sample chamber (aqueous volume approx. 0.5 ml) and a reference compartment (aqueous volume approx. 0.1 ml) and sits adjacent to a solid-state detector that monitors the incident polarized CW laser light intensity reflected through the prism from the

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Flow manifold

Prism block

To computer

Teflon block Sample chamber

Detector connector

Bare reference chamber Cam clamp (closed position) Coated side

(Bare glass portion)

Detector

Prism

Prism block removed

Prism detail

Sample deposited here

Cam clamp (open position)

CW laser

SiO2 layer (~50 nm) Silver layer (~500 nm) Bare glass

Used for angle and amplitude calibration

Figure 6.3 Diagram of the prism holder and sample compartment of a Proterion PWR instrument.

backside of the sensor surface. The sample and reference chambers can be accessed through ports in the Teflon block holder. The sensor prism surface is partially coated with thin layers of silver and SiO2 on which the sample to be characterized is deposited. The laser beam can be positioned so that it is incident either on the sample region or on the bare glass region; the latter is used for incident angle and light intensity calibration purposes (the critical angle for total internal reflection is used as a reference point). This entire unit is mounted on a rotating table that allows the incident angle of the laser beam to be continuously varied with 1 mdeg resolution so as to obtain a computer readout of the angular dependence of the reflected light intensity (see Fig. 6.4). It is important to point out that the thickness and the quality (uniformity, purity, etc.) of the prism coatings are extremely important, because these control the angular position and the line width of the resonance signal. For a sample to influence the PWR resonance spectra obtained with a sensor prism in contact with an aqueous solution, it is desirable for it to be

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Detector Computer Prism Plasmon generating film (Ag + SiO2) Sample Aqueous medium Polarizer CW Laser Rotating table

Figure 6.4

Diagram of a typical PWR spectrometer.

immobilized within a distance from the surface corresponding to less than one wavelength of the monitoring light. This influence will diminish exponentially as one moves away from this surface. A variety of methods can be used for this purpose, including covalent attachment to the silica or physicochemical adsorption to the surface. Inasmuch as one of the main interests of our laboratory has been the characterization of membrane proteins, we will focus here on the use of lipid bilayers for immobilization.

4. Lipid Bilayer Deposition With the orifice in the sample compartment of the Teflon block as a support, it is possible to create a self-assembled single lipid bilayer on the silica surface of the resonator that is attached by a Gibbs border of lipid solution (Salamon et al., 1994, 1996a), in much the same way as was done across an orifice in a Teflon sheet separating two aqueous phases in the classical Mueller-Rudin experiment (Mueller et al., 1962). Such an annulus of lipid solution not only anchors the bilayer but also acts as a reservoir of lipid molecules. Bilayer formation is accomplished by depositing a small amount of a lipid solution (typically at a concentration of approximately 10 mg/ml of lipid in a mixture of squalene and butanol; 0.15:10, v/v) on the hydrated surface of the sensor prism and then filling the chamber with aqueous buffer. The hydrophilic surface of the silica attracts the polar head groups of the lipid molecules, thus forming a lipid monolayer deposited on a layer of adsorbed water, with the hydrocarbon chains oriented toward the droplet of excess lipid solution. Filling the cell with the

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1.0

Reflectance

0.8

0.6

0.4 Buffer: p-polarization Buffer: s-polarization Bilayer: p-polarization Bilayer: s-polarization

0.2

0.0 64

65 Incident angle (mdeg)

66

Figure 6.5 Typical PWR spectra obtained with a Proterion instrument from a silver/ silica sensor in contact with an aqueous buffer before and after deposition of an egg phosphatidylcholine bilayer.

appropriate aqueous solution initiates the second step, which involves a thinning process with the formation of both the second monolayer and the Gibbs border that attaches the bilayer film to the Teflon spacer, allowing the excess of lipid and solvent to move out of the orifice (Salamon and Tollin, 1999a,b). This typically occurs in 20 to 40 min and creates a very stable, highly flexible, bilayer into which integral membrane proteins can be inserted and onto which peripheral membrane proteins can be bound by electrostatic forces (see Figure 6.5). A variety of lipid compositions can be used to form such bilayers, and the aqueous environment can also be varied over a large range. This allows one to investigate microenvironmental effects on bilayer formation and protein immobilization.

5. Spectral Data Analysis As noted previously, the optical properties of a system can be described by three parameters: refractive index (n), extinction coefficient (k), and thickness (t). It is important to note that only thickness is a scalar quantity, whereas both n and k are tensors and, therefore, in general, they

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have different values along the different measurement axes. The refractive index is a macroscopic quantity and is related to the properties of individual molecules through the molecular polarizability tensor, as well as to the environment in which these molecules are located (e.g., packing density and internal organization). Similarly, the extinction coefficient is related to the molecular optical transition tensor (Salamon and Tollin, 2001b,c; Salamon et al., 2005). The distinction between thickness and the two other optical parameters is especially important when the molecules to be investigated are located in a matrix (such as a biomembrane or lipid bilayer membrane or any thin film) that has nonrandom organization and thus possesses long-range spatial molecular order. Such molecular ordering creates an optically anisotropic system, usually having a uniaxial optical axis resulting in two different principal refractive indices: ne (also denoted as nII or np) and n0 (also referred to as n or ns), and two different extinction coefficients: kp, and ks (Salamon and Tollin, 2001b). The first of these indices is associated with a linearly polarized light wave in which the electric vector is polarized parallel to the optical axis. The second one is observed with light in which the electric vector is perpendicular to the optical axis. This is the fundamental basis on which measurement of the optical properties of anisotropic systems can lead to the evaluation of their structural parameters. In the simplified case in which a molecular shape can be approximated by a rodlike structure and the molecules are ordered such that their long axes are parallel (e.g., phospholipid molecules in a lipid bilayer membrane), one has an optically anisotropic system whose optical axis is perpendicular to the plane of the lipid bilayer (Salamon and Tollin, 2001). The values of the refractive indices and extinction coefficient measured with two light polarizations (i.e., parallel, np and kp, and perpendicular, ns and ks, to the optical axis) will describe the optical (An) and extinction coefficient (Ak) anisotropies, as follows:




An ¼ ðnp Þ  ðns Þ  Ak ¼ ðkp  ks Þ kav 2

2




ðnav Þ2 þ 2



ð6:3aÞ ð6:3bÞ

where nav and kav are the average values of the refractive index and extinction coefficient. For a uniaxial system, in which the optical axis is parallel to the membrane normal, one has the following equations for the average values:


i1=2 2 2 nav ¼ 1=3 ðnp Þ þ 2ðns Þ h

kav ¼ ðkp þ 2ks Þ=3

ð6:4aÞ ð6:4bÞ

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Furthermore, as can be seen from the Lorentz-Lorenz relation for the refractive index and the Lambert-Beer relationship for the extinction coefficient, nav is also directly related to the mass surface density (Cuypers et al., 1983; Salamon and Tollin, 2001b):



  2 ðnav Þ2 þ 2 m ¼ 0:1M=N t ðnav Þ  1

ð6:5aÞ

whereas kav is also related to the surface concentration of chromophore:

C ¼ ð4p=lÞðkav =bÞ

ð6:5bÞ

where M is molecular weight, N is molar refractivity, t is the thickness of the membrane, C the molar concentration of chromophore, and b the molar absorptivity. For lipid molecules a reasonable approximation of M/N is 3.6 (Cuypers et al., 1983). Thus, from the refractive indices and extinction coefficients measured with two polarizations (np, ns, kP, and ks), and the thickness of the membrane (t), one can calculate the following parameters describing the physical characteristics of the membrane: (1) the surface mass density (or molecular packing density), i.e., mass per unit surface area (or number of moles per unit surface area) (Salamon and Tollin, 2001b,c), which reflects the surface area occupied by a single molecule; (2) the optical anisotropy (An), which reflects the spatial mass distribution created by both the anisotropy in the molecular polarizability and the degree of long-range order of molecules within the system (Salamon and Tollin, 2001c); (3) the surface chromophore density; and (4) the spatial distribution of chromophores. As noted previously, the experimental PWR spectra can be described by three parameters: spectral position, spectral width, and resonance depth. In the case of lipid bilayers, these features depend on such physical properties as the surface mass and/or chromophore density, the spatial mass and/or chromophore distribution, and the membrane thickness. Thin-film electromagnetic theory based on Maxwell’s equations provides an analytical relationship between the experimental spectral parameters and the optical properties (Salamon and Tollin, 1999a,b; Salamon et al., 1997c, 1999a). This allows evaluation of the n, k, and t parameters from which the membrane physical properties can be assessed. The fact that there are three experimentally measured spectral parameters and three optical parameters allows, in principle, a unique determination of the n, k, and t values by fitting a theoretical resonance curve to the experimental one. We have demonstrated such an approach by applying a nonlinear least-square fitting procedure to describe the structural consequences of the interaction of the human d-opioid receptor with some of its ligands

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(Salamon et al., 2000). With this approach we demonstrated that significantly different structural changes are induced in the d-opioid receptor on binding to different ligands. We were also able to quantify these differences and to propose a model representing changes in conformation and mass distribution of lipid and receptor molecules during interaction of the receptor with either agonist or antagonist molecules. Although such an approach to analysis of experimental data works well and is able to generate enhanced understanding of both the mass and structural alterations induced by molecular interactions within a thin film, in some cases another approach is necessary. In general, this arises from three principal reasons: (1) molecular interactions leading to a complex spectrum where it is no longer possible to obtain a unique determination of the optical parameters (e.g., they occur in a heterogeneous film), resulting in a final spectrum that represents a mixed population; (2) rapid conclusions about the interactions is required, which precludes a tedious and often difficult analysis; and (3) the resonance spectra are not good enough to be fitted by the theoretical curves (poorly resolved spectra usually are good enough to do some comparative experiments, but they cannot be used to quantify them by fitting procedures). In these cases there are two other approaches available: (1) spectral simulation (Alves et al., 2005a; Salamon et al., 2005), or (2) graphical analysis (Salamon and Tollin, 2004). The spectral simulation procedure is based on the same principle as the fitting approach described previously, but is quicker and easier to apply. We have used this method to characterize the lateral segregation of lipids and proteins into microdomains (rafts) in solidsupported bilayers. We were able to measure and simulate the PWR spectra of membranes formed from a single component lipid, as well as from a mixture of lipids. Iteration in such simulations was performed by manual variation of the optical parameters until an appropriate agreement with the experimental spectra was obtained. Application of this method of data analysis allowed us to assess the most important structural parameters of a lipid membrane, such as thickness, average surface area occupied by one lipid molecule (or molecular packing density), and degree of longrange molecular order. Furthermore, we were able to characterize segregated microdomains and demonstrate preferential association of protein molecules with one type of microdomain (Salamon et al., 2005). The graphical analysis procedure (Salamon and Tollin, 2004) is based on the following consideration. PWR spectra are determined by two physical properties of a thin film such as a lipid membrane: (1) an average surface mass density, and (2) the spatial distribution of mass within the system that results from the structure of the deposited film. The separation of mass changes from those caused by structure is achieved by transforming the measured spectral changes (e.g., changes in the position of the spectra obtained either with p- or s-polarized exciting light) from an (s-p)

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orthogonal coordinate system into one reflecting (mass-structure). To perform such a transformation one must be able to place mass and structural axes within the orthogonal (s-p) coordinate system. This can be done if one knows the following two properties of the measurement system: (1) the mass sensitivity of the p- and s-axes in the (s-p) coordinate system (i.e., the sensor must be calibrated either theoretically or experimentally); (2) the optical symmetry of the measured system (i.e., whether the system is optically isotropic or anisotropic). For an anisotropic system, one must assume the direction of the optical axis (i.e., whether the optical axis is parallel to the p- or to the s-polarization direction). The axes of a new (mass/structure) coordinate system can then be scaled with the original (s/p) coordinates. Each point on the mass axis (Dm) can be expressed by changes of the original coordinates (Ds) and (Dp) as

ðDm Þ ¼ ½ðDs Þ2m þ ðDp Þ2m 1=2

ð6:6aÞ

and on the structural axis:

ðDstr Þ ¼ ½ðDs Þ2str þ ðDp Þ2str 1=2

ð6:6bÞ

In this way the contribution of structural changes and mass alterations are expressed in terms of angular shifts.

6. Membrane Protein Insertion Integral membrane protein insertion into the solid-supported bilayers can be accomplished by detergent dilution methods. Care must be taken in the choice of the detergent used so that the detergent is both able to maintain the protein in its native state and is sufficiently mild so that it does not perturb the lipid bilayer. Octylglucoside, in our experience, is able to satisfy both conditions, although in certain cases dodecyl maltoside was able to maintain the receptor active for longer periods of time. However, because dodecyl maltoside greatly perturbs the lipid bilayer, the detergent was exchanged with octylglucoside before addition to the PWR sample cell. By injecting small aliquots of a solution of a protein in an octylglucoside-containing buffer into the sample compartment, in which the detergent is above the critical micelle concentration (CMC), thereby diluting the detergent to below the CMC, spontaneous incorporation into the bilayer occurs, usually over a period of minutes. Lipid molecules that are displaced by such insertion can be transferred into the Gibbs border. Similarly, the bilayer is flexible enough so that even large extramembrane

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segments can be accommodated. It should be noted that not all the protein that is injected into the PWR sample compartment is inserted into the bilayer. Indeed, from spectral analysis we have determined that only a small amount of protein is properly reconstituted in the bilayer (approximately 5% in the case of the d-opioid receptor) (Alves et al., 2005a). Because PWR is a very sensitive technique, the insertion of picomole quantities of protein is usually sufficient to obtain a good signal. The remaining protein, which is most probably deposited into the bottom of the PWR cell, is washed from the cell compartment by flowing buffer through the system. It should be pointed out that if one wants to compare the spectral changes induced by ligand and/or G-protein binding to the lipid bilayer, one has to either incorporate similar amounts of receptor (as judged by comparable spectral shifts) or to normalize the data so that the amount of incorporated receptor is taken into account. In Fig. 6.6 are presented the PWR spectra obtained for the incorporation of a GPCR, the human d-opioid receptor (hDOR) into an egg PC bilayer. Note that the incorporation of the receptor into the bilayer leads to anisotropic increases in the resonance angle position (190 mdeg shift for the p- and 130 mdeg for the s-polarized resonance) and in the spectral depth, that are the result of an increase in the mass and the thickness of the bilayer. Because the receptor protrudes from both sides of the lipid bilayer, one should expect the bilayer to become thicker on

A

B 1.2 1.0 Reflectance

Reflectance

1.0

0.8

0.8

0.6 0.6 0.4 63.5

63.9 64.3 64.7 Incident angle, deg

67.2

67.4 67.6 Incident angle, deg

67.8

Figure 6.6 PWR spectra obtained for lipid bilayer formation and receptor incorporation with p-polarized (A) and s-polarized (B) light excitation. Solid curves represent the buffer spectra (10 mM TRIS buffer (pH 7.3), 0.5 mM EDTA, and 10 mM KCl) before bilayer formation; dotted curves correspond to PWR spectra obtained after the formation of a lipid bilayer composed of 75:25 mol% egg PC/POPG; dashed curves are PWR spectra obtained after addition of an octylglucoside-containing buffer solution of hDOR; final concentration in the cell sample compartment 0.4 nM.

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receptor incorporation. Indeed, the thickness of the proteolipid system was found by spectral fitting (Salamon et al., 1997a, 2000) to increase from 5.3 nm to 6.8 nm (the latter value corresponds to the dimension of the incorporated protein molecule perpendicular to the membrane plane). A thickness of the proteolipid system of 6.8 nm correlates well with the size determined for rhodopsin from X-ray crystallography (Palczewski et al., 2000). One should note also that spectral shifts with p-polarization were larger than with s-polarization (indicating refractive index changes in the p-direction larger than for the s-direction), which is a consequence of the anisotropic structure (i.e., cylindrical shape) of the receptor molecules. This is also evidence for the incorporation of the receptor into the bilayer with the expected orientation (i.e., long axis oriented perpendicular to the lipid bilayer), rather than just adsorbed to the surface of the bilayer, clearly reflecting a corresponding increase of the average long-range molecular order in the membrane resulting from receptor-lipid interactions. The direction of receptor insertion is usually controlled by their extramembranous domains; for example, in several cases where these are known to be small (e.g., Family A of G-protein–coupled receptors) we have found that such incorporation is bidirectional (Alves et al., 2003; Salamon et al., 1996b, 2000). This is advantageous in that it allows both sides of an inserted protein to be probed; this is important when one wants to interrogate binding to both sides as is the case with ligand and G-protein binding. This will be further discussed in the following section.

7. Ligand and G-Protein Binding by GPCRs After incorporation of the receptor into the lipid bilayer, the ligandinduced receptor conformational changes can be followed both in kinetic and thermodynamic modes. For that, small aliquots of a ligand solution (usually tens of microliters) are successively introduced into the sample cell and the PWR spectral changes followed. Spectral shifts can be used to determine a thermodynamic binding constant for the ligand-protein interaction by plotting the shifts at equilibrium as a function of the concentration of ligand in the sample compartment. Because the amount of ligand bound is very much smaller than the total ligand present, as a consequence of the large difference in the volumes of the bilayer and the aqueous medium (approximately 1000-fold), the concentration of added ligand is approximately the same as the free ligand. Because the PWR shifts are directly proportional to the bound ligand concentration, KD values can be calculated from the hyperbolic dependence of such a plot. Affinity constants obtained in this manner are in good agreement with those determined by classical binding assays with radiolabel ligands, as exemplified in Table 6.1 for ligand

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Table 6.1 Affinity constants for binding of various ligands to the hDOR obtained with PWR and from previous literature with radiolabel methods KD (nM) obtained from PWR experiments Ligands

p-polarization

s-polarization

Ligand affinities obtained from literaturea (nM)

DPDPE pClDPDPE Deltorphin II SNC80 (-)tan67 TIPPPSI NTI Naloxone TMT-LTic Morphine Etorphine

14  3 2.9  0.7

18  5 3.3  0.8

16 1.6

0.88  0.05

1.2  0.3

0.7

52  8 3.2  1.2 1.1  0.1 0.025  0.001 83 2.5  0.3

57  12 3.7  1.5 1.2  0.1 0.023  0.004 81 3.2  0.2

56 6 1.22 0.028 10 9

520  30 0.3  0.1

b

1101 0.2

b

a

The references from which such binding constants have been obtained can be found in Alves et al. (2004a). s-polarized shifts were negligible for these ligands. Note: KD values were obtained from plotting the resonance minimum position (Y) for the PWR spectra as a function of ligand concentration (X) and fitting to the following hyperbolic function that describes the binding of a ligand to a receptor: Y = (BmaxX)/(KD þ X). Bmax represents the maximum concentration bound and KD is the concentration of ligand required to reach half-maximal binding. b

binding to the hDOR (Alves et al., 2004a). Moreover, contrary to classical surface plasmon resonance (SPR) methods where only mass changes can be investigated (and binding constants obtained), because these are sensitive only to p-polarized refractive index changes, with PWR the ligand-induced conformational changes of the receptor can be monitored by obtaining spectra with both polarizations. Such studies have been performed in our laboratories with several GPCRs: rhodopsin (activated by light rather than by ligand binding) (Alves et al., 2005b; Salamon et al., 1996b), the hDOR (Alves et al., 2003, 2004a), the beta-adrenergic receptor (Devanathan et al., 2004), the neurokinin receptor (Alves et al., 2006), and the cannabinoid receptor (Georgieva et al., 2008). Here, we will not go into the details of those studies but rather point out that they have provided important insights into the type and magnitude of the conformational changes of the receptors. As an example, in the case of the neurokinin 1 receptor, for which over the past 25 years there was a large controversy in the field, radiolabel binding studies have shown that the receptor possesses two binding sites (Sagan et al., 1997), although the origin of those two binding sites was unclear

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(Beaujouan et al., 2004). PWR studies performed by successive binding of one ligand to one site after the other site was occupied and seeing how the binding of the second ligand was affected led to the conclusion that the sites were distinct and non-interconvertible (Alves et al., 2006). Furthermore, careful spectral analysis and data deconvolution, as described in one of the preceding sections, allowed one to obtain detailed information on the nature of such receptor conformations (e.g., with rhodopsin and the hDOR) (Salamon et al., 1996b, 2000). These studies have confirmed the idea that GPCRs adopt a large multitude of conformations depending on the type of bound ligand, which is important to understand GPCR signaling (Alves et al., 2004a; Salamon et al., 2000). PWR has the great advantage of being able to directly monitor G-protein binding to a receptor (in the absence or presence of ligand) because: (1) contrary to classical pharmacologic methods usually used to learn about early signal transduction events, such as cAMP and GTPgS assays, PWR does not rely on downstream events; (2) it allows one to understand the contribution of each individual G-protein subtype, rather than a global response as obtained with studies in cells where several G-proteins are expressed at different levels. To establish a network of signal transduction with the G-protein subtypes that bind to a given receptor in the presence of a specific ligand, we have inserted a pre-bound receptor into the lipid bilayer and then assay its binding to a series of different G-protein subtypes (Alves et al., 2003). In view of the fact that the receptor can orient in the lipid bilayer exposing both the extracellular and intracellular faces to the aqueous side of the bilayer (the one that can be accessed), some fraction of the inserted receptor molecules have the intracellular side available for G-protein interaction. An example of this is shown in Fig. 6.7 for the binding of a G-protein mixture to the hDOR prebound to the agonist DPDPE. In this way, the hDOR has been examined for its G-protein affinities when bound to a large variety of ligands (agonists and partial agonists, antagonists, inverse agonists). We have not only determined that the affinity of the receptor for the G-protein is highly dependent on the type of bound ligand but that a high level of specificity exists in the interaction with the different G-protein subtypes (see Table 6.2 for the binding of different G-protein subtypes to ligand-bound hDOR) and that there is no relationship between the capacity of a ligand to induce G-protein binding and to activate GTPgS exchange (Alves et al., 2003). Furthermore, we were able to study the effect of having the G-protein bound to the receptor on its capacity to bind a ligand (the so-called low- and high-affinity states corresponding to the absence and presence of G-protein, respectively). In these experiments, because neither the G-protein nor the ligand is able to cross the lipid bilayer, we have chosen a different approach, with liposomes harboring both the receptor and G-protein under study that were fused with the lipid bilayer in the PWR cell (a process induced by

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60 p-polarization

Resonance position shift, mdeg

50 s-polarization 40

30 KD = 10.4 ± 0.4 nM

20

10

0 0

20

40

60

80

100

[G proteins], nM

Figure 6.7 Binding of a G-protein mixture (obtained from human brain) to the hDOR prebound to the agonist DPDPE: p-polarization (closed circles); s-polarization (open circles); solid lines correspond to hyperbolic fits to data points yielding KD value shown in figure.

calcium ions) (Alves et al., 2004b). In this way, some G-protein could be delivered to the side of the bilayer facing the prism. Subsequent ligand binding to the G-protein–receptor complex showed that the ligand affinity is greatly enhanced by the presence of G-protein bound to the receptor (Alves et al., 2004b). Such studies have provided important information on the initial signal transduction events.

8. Conclusions In this chapter we have provided a description of a novel biophysical method, plasmon-waveguide resonance (PWR) spectroscopy, that allows evaluation of the anisotropic structural properties of lipid and proteolipid bilayers, with s- and p-polarized light, as well as the structural changes that accompany perturbation of these structures by other biologically relevant molecules such as agonist and antagonist ligands, and other proteins that are involved in a biologic cascade, that directly interact with the integral membrane protein. For the first time, all of these studies, including the thermodynamic and kinetic parameters involved in these interactions, can be accomplished directly without the need for any modification of structure

Table 6.2 Binding affinities between the individual G-protein subtypes and the hDOR either unliganded or bound to various ligands and between GTPgS and the receptor–G-protein complex G-protein subtype

Goa

Polarization

p

Gia1

Gia2

Gia3

s

p

s

p

s

P

S

DPDPE bound KDG-protein (nM) 10  1 KDGTPgS (nM) 404  37

91 394  71

302  24 4.7  0.3

306  28 3.7  0.7

71 9.9  0.5

71 8.3  0.8

45  5 80  11

41  5 83  9

Unliganded receptor KDG-protein (nM) 20  1.7 KDGTPgS (nM) 1917  177

22  1.8 1883  219

79  9 *

81  8 *

598  70 *

574  70 *

95  10 *

96  10 *

45  4 880  135

36  8 89  11

31  7 96  9

298  29 1589  129

322  25 1712  135

18  3 925  139

16  3 896  145

4.8  0.4 12.5  1.9

215  33 2.2  0.4

209  33 2.4  0.3

13.1  1.1 92  13

14.2  1.1 102  15

18  2 26  3

19  2 23  3

Morphine bound KDG-protein (nM) 40  5 KDGTPgS (nM) 910  120 SNC 80 bound KDG-protein (nM) 5.2  0.5 KDGTPgS (nM) 8.9  1.8

* No PWR spectral shifts were obtained on addition of GTPgS up to 5 mM. Note: KD values were obtained from plotting the resonance minimum position for the PWR spectra as a function of G-protein concentration and fitting to the following hyperbolic function that describes the binding of a ligand to a receptor: Y = (Bmax  X)/(KD þ X). Bmax represents the maximum concentration bound and KD is the concentration of ligand required to reach half-maximal binding. The table has been obtained from Alves et al. (2004b).

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(radiolabels, fluorescent probes, dyes, etc.) that are necessary for evaluation of such interactions by other currently available biophysical and bioanalytical methods. Perhaps an even greater advantage of this new method is that it is exquisitely sensitive, requiring only femtomole quantities of integral membrane proteins. This is critical at the present time because most integral membrane proteins are not produced in large quantities naturally, and efforts to use more robust methods of molecular biology to obtain large amounts of these proteins have been only rarely successful because of the fragile stability of most integral membrane proteins and the lack of general biochemical methods to refold denatured membrane proteins to their biologically active structures. In this overview, we have illustrated the application of PWR to examine the structures of G-protein–coupled receptors (GPCRs), the largest class of integral membrane proteins in the human genome, and the targets for nearly 50% of all current drugs. We have demonstrated that we can directly evaluate the conformational changes that occur on interactions of these receptors with agonists, antagonists, partial agonists, and inverse agonists and have been able to determine the equilibrium-binding affinities of these ligands for the receptors, as well as the kinetics of these processes. Unlike SPR, the use of two polarizations to carry out these measurements allows one to obtain information about structural changes as well as changes in mass density. A possible criticism of these studies is that they are done in artificial model membrane structures, and thus the changes in structure observed, and the corresponding thermodynamic and kinetic properties obtained, may not correspond to what is happening in biologic cellular systems. We have addressed these concerns in detail previously (Hruby and Tollin, 2007). Most importantly, the binding affinities of ligands to the G-protein– coupled receptors we have examined, whether agonist, antagonist, partial agonist, or inverse agonist, have been the same in studies with PWR as those reported in the literature with membrane preparations from living systems or cellular assays (see Table 6.1). Furthermore, GTPgS assay results have been comparable to those seen in membrane and whole cell preparations from living systems. This is clear evidence that no major changes in protein conformation have occurred during the course of these measurements. On the other hand, it cannot be ruled out that the microenvironment that exists within a cell may modulate the detailed properties of the receptors. In fact, in those cases in which we have changed the lipid compositions of the bilayer, we have, indeed, observed quantitative changes in receptor thermodynamic properties (Alves et al., 2005a,b). Thus, care must be used in extrapolation of these results to in vivo systems. These studies also have far-reaching implications in a number of areas of biochemistry, pharmacology, and drug design. For example, our studies have demonstrated that GPCRs have multiple biologically relevant conformations, which are likely important for signaling. Interactions of

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agonists, antagonists, partial agonists, and inverse agonists all give different functional conformations than the unoccupied receptor. Thus, textbook depictions of the typical GPCR with a two-state model (i.e., inactive state and agonist-occupied active state) will need to be modified. Structurally, GPCRs are much more heterogeneous with respect to the structures that result from ligand-receptor interactions than implied by such a simplistic model. Indeed, in some cases peptide agonists and nonpeptide agonists for the same receptor can lead to different conformational states. Furthermore, the conformational diversity of receptor-ligand structure manifests itself in different effects on G-protein interactions with the receptor (Alves et al., 2004b; see Table 6.2). The implications of these and other observations that use PWR are still not completely clear and will require further studies. However, even at this early date it seems possible that they may provide new insights into the subclassification of some GPCRs (as for example mu1, mu2, etc. opioid receptors) based on the different structure-activity relationship of different classes of ligands. In this regard, it can be suggested that these varieties of structural changes might lead to different signaling pathways depending on the ways in which these different ligand-receptor conformations affect interactions with other signaling proteins such as adenylate cyclase, protein kinases, inositol phosphate, protein phosphatases, b-arrestins, ion channels and other proteins involved in GPCR signaling. These and other potential studies offer exciting new possibilities for the application of PWR spectroscopy to a variety of important biologic problems involving membranes, integral membrane proteins, and the biochemical machinery associated with their bioactivities.

Conflicts of Interests The authors have patents or patents pending on the instrumentation and applications of PWR spectroscopy.

ACKNOWLEDGMENTS This work was supported previously by grants from the National Science Foundation and the U. S. Public Health Service, National Institutes of Health, and National Institute of Drug Abuse.

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Tyrosine Sulfation of HIV-1 Coreceptors and Other Chemokine Receptors Hyeryun Choe* and Michael Farzan† Contents 1. Introduction 1.1. Disovery of tyrosine-sulfated peptides and proteins 1.2. Identification of the tyrosyl-protein sulfotransferases 1.3. Tyrosine sulfation of HIV-1 coreceptors 1.4. Tyrosine sulfation of other chemokine and related receptors 1.5. Tyrosine sulfation of coreceptor-binding site antibodies 1.6. Useful properties of sulfotyrosine 1.7. Approaches to studying tyrosine sulfation of chemokine receptors 2. Production and Use of Tyrosine-Sulfated Peptides Derived from Chemokine Receptors 2.1. Chemically synthesized tyrosine-sulfated peptides 2.2. Production of sulfated peptides in mammalian cells 2.3. Cell-free sulfation 2.4. Modulation of peptide sulfation 2.5. Uses of tyrosine-sulfated peptides 3. Study of Chemokine-Receptor Sulfation on the Plasma Membrane 3.1. Modulation of chemokine-receptor sulfation 3.2. Mutagenesis of candidate sulfotyrosines 4. Bacterial Expression of Tyrosine-Sulfated Peptides and Proteins 4.1. A system to introduce non-native amino acids into proteins 4.2. Advantages and uses of the system

* {

148 148 149 149 150 153 154 155 155 155 156 158 158 159 160 160 160 161 161 162

Department of Pediatrics, Harvard Medical School, Perlmutter Laboratory, Children’s Hospital, Boston, Massachusetts, USA Department of Microbiology and Molecular Genetics, Harvard Medical School, New England Primate Research Center, Southborough, Massachusetts, USA

Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05407-X

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2009 Elsevier Inc. All rights reserved.

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5. Protocols 5.1. Detection of sulfotyrosine of chemokine receptors 5.2. Metabolic labeling of peptide-Fc fusion proteins and removing the Fc domain 5.3. Inhibiting tyrosyl-protein sulfotransferae activity with smallhairpin RNAs 5.4. Cell-free sulfation of tyrosine-containing peptides 5.5. Expression of tyrosine-sulfated proteins in E. coli 6. Conclusions References

163 163 164 165 165 166 167 168

Abstract Sulfotyrosines contribute to a number of critical extracellular protein–protein interactions, including the association of the HIV-1 envelope glycoprotein with the HIV-1 coreceptor CCR5, a similar association between the Duffy binding protein of Plasmodium vivax and the Duffy antigen/receptor for chemokines, between complement components C5a and C3a and their respective receptors, and between many CC- and CXC-chemokines and their receptors. In addition, the antigen-combining regions of a number of human antibodies include sulfotyrosines that are necessary for antigen recognition. The study of sulfotyrosines requires an array of techniques, each with its advantages and limitations. These include modulation of tyrosyl-protein sulfotransferase activity in mammalian cell lines, production of tyrosine-sulfated peptides with direct chemical synthesis or enzymatic addition of sulfate to tyrosines in cell-culture or cell-free systems, and use of a novel tRNA/tRNA-synthetase pair capable of introducing sulfotyrosines at specific sites into bacterially expressed proteins. Here we describe the use of these various approaches to study the role of tyrosine sulfation of chemokine receptors in ligand binding and HIV-1 entry.

1. Introduction 1.1. Disovery of tyrosine-sulfated peptides and proteins Tyrosine sulfation was first described on fibrinopeptide B in 1954, and by the 1970s, a handful of secreted peptides had been shown to include functionally important sulfotyrosines (reviewed in Kehoe and Bertozzi [2000], Moore [2003], and Seibert and Sakmar [2008]). A greater appreciation for full extent of tyrosine sulfation came in 1982, with the observation that many proteins from mammalian cells could incorporate metabolically labeled sulfate on what amino-acid analysis showed to be tyrosines (Huttner, 1982, 1984). The first membrane protein shown to be modified by tyrosine sulfate was the P-selectin glycoprotein ligand (PSGL-1) (Pouyani and Seed, 1995; Sako et al., 1995). The flexible PSGL-1 aminoterminus includes both sulfotyrosines and adjacent O-glycosylation

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moieties. As was later shown for CCR5 and its natural ligands, these aminoterminal modifications cooperate in binding the PSGL-1 ligand, P-selectin.

1.2. Identification of the tyrosyl-protein sulfotransferases The identification of the enzymes responsible for addition of sulfate to tyrosines was a critical contribution to the study and production of sulfated proteins. Two enzymes, tyrosyl protein sulfotransferases 1 and 2 (TPST1, TPST2), mediate tyrosine sulfation in mammalian cells (Beisswanger et al., 1998; Ouyang and Moore, 1998; Ouyang et al., 1998). These enzymes, type II membrane proteins active in the trans-Gogi network, bind the universal sulfate doner 30 -phosphoadenosine 50 -phosphosulfate (PAPS) and transfer sulfate to tyrosines of exposed and flexible regions of lumenal domains of proteins (Suiko et al., 1992). The TPSTs recognize accessible tyrosines usually adjacent to several acidic residues, although in a proper context glycines or asparagines can also promote or at least permit sulfation. In contrast, glycosylated asparagines, basic residues, and phenylalanines seem to hinder sulfation and are typically not found adjacent to a sulfotyrosine (Bundgaard et al., 1997). To date, no clear specificity difference between the two TPSTs has been observed, nor has any tyrosine-specific sulfatase activity been described in mammalian cells (Moore, 2003).

1.3. Tyrosine sulfation of HIV-1 coreceptors The first chemokine receptor shown to be modified by tyrosine sulfation was the CC chemokine receptor and HIV-1 coreceptor CCR5 (Farzan et al., 1999). Figure 7.1 shows specific incorporation of sulfate into CCR5. CCR5 binds and signals in response to the CC-chemokines CCL3 (MIP1a, CCL4 (MIP-1b) and CCL5 (RANTES), and the receptor’s aminoterminal sulfotyrosines and O-linked glycosylation are essential for this association (Bannert et al., 2001). In addition, CCR5 together with the cellular receptor CD4 is essential for most primary HIV-1 isolates to enter a target T cell or macrophage (Choe et al., 1996). The HIV-1 envelope glycoprotein gp120, which with gp41 mediates the viral entry process, first binds CD4. CD4 association induces a conformational change in gp120 that permits subsequent association with CCR5 (Wu et al., 1996). After CCR5 association with gp120, gp41 mediates a large-scale structural rearrangement of the envelope glycoprotein resulting in mixing of the virion and target cell lipids and ultimately entry of the viral core into the cell. CCR5 sulfotyrosines, but not its O-glycosylation, are critical to gp120 binding and HIV-1 entry (Farzan et al., 1999). In some infected individuals, HIV-1 variants can emerge that gain the ability to enter cells by way of the chemokine receptor CXCR4 in addition to or instead of CCR5. Although tyrosine sulfation of CXCR4 is essential for association with this receptor’s

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1

2

3

4

5

6

7

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100 67 CD4 46 CCR5 30 cys/met

SO4

Figure 7.1 CCR5 incorporation of [35S]-sulfate. Cf 2Th cells (lanes 1, 3, 5, and 7) or Cf 2Th cells stably expressing CD4 and CCR5 (lanes 2, 4, 6, and 8) were incubated with [35S]-cysteine and -methionine (lanes 1 to 4) or [35S]-sulfate (lanes 5 to 8). Cells were lysed, and lysates were immunoprecipitated with the anti-CCR5 antibody 5C7 (lanes 1, 2, 5, and 6) or the anti-CD4 antibody OKT4a (lanes 3, 4, 7, and 8), and analyzed by SDSPAGE. Lanes 1 to 4 represent a 12-h film exposure, whereas lanes 5 to 8 represent the same gel exposed for 48 h. Numbers at the left of the figure indicate molecular weight in kilodaltons.

natural ligand, CXCL12 (SDF-1), this modification and, indeed, the entire CXCR4 amino-terminus play a much less pronounced role in the entry of CXCR4-using HIV-1 isolates than with CCR5-using isolates (Farzan et al., 2002a). In addition to these principal coreceptors, a number of chemokine receptors and similar proteins have been shown to support HIV-1 entry in cell-culture systems (Table 7.1). For example, the chemokine receptors CCR2b, CCR3, CCR8, CXCR6, and the related receptors gpr1, gpr15, and apj can support infection by one or more isolates (reviewed in Choe et al. [1998]). In contrast to CCR5 and CXCR4, physiologic roles for these minor coreceptors have not been described. However, their identification has highlighted biochemical properties essential for HIV-1 entry, namely the presence of an acidic and tyrosine-rich amino-terminal motif similar to CCR5 and CXCR4. In those cases that have been examined (CCR2b, CCR3, CCR5, CCR8, and CXCR4) these receptors are sulfated at their amino-terrminal tyrosines (Farzan et al., 1999, 2002a; Fong et al., 2002; Gutierrez et al., 2004; Preobrazhensky et al., 2000).

1.4. Tyrosine sulfation of other chemokine and related receptors In addition to the principal and minor HIV-1 coreceptors, a number of chemokine and related receptors rely on amino-terminal sulfotyrosines to bind their natural ligands, including CX3CR1, the C5a and C3a receptors,

Table 7.1 Sequences of amino-termini of chemokine receptors and related proteins. The amino-terminal sequences of the chemokine receptors and selected related receptors are shown. Bold receptor name indicates experimental demonstration of tyrosine sulfation. Sequences predicted or demonstrated to contain sulfotyrosines are indicated in bold. Receptors that function as principal or minor HIV-1 coreceptors are indicated with, respectively, triple or single plus signs Chemokine Receptors

ccr1 ccr2 ccr3 ccr4 ccr5 ccr6 ccr7 ccr8 ccr9 ccr10 cxcr1 cxcr2 cxcr3 cxcr4 cxcr5 cxcr6 cxcr7 cx3cr1 xcr1

HIV-1 coreceptor?

METPNTTEDYDTTTEFDYGDATPC MLSTSRSRFIRNTNESGEEVTTFFDYDYGAPC MTTSLDTVETFGTTSYYDDVGLLC MNPTDIADTTLDESIYSNYYLYESIPKPC MDYQVSSPIYDINYYTSEPC MSGESMNFSDVFDSSEDYFVSVNTSYYSVDSEMLLC MDLGKP. . .LLVIFQVCLCQDEVTDDYIGDNTTVDYTLFESLC MDYTLDLSVTTVTDYYYPDIFSSPC MTPTDFTSPIPNMADDYGSESTSSMEDYVNFNFTDFYC MGTEATEQVSWGHYSGDEEDAYSAEPLPELC MSNITDPQMWDFDDLNFTGMPPADEDYSPC MEDFNMESDSFEDFWKGEDLSNYSYSSTLPPFLLDAAPC MVLEVSDHQVLNDAEVAALLENFSSSYDYGENESDSCCTSPPC MEGISIYTSDNYTEEMGSGDYDSMKEPC MNYPLTLEMDLENLEDLFWELDRLDNYNDTSLVENHLC MAEHDYHEDYGFSSFNDSSQE MDLHLFDYSEPGNFSDISWPCNSSDCIVVDTVMC MDQFPESVTENFEYDDLAEAC MESSGNPESTTFFYYDLQSQPC

 þ þ  þþþ   þ þ     þþþ  þ    (continued )

Table 7.1 (continued) Related 7TMS receptors

TSHR DARC c5aR ccrl1 d6 chemr23 apj gpr1 gpr15

HIV-1 coreceptor?

GMGCSS. . .IGFGQELKNPQEETLQAFDSHYDYTICGDSEDMVC MGNCLH. . .LDFEDVWNSSYGVNDSFPDGDYDANLEAAAPCHSC MNSFNYTTPDYGHYDDKDTLDLNTPVD MALEQNQSTDYYYEENEMNGTYDYSQYELIC MAATASPQPLATEDADSENSSFYYYDYLDEVAFMLC MEDEDYNTSISYGDEYPDYLDSIVVLED MEEGGDFDNYYGADNQSE MEDLEETLFEEFENYSYDLDYYSLESDL MDPEETSVYLDYYYATSPNSDIR

     þ þ þ þ

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the thyroid-stimulating hormone receptor (TSHR), and the Duffy antigen/ receptor for chemokines (DARC) (Choe et al., 2005; Costagliola et al., 2002; Farzan et al., 2001; Fong et al., 2002; Gao et al., 2003). DARC, thought to be a scavanger receptor for several CC- and CXC-chemokines, mediates the invasion of reticulocytes by the malaria parasite Plasmodium vivax, and the association of DARC with the P. vivax Duffy-binding protein requires one of two sulfate groups present on DARC’s amino-terminal tyrosines (Choe et al., 2005). Table 7.1 shows tyrosine-sulfated regions of each of these seven-transmembrane segment receptors, as well as regions of related chemokine receptors also predicted to include sulfotyrosines.

1.5. Tyrosine sulfation of coreceptor-binding site antibodies Sulfotyrosines also play a necessary role in the recognition by some antibodies of a highly conserved coreceptor-binding region of the HIV-1 envelope glycoprotein (Choe et al., 2003). Tyrosine sulfate has been observed at the third complementarity detemining region (CDR3) of the heavy chain of at least five neutralizing antibodies that recognize this region (Table 7.2). However, sulfation is unlikely to be limited to antibodies of HIV-positive individuals. The heavy chain CDR3 is encoded by one of approximately 25 diversity genes, a number of which encode sequences rich in tyrosines and acidic residues. Although antibody sulfation likely requires relatively long and flexible CDR3 regions, the frequency of tyrosines and aspartic acids encoded by antibody heavy-chain diversity genes suggests that tyrosine sulfation of antibodies is not rare. Tyrosine-sulfated antibodies have become an important tool in understanding the coreceptor interaction with HIV-1 envelope glycoprotein. For example, a structure of the HIV-1 gp120 with CD4 and one such antibody has defined two gp120 Table 7.2 The heavy-chain CDR3 regions of coreceptor-binding site antibodies. The heavy-chain CDR3 regions of five tyrosine-sulfated HIV-1 neutralizing antibodies and of 17b, an antibody that recognizes the same gp120 epitope, are shown. Note that 17b is not sulfated despite the presence of an apparent sulfation motif, likely because of the inaccessibility of its tyrosines. The names of sulfated antibodies are shown in bold, as are CDR3 regions shown to include sulfotyrosines Antibody

Heavy-chain CDR3 sequence

E51 412d 47e C12 Sb1 17b

NSIAGVAAAGDYADYDGGYYYDMD PYPNDYNDYAPEEGMSWYFD GGEDGDYLSDPFYYNHGMD DVGPDWDNDDYYDRSGRGVFD RNPNEYYDENADYSTVYHYMD VYEGEADEGEYDNNGFLK

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Table 7.3 Alignment of a the CCR5 amino-terminus with a peptide derived from the heavy-chain CDR3 region of the tyrosine-sulfated HIV-1 neutralizing antibody E51. An alignment of residues suggesting that the basis of pDE51’s activity may be its homology with CCR5 in aspartic acids and sulfotyrosines that contact gp120. Homologous residues are shown in bold. The higher affinity of pDE51 for gp120 may be due to the absence of hydrophobic CCR5 residues that interact with CCR5 transmembrane regions

pDE51 CCR5

GDYA. . ..DYDGGYYYDMD MDYQVSSPIYDINYYTSEP. . .

sulfotyrosine-binding pockets that presumably bind CCR5 sulfotyrosines (Huang et al., 2007). In addition, a CCR5-mimetic peptide shown in Table 7.3 and derived from tyrosine-sulfated CDR3 region of another HIV-1 neutralizing antibody inhibits viral replication more efficiently than any CCR5-derived peptides and suggests ways in which affinity of sulfated peptides for chemokine ligands might be enhanced (Dorfman et al., 2006).

1.6. Useful properties of sulfotyrosine The observations of key roles for tyrosine sulfation in the entry processes of both HIV-1 and one of the two major forms of human malaria suggests that properties of sulfotyrosine are useful to these pathogens. One possibility is that sulfotyrosines, with a number of highly polarizable electrons distributed between the sulfate and phenyl groups, can bind specifically and with high affinity a diverse set of proteins. The specificity with which sulfotyrosinecontaining peptides bind various ligands is clear. For example HIV-1 gp120 will not bind a C5aR-derived peptide with two sulfotyrosines or a CCR5derived peptide with two phosphotyrosines, but it will associate with a CCR5-peptide with two sulfotyrosines (Cormier et al., 2000; Farzan et al., 2000). Similarly, C5a will not bind tyrosine-sulfated CCR5-derived peptides, but it will bind a C5aR peptide with the two sulfotyrosines found on the native receptor (Farzan et al., 2001). Despite this specificity, sulfotyrosines may permit a wider array of protein interactions than unmodified amino acids. The chemokine/receptor interaction is an example of this; most chemokines bind more than one receptor, and most chemokine receptors bind more than one chemokine. P. vivax and especially HIV-1 may also use this property of sulfotyrosine to modify their respective entry proteins in response to immune pressure while at the same time retaining a high affinity for their cellular receptors. Sulfate modification may have an additional function for chemokine receptors, because it distinguishes immature receptors from receptors that have passed through the trans-Golgi network, perhaps preventing intracellular association of chemokine and

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receptor in cells that produce both. A premature interaction between the HIV-1 envelope glycoprotein and CCR5 may be similarly avoided.

1.7. Approaches to studying tyrosine sulfation of chemokine receptors Despite central roles for tyrosine sulfation in many protein–protein interactions, working with sulfated peptides and proteins has proved challenging. Chemical synthesis of peptides bearing sulfotyrosines requires modification of standard synthesis protocols, is consequently of high cost, and typically limited to two sulfotyrosines on relatively short peptides. Because bacteria do not naturally sulfate proteins and because chemokine receptors, like other mammalian seven-transmembrane segment receptors, cannot be expressed in bacteria, most studies of chemokine-receptor sulfation have been performed in mammalian systems, with RNA interference or overexpression of the TPST enzymes. Cell-free systems for sulfating peptides and soluble receptor fragments have also been used, but peptide yields are typically limiting. Recently, an important contribution to the field has been made by Peter Schultz’s group (Liu and Schultz, 2006; Ryu and Schultz, 2006), who have developed a novel technology for introducing synthetic amino acids, including sulfotyrosines, into bacterially expressed proteins. The uses and limitations of each of these approaches are described below.

2. Production and Use of Tyrosine-Sulfated Peptides Derived from Chemokine Receptors 2.1. Chemically synthesized tyrosine-sulfated peptides Because chemokine receptors cannot be expressed as soluble proteins, biochemical characterization of tyrosine-sulfated peptides based on receptor amino-termini will likely play an increasingly important role in the study and inhibition of chemokine-receptor function. Building on earlier studies of secreted peptides such as gastrin and cholecystokinin, several groups investigated the potential of commercially synthesized tyrosine-sulfated peptides based on the sequence of the CCR5 amino-terminus. These peptides were useful in demonstrating the specificity of sulfotyrosine association with gp120, in clarifying the role of individiual CCR5 sulfotyrosines in this association, and in localizing the sulfate-binding region on gp120 (Cormier and Dragic, 2002; Cormier et al., 2001; Farzan et al., 2000; 2002b). They proved, however, dissappointing as inhibitors of HIV-1 entry, blocking viral replication with IC50s greater than 100 mM. There are several reasons for the poor performance of these first-generation CCR5 mimetics. First, although CCR5 has four sulfated tyrosines in its

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amino-terminus, difficulties in synthesis limited to two the number of sulfotyrosines that could be included in high-quality peptide preparations. Moreover, only shorter peptides (<25 amino acids) could be generated in this manner, and introduction of an adjacent disulfide bond proved prohibitively difficult and expensive. In addition to these problems of chemical synthesis, first-generation peptides, based directly on the CCR5 aminoterminus, retained a number of hydrophobic amino acids that do not interact with the HIV-1 envelope glycoprotein. Instead, experimental evidence suggested they interact with the remainder of the CCR5 protein, and, indeed, these peptides can mediate a interaction of a CCR5 variant lacking its amino-terminus with the HIV-1 envelope glycoprotein or with the chemokine CCL5 (Farzan et al., 2002b). It is likely that hydrophobic residues from the CCR5 amino-terminus contribute to inappropriate conformations or small-scale aggregation of these peptides in solution. Second-generation peptides, produced in mammalian cells and lacking these hydrophobic residues, are considerably more effective at blocking HIV-1 infection (Dorfman et al., 2006), although further improvements will be necessary before they can be considered as potential therapeutics. One such peptide is shown in Table 7.3 aligned with CCR5.

2.2. Production of sulfated peptides in mammalian cells The limitations and expense of chemically synthesized peptides prompted exploration of mammalian expression systems for studying the interaction between CCR5 and the HIV-1 envelope glycoprotein. To promote efficient production and secretion of these peptides, they have typically been expressed as fusion proteins with the Fc region of an antibody, usually human or mouse IgG (Fig. 7.2). Because these overexpression systems typically saturate the endogenous TPST activities of cell lines such as HEK293T cells, plasmids expressing a peptide-Fc fusion protein are cotransfected with a plasmid expressing one of the TPSTs. In our hands, TPST2 is the more efficient at sulfating CCR5-based or CCR5-like peptides. Fc-fusion proteins generated in this way can be purified on protein-A Sepharose beads with protocols standard for antibody purification. Note, however, if elution is performed with a highly acidic buffer, the eluant must be neutralized immediately. Otherwise, the acid-labile sulfates are likely to be lost. Elution with high salt, for example 3 M MgCl, can be used and may be preferable to avoid this possibility. The efficiency of sulfate incorporation can be monitored as in Fig. 7.1 by [35S]-sulfate labeling of cells in parallel, before harvesting of peptide, or through functional assays if function has already been established. If necessary, removal of the Fc domain can be accomplished through introduction of a thrombin cleavage site and incubation with thrombin (see section 5.2). Figure 7.3 shows an

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mock

patient sera

pCCR5-Ig

pE51-Ig

p47e-Ig

pC12-Ig

p17b-Ig

p412-Ig

Peptide-Fc fusion

ADA-Ig

S1-Ig

Beads alone

ADA-Ig

S1-Ig

Beads alone

DM1

CD4mim

Figure 7.2 The CDR3 regions of the sulfated antibody E51 functions as a CCR5 mimetic. Radiolabeled HIV-1 gp120 (ADA isolate) was incubated with radiolabeled supernatants from 293Tcells transfected with plasmid expressing the sulfated (412d C12, 47e, E51) or unsulfated (17b) heavy-chain CDR3 regions of the indicated antibody, or the CCR5 amino-terminus, or with patient sera. Each peptide was expressed as a fusion protein with the Fc region of human IgG1 and produced in the presence of exogenous TPST2. Mixtures of peptide-Fc and gp120 were precipitated with protein A-Sepharose and analyzed by SDS-PAGE. Numbers to the left of the panel indicate molecular weight in kilodaltons. Bands that migrate with the 67-kDa marker indicate residual dimeric forms of the peptide-Ig proteins. As shown in Table III, pE51 is highly similar to the CCR5 amino-terminus, but it lacks a number of the hydrophobic residues of CCR5 that may interfere with gp120 association.

DM1 CD4mim

8.5 3.5

Total

DM1 CD4mim Bound

Figure 7.3 A combined CCR5/CD4 mimetic peptide efficiently binds gp120. Plasmids encoding peptides formed of a 27-amino-acid CD4 mimetic peptide, with or without a 12-amino-acid CCR5 mimetic peptide, and fused to the Fc region of human IgG1were transfected into HEK293T cells, and culture supernatants were precipitated with protein-A Sepharose. CD4 mimetic and CD4/CCR5 mimetic peptides were eluted in 3 M MgCl2, dialyzed against PBS and incubated with thrombin, which cleaves a specific sequence engineered between the mimetic peptide and the Fc domain (see protocols). Free peptide was concentrated and incubated with HIV-1 gp120 and protein sera. Note that only the double CD4/CCR5 mimetic, but not the CD4 mimetic peptide, bound gp120 efficiently.

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60 min

20 min

Figure 7.4 Cell-free sulfation of a CCR5-derived peptide. Protein-A Sepharose beads bound to 200 ngTPST2 fused to the Fc region of human IgG1were incubated with 4 mCi (2 nM ) [35S]-PAPS (30 -phosphoadenosine 50 -phosphosulfate, the sulfate donor) and 2.5 mg (1 nM) unsulfated peptide derived from the first 22 amino acids of CCR5 (MDYQVSSPIYDINYYTSEPSQK) in a total volume of 100 ml for 20 or 60 min, as indicated. An aliquot of supernatant was analyze by SDS-PAGE on a 10 to 20% gradient Tricine gel. Note that longer incubation times generate more of the multiply sulfated peptides, which have greater mobility in these conditions.

example of a CCR5-mimetic peptide fused to a CD4-mimetic peptide that was prepared in this manner and precipitated with HIV-1 gp120.

2.3. Cell-free sulfation An alternative protocol for generation of sulfated peptides was orginally developed to study PSGL-1 and later applied to CCR5. Chemically synthesized or cell-produced peptides can be incubated with PAPS and a soluble form of TPST2 (see protocols). However, the cell-free TPST activity is significantly attenuated relative to TPST activity in a cell, so peptides generated in this way are usually studied in small quantities and with radiolabeling with [35S]-PAPS (Fig. 7.4). Because of the lower efficiency of cell-free TPST2, peptides with multiple tyrosines can be variably sulfated. This heterogeniety is usually not desired, but it has been useful for determining the order of sulfate addition to CCR5 and CXCR4 by mass spectrometry (Seibert et al., 2002, 2008).

2.4. Modulation of peptide sulfation With mammalian expression systems, unsulfated or partially sulfated control peptides can be generated in several ways. The first is through mutation of the sulfated tyrosines. Frequently, phenylalanine is chosen to replace the

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ST-shRNA

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ST-shRNA

Tyrosine Sulfation of Chemokine Receptors

37 scFv 25 cys/met

SO4

Figure 7.5 TPST-2 enhancement and shRNA inhibition of sulfation of a single-chain antibody E51. Plasmid encoding E51scFv was cotransfected into 293Tcells with plasmids expressing shRNAs complementary to the messages of TPST1 and TPST2 (STshRNA), with pcDNA3.1 (vector) or with a plasmid encoding TPST2.Transfected cells were divided and radiolabeled with [35S]-cysteine and -methionine or [35S]-sulfate. Radiolabeled cell supernatants were immunoprecipitated with protein L-Sepharose and analyzed by SDS-PAGE.

sulfated tyrosine, but for several reasons this may be an innappropriate choice. First, because of its greater hydrophobicity, phenylalanine led to aggregation or nonbiologic conformations of a peptide in solution. Second, when more than one sulfotyrosine is present on a peptide, and elimination of a single sulfate is desired, phenylalanine is especially inappropriate, because it will efficiently prevent sulfation of adjacent tyrosines. For these reasons, aspartic acid, which maintains peptide solubility and promotes sulfation of nearby tyrosines, can be a better choice. An alternative means by which unsulfated peptides can be generated is through the use of RNA interference targeting the cellular TPSTs (see protocols). As shown in Fig. 7.5, targeting of TPST1 and TPST2 by small hairpin RNAs 2 days before transfection of plasmid expressing an scFv or peptide-Fc fusion protein can prevent incorporation of up to 90% of the sulfate incorporated into same peptide produced in the presence of exgoneous TPST2 (Choe et al., 2003).

2.5. Uses of tyrosine-sulfated peptides Once generated, tyrosine-sulfated peptides and Fc-fusion proteins are used in a variety of assays. For example, they can block the binding of iodinated or [35S]-labeled ligand with receptor. Similarly, peptide can be used to inhibit the association of gp120/soluble CD4 complexes with CCR5 on the cell membrane, or an antibody recognizing the coreceptor-binding site of gp120. Sulfated peptides have been shown to complement a CCR5variant lacking its amino-terminus, but only in assays that can detect a transient association with receptor such as chemokine-mediated signaling

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monitered by calcium mobilization or an HIV-1 entry assay. Peptide-Fc fusion proteins can be used as antibodies to precipitate gp120, neutralize HIV-1 entry, or monitor conformational changes of the HIV-1 envelope glycoprotein expressed on the cell surface. With the introduction of structural elements such as disulfide bonds into these peptides and through elimination of hydrophobic residues that do not contribute to ligand binding, these peptides will be of further use in characterizing and inhibiting chemokine receptor and coreceptor function.

3. Study of Chemokine-Receptor Sulfation on the Plasma Membrane 3.1. Modulation of chemokine-receptor sulfation The study of tyrosine sulfation of intact chemokine receptors present an additional set of challenges, because the structure of the receptor ectodomain is determined by the transmembrane helices and cannot be manipulated in solution. These studies include binding assays with radiolabeled chemokine or gp120/CD4 complexes, signaling assays with an indicator of calcium mobilization, and HIV-1 entry assays quantified with an integrated reporter gene such as luciferase or green-fluorescent protein. Early study of tyrosine sulfate of cell-surface receptors used the TPST inhibitor chlorate to establish a functional role for tyrosine sulfation in receptor activity. However, because of its cytotoxity, chlorate can be used only in a narrow time window. Chlorate also inhibits the sulfation of O-glycosylation moieties and glycosaminoglycans, which typically promote lower affinity, low specificity interactions with chemokines and gp120. As with tyrosine sulfated peptides, a better current alternative for blocking TPST activity is the use of siRNA or shRNA targeting both sulfotransferases. As mentioned, shRNA can substantially diminish TPST activity to an extent detectable in functional assays. Because protein overexpressed in HEK293T cells can overwhelm the endogenous TPST activity of these cells, a functional role for tyrosine sulfate can also be established by comparison of receptor function in cells expressing exogenous TPST2 with shRNA or mock-transfected cells.

3.2. Mutagenesis of candidate sulfotyrosines Although these aforementioned assays can establish a role for tyrosine sulfation in chemokine receptor function, it is frequently important to know which receptor tyrosines are sulfated and which sulfotyrosines

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contribute most to function. Here mutagenesis is essential, and, as in peptide studies, alteration of tyrosine to aspartic acid is frequently preferable to alterations to phenylalanine. The extent of sulfation of wild-type and variant receptors can be quantitatively determined through comparison of [35S]cysteine and -methionine labeling to [35S]-sulfate incorporation into receptor. Functional assays of modified chemokine receptors require some means of measuring cell-surface expression, because removal of a sulfotyrsosine can have a suprisingly pronounced effect on receptor expression at the plasma membrane. Flow cytometric analysis can be used to quantify expression on the plasma membrane. This analysis can be performed with either an antibody recognizing a tag at the receptor amino-terminus, or, preferably, an antibody insensitive to chlorate treatment or to shRNA inhibition of the TPSTs. If a particular receptor variant expresses less efficiently than its wildtype counterpart, wild-type receptor expression should be varied by decreasing the amount of plasmid transfected. Although sometimes found in the literature, functional results should not be numerically normalized by dividing by expression levels, because expression and function rarely have a linear relationship.

4. Bacterial Expression of Tyrosine-Sulfated Peptides and Proteins 4.1. A system to introduce non-native amino acids into proteins A recent, major contribution to the study of tyrosine sulfation was made by Liu and Schultz (2006), who extended a novel system developed in the Schultz laboratory that permits introduction of synthetic amino acids into bacterially expressed proteins. This system uses a modified heterologous tRNA synthetase derived from Methanococcus jannashii to charge an M. jannashii amber tRNA with a nonnative amino acid. Liu and Schultz isolated a tRNA synthetase from a library of such enzymes positively selected to charge sulfotyrosine and negatively selected to avoid charging any other amino acids. The selected tRNA synthetase charges the M. jannashii amber tRNA, which then introduces sulfotyrosine during translation at any amber codon introduced into the gene. With this system, sulfotyrosine can be introduced into any site by modifying the corresponding codon to amber. To produce protein, plasmid bearing a modified gene is transformed into E. coli together with an additional plasmid encoding the heterologous tRNA/tRNA synthetase pair. Bacteria are grown in defined media in the presence of sulfotyrosine (see protocol below). As shown in Fig. 7.6, when sulfotyrosines are incorporated in this

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37 scFv 25

Figure 7.6 Bacterial expression of a tyrosine-sulfated 412d scFv. The open-reading frame of 412d scFv was cloned into bacterial expression vector pET11 (Novagen). A variant of this construct was made in which codons for two 412d heavy-chain CDR3 tyrosines were modified to amber (412d**, asterisks indicate amber-encoded sulfotyrosine). BL21(DE3) bacteria (Invitrogen) were transformed with pSUP-SY plasmid, which expresses both the M. jannashii amber suppressing tyrosyl tRNA (MjtRNATyrCUA) and the M. jannashii tyrosyl-transfer RNA synthetase (MjTyrRS) selected to charge the MjtRNATyrCUAwith sulfotyrosine. Bacteria were subsequently transformed with pET-scFv-412d or pET-scFv-412d**. Bacteria were grown overnight, diluted 1:100 in a defined media containing antibiotics and 10 mM sulfotyrosine, grown to an OD600 of 0.7, and induced with IPTG. Induction was continued for 24 h at room temperature; 15-ml aliquots of induced bacteria were lysed by boiling in SDS sample buffer, separated by SDS-PAGE, and blotted with an antibody that recognizes C9 tag at the scFv-C-terminus. Note that sulfotyrosines shift the mobility of the scFv variant and that no unsulfated scFv-412d** was detected.

manner at two sites in a single-chain antibody (scFv), the scFv expresses with comparable efficiency as the otherwise identical scFv encoding two unsulfated tyrosines.

4.2. Advantages and uses of the system This system has a number of advantages relative to chemical synthesis and mammalian expression systems for generating tyrosine-sulfated peptides and proteins. First, unlike chemical synthesis, there seems to be little limitation on number of sulfotyrosines incorporated, because peptides with as many as five sulfotyrosines express efficiently in this system. Second, in contrast to both mammalian expression and chemical synthesis, large quantities of proteins and peptides can be produced with standard bacterial expression protocols. Third, sulfotyrosine is incorporated homogenously at all desired sites. Fourth, the system does not depend on TPST activity, and it therefore allows introduction of a sulfotyrosine in amino-acid sequences that the mammalian TPSTs may not recognize. Finally, this system permits the phage-based selection of tyrosine-sulfated antibodies and peptides and has been recently used with phage display to generate de novo a tyrosine sulfated antibody that recognizes HIV-1 gp120 (Liu et al., 2008). Thus this system can begin to change the way tyrosine-sulfated proteins are produced, improved, and studied, ultimately allowing biologists to fully exploit the useful properties of sulfotyrosine.

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5. Protocols 5.1. Detection of sulfotyrosine of chemokine receptors 5.1.1. General considerations Sulfotyrosine can be detected through incorporation of radiolabeled sulfate into a chemokine receptor in which all glycosylation moieties have been removed. Removal of glycosylation is essential because carbohydrates are also frequently sulfated. Note that in conjunction with use of shRNA or exogenous TPSTs, this approach can obviate the need to show that sulfate is specifically incorporated at tyrosines with thin-layer chromotography, a more difficult and time-consuming approach (Corbeil et al., 2005). With a few exceptions it is difficult to immunoprecipitate endogenously expressed chemokine receptors from cell lysates, because few antibodies are available to do so. One exception is CXCR4, which can be precipitated from a number of cell lines, including HeLa cells, with the antibody 12G5 (R&D Systems). Generally instead a tag is added to the N- or C-terminus of the chemokine receptor. In most cases in our experience these tags do not disrupt function. Flag tag should be avoided at the amino-terminus, however, because the tag is highly acidic and its tyrosine itself is prone to sulfation. Note also that cells are lysed with sugar-based detergents such as DDM, cymal-5, or cymal-6 (Anatrace) rather than lipid-based detergents such as NP40 or Triton X100, because the former detergents permit more efficient extraction of the hydrophobic chemokine receptors. In addition, receptors are eluted off of protein A-Sepharose beads at 50 to 60  C rather than at boiling temperature to prevent aggregation and artifactual higher order bands in SDS-PAGE. 5.1.2. Radiolabeled reagents and labeling media To assess tyrosine sulfation of a given protein or peptide, we use [35S]cysteine and -methionine labeling (Perkin Elmer) to determine expression of the protein-of-interest and control proteins. Sulfate incorporation is measured with parallel labeling with [35S]-sulfate (Perkin Elmer). [35S]cysteine and -methionine labeling media is made by adding 50 mCi each of [35S]-cysteine and -methionine per ml of cysteine- and methionine-free cellculture media (Invitrogen or Sigma) containing, penicillin, streptomycin, glutamine, and 10% dialyzed fetal bovine serum (FBS, Invitrogen). [35S]sulfate labeling media is made by adding100 mCi of [35S]-sulfate per ml of sulfate-minus media (MEM, Sigma) containing 10% dialyzed FBS. Because cells need more sulfate than provided in 100 mCi [35S]-sulfate, for overnight labeling, unlabeled sodium sulfate is added to a final 1 mM concentration. Protocol: Detecting tyrosine sulfation of chemokine receptors 1. 3 to 4  106 HEK293T cells are plated onto T75 flasks. The following day, cells are transfected with a plasmid encoding tagged chemokine receptor and a plasmid encoding TPST2 at 10:1 ratio.

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2. The following day, cells are trypsinized, washed twice in phosphatebuffered saline (PBS), split at 1:5 ratio, resuspended in 1.5 ml of cysteine/ methionine-labeling media or 5 to 10 ml of sulfate-labeling media, and plated onto 1 (cysteine/methionine) or 3 (sulfate) wells of a 6-well plate. Because transfected HEK293T cells easily detach, polylysine-coated plates are recommended. 3. After overnight labeling, radioactive media is removed, and cells are either directly lysed in the wells or harvested in PBS containing 1 mM EDTA, and lysed in tubes, with 0.3 to 0.5 ml per well of 0.5% DDM or Cymal-5 (Anatrace) in PBS containing protease inhibitor cocktail (Sigma). 4. Cell debris is removed by microfuge centrifugation at 13,000 rpm for 10 min at 4  C, and tagged chemokine receptors are precipitated from the cleared lysate with anti-tag antibody and protein A-Sepharose at room temperature for 1 h. 5. Precipitates are washed three times with the same detergent used to lyse cells, once with PBS, and digested in a small volume (5 ml) containing protease inhibitor cocktail with 0.5 to 1 ml of N-glycosidase, endo F (PNGase F, New England Biolab), for 30 min at 37  C with intermittent mixing. Removal of N-glycosylation is useful, because sulfate can be associated with N- and O-glycosylation. 6. If removal of O-glycosylation is also necessary, wash N-glycosidase– digested samples once with 150 mM NaCl containing low concentration buffer (5 mM ) of neutral pH. Remove liquid completely, add a small volume (10 to 15 ml) of 1 O-glycosidase buffer containing O-glycosidase, neuraminidase, and protease inhibitor cocktail, and incubate for 1 h at 37  C with intermittent mixing. 7. At the end of digest, add reducing SDS-PAGE sample buffer directly to the samples, and incubate for 10 min at 50 to 60  C. As mentioned previously, boiling the samples will cause aggregation of chemokine receptors. 8. Resolve samples by SDS-PAGE, dry the gel, and expose it to an X-ray film. In most cases, chemical enhancing of signals is not necessary. Both [35S]-cysteine/-methionine and [35S]-sulfate signals should be detectable after overnight exposure.

5.2. Metabolic labeling of peptide-Fc fusion proteins and removing the Fc domain 5.2.1. General considerations 1. Peptide-Fc fusion proteins precipitated onto protein A-Sepharose beads are washed an additional time with 150 mM NaCl/20 mM Tris, pH 8.4. Liquid is removed completely, and 10 to 20 U of thrombin (Novagen) in a

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small volume is added into beads, and incubate at 37  C for 1 h with intermittent mixing. If protease inhibitors are used during immunoprecipitation and thrombin digestion, those of serine protease should be excluded. 2. Add 50 to 200 ml of PBS, vortex, spin beads, and transfer supernatant to a new tube. Repeat this step, combine the supernatant, and spin once again to pellet residual beads. Care should be taken not to include the beads, because they clog the membrane used in the next step. 3. Transfer this supernatant into Microcon centrifugal filter unit (Millipore) with 3 or 10 kDa molecular-weight cutoff (YM-3 or YM-10), spin in the microcentrifuge, and collect filter-through containing cleaved peptides. Thrombin and any contaminating Ig portion of the molecules are retained above the filter. The peptide can then be used in proteinbinding assays (as in Fig. 7.3) or for other functional studies.

5.3. Inhibiting tyrosyl-protein sulfotransferae activity with small-hairpin RNAs 5.3.1. General considerations The functional role of sulfotyrosines of chemokine receptors can be examined by modulating receptor sulfation or by mutating sulfated tyrosines. As shown in Fig. 7.5 for an scFv, tyrosine sulfation can be modulated with RNA interference with shRNAs, or, if the protein is overexpressed, by including exogenous TPST2. If interefering RNAs are used, cells have to be grown for at least 2 days after transfection with shRNA before they are used in functional assays to reduce the effect of previously translated TPST. In our experience, shRNA sequences derived from nucleotides 259 to 276 of the TPST1 gene (GCCATGCTGGACGCACAT) and nucloetides 73 to 92 of the TPST2 (GGACAGCAGGTGCTAGAGTG), transcribed by U6 promoter, substantially reduce TPST activities.

5.4. Cell-free sulfation of tyrosine-containing peptides 5.4.1. General considerations Tyrosine sulfation of peptides can be carried out in vitro with purified TPSTs and the sulfate donor, PAPS. Ig-fusion forms of TPST ectodomains (residues 25 through 370 of TPST1, and 25 through 396 of TPST2) can be produced efficiently and retain their enzymatic activities. Because in our hands TPST2-Ig more efficiently sulfates tyrosines on a CCR5-derived peptide than TPST1-Ig, the remainder of this protocol used TPST2-Ig. TPST2-Ig proteins are produced from transfected HEK293T cells into a serum-free media (Free-Style; Invitrogen). Figure 7.4 shows TPST2-Ig sulfation of four tyrosines on synthetic peptides derived from amino terminal 22 residues of CCR5 (MDYQVSSPIYDINYYTSEPSQK).

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Protocol: Cell-free sulfation of syntheticpeptides 1. TPST-Ig proteins produced in Free-Style media are bound to protein A-Sepharose and washed twice with 0.5 M NaCl/PBS and twice with PBS. 2. Beads containing approximately 0.2 ug of TPST-Ig, determined by analyzing an aliquot by SDS-PAGE, are mixed with 2.5 ug of peptide (1 nMoles) and 4 mCi (2 nMoles) of [35S]-PAPS (Perkin Elmer) in total volume of 100 ml, and incubated at 37  C. 3. At various time points up to 24 h, aliquots are taken, beads pelleted, and supernatants are analyzed by SDS-PAGE on a 10 to 20% gradient Tricine gel. Note that sulfation efficiency varies with sequence and number of target tyrosines.

5.5. Expression of tyrosine-sulfated proteins in E. coli 5.5.1. General considerations Although bacteria do not have a mechanism to incorporate sulfate into tyrosines, a novel system for introducing sulfotyrosines during translation in bacteria has been developed. Bacteria are sequentially transformed with two plasmids: one expressing the tRNA/tRNA synthetase pair (pSUP-SY, a generous gift from Chang Liu and Peter Schultz), and one encoding the protein-of-interest, whose gene is modified so that the amber codon (TAG) is introduced at sites where sulfotyrosine is desired. (Note that the natural stop codon should be altered if it is an amber.) Apart from supplementing culture media with sulfotyrosine (Bachem), standard bacterial expression protocols can be used. We transformed BL21(DE3) bacteria (Invitrogen) with pSUP-SY, made these transformed bacteria chemically competent, and transformed them with a pET11 vector expressing a single-chain form of the tyrosine-sulfated antibody 412d, modified to amber at sites encoding its sulfated tyrosines (pET11-scFv-412d**). A control plasmid in which the tyrosine codons are not replaced is also characterized (pET11-scFv-412d). The results are shown in Fig. 7.6. Protocol: Expression of a sulfated single-chain antibody in E. coli 1. Transform BL21(DE3) with pSUP-SY plasmid, and grow at 37  C in the presence of 30 mg/ml chloramphenicol. Wait for 2 days, because any colonies that appear within 1 day tend to have deletion of tRNA gene, because overexpressed tRNA is toxic. Also, large colonies likely have tRNA-gene deletion. 2. Expand a few colonies, extract plasmids, and sequence these to confirm that tRNA and synthetase genes are not deleted. 3. Prepare competent bacteria from a selected colony with standard methods, transform pET11 plasmid containing the gene-of-interest

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6. 7.

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bearing amber codons, and grow on plates containing 30 mg/ml of chloramphenicol and 100 mg/ml of ampicillin. A colony resistant to both antibiotics and, therefore, harboring both plasmids is grown overnight in LB containing 30 mg/ml of chloramphenicol and 100 ug/ml of ampicillin. Dilute the overnight culture 1:100 into GMML (1  M9 salt, 1% glycerol, and 0.3 mM leucine), and grow to saturation at 37  C. Dilute this culture 1:20 in GMML supplemented with up to 15 mM sulfotyrosine (Bachem), and grow with shaking at room temperature. During this step and induction, use 50 mg/ml ampicillin and 15 mg/ml chloramphenicol. At 0.6 to 0.7 of OD600, add IPTG to final 1 mM. Grow bacteria transformed with the wild-type gene for 2 h at 37  C, and those containing amber mutations for 12 to 48 h at room temperature. Check induction level by aliquoting 20 ml of bacteria suspension at various time points with and without induction. Boil these aliquots directly in reducing SDS-PAGE sample buffer, separate by SDS-PAGE, and visualize by Coomassie staining. To visualize the induced protein, the gel is transferred to PVDF membrane and blotted with an antibody that recognizes the protein or a tag. Sulfated and unsulfated proteins can be usually distinguished by their mobilities on a polyacrylamide gel; larger sulfated proteins tend to migrate more slowly (Fig. 7.6), whereas sulfated peptides tend to run faster (Fig.7.4). In both cases, this mobility shift is proportional to the number of sulfates.

6. Conclusions Despite the observation by Hutter in 1982 that the tyrosines of many proteins expressed in mammalian cells are modified by sulfate, it took a number of years to recognize the functional importance of these sulfotyrosines. Perhaps because of their unique biophysical properties, sulfotyrosines are frequently found at sites of protein–protein interactions, including interactions necessary for at least two important human pathogens. The discovery of the TPST enzymes began to make possible not just the scientific characterization but also the production and manupulation of tyrosine-sulfated proteins. The development of bacterial systems in which sulfotyrosine is genetically encoded will likely transform the way we improve, produce, and use tyrosine-sulfated peptides and proteins. This technology may make possible novel therapeutics including peptides and antibodies that control replication of pathogens or modulate inflammatory responses mediated by chemokine receptors and related proteins.

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Tyrosine-sulfated peptides functionally reconstitute a CCR5 variant lacking a critical amino-terminal region. J. Biol. Chem. 277, 40397–40402. Farzan, M., Mirzabekov, T., Kolchinsky, P., Wyatt, R., Cayabyab, M., Gerard, N. P., Gerard, C., Sodroski, J., and Choe, H. (1999). Tyrosine sulfation of the amino terminus of CCR5 facilitates HIV-1 entry. Cell 96, 667–676. Farzan, M., Schnitzler, C. E., Vasilieva, N., Leung, D., Kuhn, J., Gerard, C., Gerard, N. P., and Choe, H. (2001). Sulfated tyrosines contribute to the formation of the C5a docking site of the human C5a anaphylatoxin receptor. J. Exp. Med. 193, 1059–1066. Farzan, M., Vasilieva, N., Schnitzler, C. E., Chung, S., Robinson, J., Gerard, N. P., Gerard, C., Choe, H., and Sodroski, J. (2000). A tyrosine-sulfated peptide based on the N terminus of CCR5 interacts with a CD4-enhanced epitope of the HIV-1 gp120 envelope glycoprotein and inhibits HIV-1 entry. J. Biol. Chem. 275, 33516–33521. Fong, A. M., Alam, S. M., Imai, T., Haribabu, B., and Patel, D. D. (2002). CX3CR1 tyrosine sulfation enhances fractalkine-induced cell adhesion. J. Biol. Chem. 277, 19418–19423. Gao, J., Choe, H., Bota, D., Wright, P. L., Gerard, C., and Gerard, N. P. (2003). Sulfation of tyrosine 174 in the human C3a receptor is essential for binding of C3a anaphylatoxin. J. Biol. Chem. 278, 37902–37908. Gutierrez, J., Kremer, L., Zaballos, A., Goya, I., Martinez, A. C., and Marquez, G. (2004). Analysis of post-translational CCR8 modifications and their influence on receptor activity. J. Biol. Chem. 279, 14726–14733. Huang, C. C., Lam, S. N., Acharya, P., Tang, M., Xiang, S. H., Hussan, S. S., Stanfield, R. L., Robinson, J., Sodroski, J., Wilson, I. A., Wyatt, R., Bewley, C. A., and Kwong, P. D. (2007). Structures of the CCR5 N terminus and of a tyrosine-sulfated antibody with HIV-1 gp120 and CD4. Science 317, 1930–1934. Huttner, W. B. (1982). Sulphation of tyrosine residues-a widespread modification of proteins. Nature 299, 273–276. Huttner, W. B. (1984). Determination and occurrence of tyrosine O-sulfate in proteins. Methods Enzymol. 107, 200–223. Kehoe, J. W., and Bertozzi, C. R. (2000). Tyrosine sulfation: A modulator of extracellular protein-protein interactions. Chem. Biol. 7, R57–R61. Liu, C. C., Mack, A. V., Tsao, M. L., Mills, J. H., Lee, H. S., Choe, H., Farzan, M., Schultz, P. G., and Smider, V. V. (2008). Protein evolution with an expanded genetic code. Proc. Natl. Acad. Sci. USA 105, 17688–17693. Liu, C. C., and Schultz, P. G. (2006). Recombinant expression of selectively sulfated proteins in Escherichia coli. Nat. Biotechnol. 24, 1436–1440. Moore, K. L. (2003). The biology and enzymology of protein tyrosine O-sulfation. J. Biol. Chem. 278, 24243–24246. Ouyang, Y., Lane, W. S., and Moore, K. L. (1998). Tyrosylprotein sulfotransferase: purification and molecular cloning of an enzyme that catalyzes tyrosine O-sulfation, a common posttranslational modification of eukaryotic proteins. Proc. Natl. Acad. Sci. USA 95, 2896–2901. Ouyang, Y. B., and Moore, K. L. (1998). Molecular cloning and expression of human and mouse tyrosylprotein sulfotransferase-2 and a tyrosylprotein sulfotransferase homologue in Caenorhabditis elegans. J. Biol. Chem. 273, 24770–24774. Pouyani, T., and Seed, B. (1995). PSGL-1 recognition of P-selectin is controlled by a tyrosine sulfation consensus at the PSGL-1 amino terminus. Cell 83, 333–343. Preobrazhensky, A. A., Dragan, S., Kawano, T., Gavrilin, M. A., Gulina, I. V., Chakravarty, L., and Kolattukudy, P. E. (2000). Monocyte chemotactic protein-1 receptor CCR2B is a glycoprotein that has tyrosine sulfation in a conserved extracellular N-terminal region. J. Immunol. 165, 5295–5303.

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Ryu, Y., and Schultz, P. G. (2006). Efficient incorporation of unnatural amino acids into proteins in Escherichia coli. Nat. Methods 3, 263–265. Sako, D., Comess, K. M., Barone, K. M., Camphausen, R. T., Cumming, D. A., and Shaw, G. D. (1995). A sulfated peptide segment at the amino terminus of PSGL-1 is critical for P-selectin binding. Cell 83, 323–331. Seibert, C., Cadene, M., Sanfiz, A., Chait, B. T., and Sakmar, T. P. (2002). Tyrosine sulfation of CCR5 N-terminal peptide by tyrosylprotein sulfotransferases 1 and 2 follows a discrete pattern and temporal sequence. Proc. Natl. Acad. Sci. USA 99, 11031–11036. Seibert, C., and Sakmar, T. P. (2008). Toward a framework for sulfoproteomics: Synthesis and characterization of sulfotyrosine-containing peptides. Biopolymers 90, 459–477. Seibert, C., Veldkamp, C. T., Peterson, F. C., Chait, B. T., Volkman, B. F., and Sakmar, T. P. (2008). Sequential tyrosine sulfation of CXCR4 by tyrosylprotein sulfotransferases. Biochemistry 47, 11251–11262. Suiko, M., Fernando, P. H., Sakakibara, Y., Nakajima, H., Liu, M. C., Abe, S., and Nakatsu, S. (1992). Post-translational modification of protein by tyrosine sulfation: active sulfate PAPS is the essential substrate for this modification. Nucleic Acids Symp. Ser. 183–184. Wu, L., Gerard, N. P., Wyatt, R., Choe, H., Paralin, C., Ruffing, N., Borsetti, A., Cardosa, A. A., Desjardin, E., Newman, W., Gerard, C., and Sodroski, J. (1996). CD4-induced interactions of primary HIV-1 gp120 glycoproteins with the chemokine receptor CCR-5. Nature 184, 179–183.

C H A P T E R

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Activation Mechanisms of Chemokine Receptors Pia C. Jensen and Mette M. Rosenkilde Contents 1. 2. 3. 4.

Introduction Current Models for 7TM Receptor Activation Constitutive Activity of 7TM Receptors Activation of Chemokine Receptors by Small-Molecule Agonists 4.1. CCR8 small molecules–semiclassic pharmacophores for CC-chemokine receptors 4.2. CXCR3—small-molecule anchorage 4.3. CCR1—allosteric modulators 5. Experimental Procedures 6. IP3 Assay in Transiently Transfected COS-7 Cells 6.1. Detailed protocol References

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Abstract Chemokine receptors belong to the large family of 7-transmembrane (7TM) G-protein–coupled receptors. These receptors are targeted and activated by a variety of different ligands, indicating that activation is a result of similar molecular mechanisms but not necessarily similar modes of ligand binding. Attempts to unravel the activation mechanism of 7TM receptors have led to the conclusion that activation involves movements of the transmembrane segments VI and VII in particular, as recently gathered in the Global Toggle Switch Model. However, to understand the activation mechanism completely, more research has to be done in this field. Chemokine receptors are interesting tools in this matter. First, the chemokine system has a high degree of promiscuity that allows several chemokines to target one receptor in different ways, as well as a single chemokine ligand to target several receptors in different ways. Second, the endogenous ligands are large proteins that mainly activate their cognate receptors by interacting with various extracellular-located receptor regions.

Department of Neuroscience and Pharmacology, Laboratory for Molecular Pharmacology, The Panum Institute, University of Copenhagen, Copenhagen, Denmark Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05408-1

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It is, however, also possible to introduce agonism of simple ligands like metal ions. Thus, the chemokine system offers the possibility to test and compare the activation profiles of several chemically diverse ligands. This also brings up the interesting discussion of allosterism, because small molecules in the chemokine field often interact with allosteric receptor sites.

1. Introduction With more than 700 members, 7-transmembrane (7TM) G-protein– coupled receptors (GPCR) make up the largest superfamily of proteins in the human organism. Of the six subgroups (family A to F, based on sequence homology), family A (also known as the rhodopsin-like receptors) is by far the largest and most exploited (Gether, 2000; Schwartz, 1996). Chemokine receptors are found as a subgroup of the latter and count up to approximately 25 members (Murphy et al., 2000). 7TM receptors, in general, are activated by a diversity of endogenous chemical agents and exogenous stimuli such as peptides, ions, biogenic amines, lipids, nucleotides, odors, light, and taste (Okuno et al., 2008). Chemokines contain 70 to 80 amino acids, yet despite this (relatively) large ligand size, several nonpeptide agonists and antagonists have been identified within the chemokine system (Fig. 8.1). The huge ligand diversity within 7TM receptors indicates that receptor activation is not necessarily dependent on agonists binding to a specific set of residues in the conserved structure. That is, there is no common lock for all agonists (Schwartz and Rosenkilde, 1996a). It is, however, believed that a general activating mechanism is common for all 7TM receptors, irrespective the chemical nature of the agonist or the binding site (Schwartz and Rosenkilde, 1996b; Schwartz et al., 2006). Palczewski and colleagues (2000) published the first crystal structure of a mammalian 7TM receptor, the bovine rhodopsin in an inactive state, and in the following years the crystallization field was relatively silent. However, in 2007 and 2008 the field ‘‘exploded’’ with several convincing crystals of not only rhodopsin but also the b2- and b1-adrenergic receptors and the Adenosin A2 receptor (Hanson et al., 2008; Jaakola et al., 2008; Palczewski et al., 2000; Rasmussen et al., 2007; Warne et al., 2008). These crystals are all depicting receptors in inactive states constrained by antagonists or inverse agonists and by antibodies, T4-lysozyme fusion-constructs, or stabilizing receptor mutations (Hanson et al., 2008; Jaakola et al., 2008; Palczewski et al., 2000; Rasmussen et al., 2007; Warne et al., 2008). As a temporary climax, the first crystal structure of a 7TM receptor in a G-protein–interacting (active!!) conformation was presented in September 2008 (Scheerer et al., 2008). In this structure, the activated state of the receptor was mimicked by crystallizing opsin in complex with the main

CCR2 antagonist

CCR5 antagonists

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Pfizer UK-427,857 (Maraviroc)

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Millenneum LMD-009

ZK 756326 (uM)

Figure 8.1 Selection of small-molecule CC-chemokine receptor antagonists and agonists. The chemical compounds share a similar pharmacophore with a centrally located amine flanked by aromatic groups (except for ZK 756326 with aromatic groups only at one side). From left, the CCR2 antagonist: RS-504393 (10-[2-(5-methyl-2-phenyloxazol-4-yl)ethyl]-6-methylspiro{4H-3,1-benzoxazine-4,40 -piperidin}2(1H)-one) (Berkhout et al., 2003; Mirzadegan et al., 2000), the CCR5 antagonists: UK-427,857 (Maraviroc or 4,4-difluoro-N-((1S)3-{(3-endo)-3-[3-methyl-5-(1-methylethyl)-4H-1,2,4-triazol-4-yl]-8-azabi