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 ...

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METHODS IN ENZYMOLOGY Editors-in-Chief

JOHN N. ABELSON AND MELVIN I. SIMON Division of Biology California Institute of Technology Pasadena, California, USA Founding Editors

SIDNEY P. COLOWICK AND NATHAN O. KAPLAN

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32 Jamestown Road, London NW1 7BY, UK First edition 2009 Copyright # 2009, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: permissions@ elsevier.com. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made For information on all Academic Press publications visit our website at elsevierdirect.com ISBN: 978-0-12-374907-9 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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Contributors

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 Biotecnologıá/ 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 Munõz Department of Immunology and Oncology, Centro Nacional de Biotecnologıá/ 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 Biotecnologıá/ 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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Contributors

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

METHODS IN ENZYMOLOGY

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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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

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


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VOLUME 451. Autophagy: Lower Eukaryotes and Non-Mammalian Systems (Part A) Edited by DANIEL J. KLIONSKY VOLUME 452. Autophagy in Mammalian Systems (Part B) Edited by DANIEL J. KLIONSKY VOLUME 453. Autophagy in Disease and Clinical Applications (Part C) Edited by DANIEL J. KLIONSKY VOLUME 454. Computer Methods (Part A) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 455. Biothermodynamics (Part A) Edited by MICHAEL L. JOHNSON, JO M. HOLT, AND GARY K. ACKERS (RETIRED) VOLUME 456. Mitochondrial Function, Part A: Mitochondrial Electron Transport Complexes and Reactive Oxygen Species Edited by WILLIAM S. ALLISON AND IMMO E. SCHEFFLER VOLUME 457. Mitochondrial Function, Part B: Mitochondrial Protein Kinases, Protein Phosphatases and Mitochondrial Diseases Edited by WILLIAM S. ALLISON AND ANNE N. MURPHY VOLUME 458. Complex Enzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles and Peptides Edited by DAVID A. HOPWOOD VOLUME 459. Complex Enzymes in Microbial Natural Product Biosynthesis, Part B: Polyketides, Aminocoumarins and Carbohydrates Edited by DAVID A. HOPWOOD VOLUME 460. Chemokines, Part A Edited by TRACY M. HANDEL AND DAMON J. HAMEL VOLUME 461. Chemokines, Part B Edited by TRACY M. HANDEL AND DAMON J. HAMEL

C H A P T E R

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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

Posttranslational Modification of Chemokines

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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.


13

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 12,500

[NaCl] (M) (---)

A

8800

9200

0.4 0.2 0.0

8400

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


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

X

5 4

3 P

2

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

0

56 57 58 59 60 61

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


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

18

Tamara Loos et al.

O

O Fmoc-NH-CH-C – OH

Fmoc -NH-CH-C-O- resin

R2

R1

Y

X Deprotection (piperidine)

Activation (HBTU) O Fmoc-NH-CH-C-O-HBTU

Coupling

O H2N-CH-C-O- resin

R2

R1

Y

X Acetic anhydride capping

O

O

Repeat cycles

Fmoc-NH-CH-C-NH-CH-C-O- resin R2

R1

Y

X O

O

O

Fmoc - HN-CH-C-...-NH-CH-C-NH-CH-C-O - resin Rn

R2

R1

Z

Y

X Piperidine

O

O

O

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

R2

Z

Y

O

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.


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


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.

22

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

8400

8300 8350

8450

Mr

8+ 1057.8

0.50

7+ 1208.3

13+ 0.25 651.2

0.00 700

800

900

1000

1100

1200

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

8400

8450

Mr

1.0 13+ 646.1 8+ 1049.2

0.5

7+ 1198.8

6+ 1398.6

0.0 700

800

900

1000

1100

1200

1300

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)


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.

24

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


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.

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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).

Characterization of Chemokine Oligomerization

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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 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

38


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. 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

Chemokine and Glycosaminoglycan Interactions

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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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Damon J. Hamel et al.

% Hot ligand bound

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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


85

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 106 Daltons, and associating systems characterized by KD values between 104 to 108 M1 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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0 5.9

0.07

6.0 Radius

Single species

Residuals

0.14

Variance

18.0 ⫻ 10−5

Monomer-dimer

0 −0.14

6.1

7.0

0.06

7.1 Radius

Monomer-dimer-tetramer

0

0 −0.07

0.5

−0.06 0.05

Variance 81.5 ⫻ 10−5

Monomer-tetramer

0 Variance 92.0 ⫻ 10−5

−5 −0.05 Variance 3.7 ⫻ 10

7.2


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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+

1300

1500

1700

1900

2100

[M(CCL8)]4+

2300

2500

m/z

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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van Berkel, V., Barrett, J., Tiffany, H. L., Fremont, D. H., Murphy, P. M., McFadden, G., Speck, S. H., and Virgin, H. I. (2000). Identification of a gammaherpesvirus selective chemokine binding protein that inhibits chemokine action. J. Virol. 74, 6741–6747. Vives, R. R., Sadir, R., Imberty, A., Rencurosi, A., and Lortat-Jacob, H. (2002). A kinetics and modeling study of RANTES(9-68) binding to heparin reveals a mechanism of cooperative oligomerization. Biochemistry 41, 14779–14789. Webb, L. M., Ehrengruber, M. U., Clark-Lewis, I., Baggiolini, M., and Rot, A. (1993). Binding to heparan sulfate or heparin enhances neutrophil responses to interleukin 8. Proc. Natl. Acad. Sci. USA 90, 7158–7162. Xu, G., and Chance, M. R. (2007). Hydroxyl radical-mediated modification of proteins as probes for structural proteomics. Chem. Rev. 107, 3514–3543. Yu, Y., Sweeney, M. D., Saad, O. M., Crown, S. E., Hsu, A. R., Handel, T. M., and Leary, J. A. (2005). Chemokine-glycosaminoglycan binding: Specificity for CCR2 ligand binding to highly sulfated oligosaccharides using FTICR mass spectrometry. J. Biol. Chem. 280, 32200–32208. Zhuang, T., Leffler, H., and Prestegard, J. H. (2006). Enhancement of bound-state residual dipolar couplings: Conformational analysis of lactose bound to Galectin-3. Protein Sci. 15, 1780–1790.

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 Munõz, 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 Biotecnologıá/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,

Analysis of Chemokine Receptor Dimerization

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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-Falcoń 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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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

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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

Pulsed laser

CFP

CFP

Lifetime 1

Lifetime 2

YFP

B 150

150

125

125 2.5

Lifetime (ns)

1 0.5

Pixels

Lifetime (ns)

2 1.5

100 75 50

2 1.5 1 0.5

0

100 75 50

0

25

25

0

0 0

CCR5-CFP 2.35 ± 0.01 ns

Pixels

3

3 2.5

1

2

3

Fitted lifetime (ns)

0

CCR5-CFP/ CCR2-YFP 1.82 ± 0.03 ns

1

2

3

Fitted lifetime (ns)

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

59

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

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Zdzislaw Salamon et al.

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.

Applications of Plasmon Resonance Imaging to GPCR Signaling

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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


135

(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)

136


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

137


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

140


(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

141


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.

REFERENCES Alves, I. D., Salamon, Z., Varga, E., Yamamura, H. I., Tollin, G., and Hruby, V. J. (2003). Direct observation of G-protein binding to the human delta-opioid receptor using plasmon-waveguide resonance spectroscopy. J. Biol. Chem. 278, 48890–48897.


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Alves, I. D., Cowell, S. M., Salamon, Z., Devanathan, S., Tollin, G., and Hruby, V. J. (2004a). Different structural states of the proteolipid membrane are produced by ligand binding to the human d-opioid receptor as shown by plasmon-waveguide resonance spectroscopy. Mol. Pharmacol. 65, 1248–1257. Alves, I. D., Ciano, K. A., Boguslavski, V., Varga, E., Salamon, Z., Yamamura, H. I., Hruby, V. J., and Tollin, G. (2004b). Selectivity, cooperativity, and reciprocity in the interactions between the delta-opioid receptor, its ligands, and G-proteins. J. Biol. Chem. 279, 44673–44682. Alves, I. D., Salamon, Z., Hruby, V. J., and Tollin, G. (2005a). Ligand modulation of lateral segregation of a G-protein-coupled receptor into lipid microdomains in sphingomyelin/ phosphatidylcholine solid-supported bilayers. Biochemistry 44, 9168–9178. Alves, I. D., Salgado, G. F., Salamon, Z., Brown, M. F., Tollin, G., and Hruby, V. J. (2005b). Phosphatidylethanolamine enhances rhodopsin photoactivation and transducin binding in a solid supported lipid bilayer as determined using plasmon-waveguide resonance spectroscopy. Biophys. J. 88, 198–210. Alves, I. D., Delaroche, D., Mouillac, B., Salamon, Z., Tollin, G., Hruby, V. J., Lavielle, S., and Sagan, S. (2006). The two NK-1 binding sites correspond to distinct, independent, and non-interconvertible receptor conformational states as confirmed by plasmon-waveguide resonance spectroscopy. Biochemistry 45, 5309–5318. Anafi, D., Ramsay, G., MacDonald, J., Halatin, P., and Schwartz, Ch. (2004). Beam shifting surface plasmon resonance system and methods. US Patent #: 6,768,550 B2. Beaujouan, J. C., Torrens, Y., Saffroy, M., Kemel, M. L., and Glowinski, J. A. (2004). 25 year adventure in the field of tachykinins. Peptides 25, 339–357. Cuypers, P. A., Corsel, J. W., Janssen, M. P., Kop, J. M. M., Hermens, W. T., and Hemker, H. C. (1983). The adsorption of prothrombin to phosphatidylserine multilayers quantitated by ellipsometry. J. Biol. Chem. 258, 2426–2431. Devanathan, S., Yao, Z., Salamon, Z., Kobilka, B., and Tollin, G. (2004). Plasmonwaveguide resonance studies of ligand binding to the human beta 2-adrenergic receptor. Biochemistry 43, 3280–3288. Georgieva, T., Devanathan, S., Stropova, D., Park, C. K., Salamon, Z., Tollin, G., Hruby, V. J., Roeske, W. R., Yamamura, H. I., and Varga, E. (2008). Unique agonist-bound cannabinoid CB1 receptor conformations indicate agonist specificity in signaling. Eur. J. Pharmacol. 581, 19–29. Hruby, V. J., and Tollin, G. (2007). Plasmon-waveguide resonance (PWR) spectroscopy for directly viewing of GPCR/G-protein interactions and quantifying affinities. Curr. Opin. Pharmacol. 7, 1–8. Mueller, P., Rudin, D. O., Tien, H. T., and Wescott, W. C. (1962). Reconstitution of cell membrane structure in vitro and its transformation into an excitable system. Nature 194, 979–980. Palczewski, K., Kumasaka, T., Hori, T., Behnke, C. A., Motoshima, H., Fox, B. A., Le Trong, I., Teller, D. C., Okada, T., BStenkamp, R. E., Yamamoto, M., and Miyano, M. (2000). Crystal structure of rhodopsin: A G protein-coupled receptor. Science 289, 739–745. Sagan, S., Beaujouan, J.-C., Torrens, Y., Saffroy, M., Chassaing, G., Glowinski, J., and Lavielle, S. (1997). High affinity binding of [3H]propionyl-[Met(O2)11]substance P(7-11), a tritiated septide-like peptide, in Chinese hamster ovary cells expressing human neurokinin-1 receptors and in rat submandibular glands. Mol. Pharmacol. 52, 120–127. Salamon, Z., Wang, Y., Tollin, G., and Macleod, H. A. (1994). Assembly and molecular organization of self-assembled lipid bilayers on solid substrate monitored by surface plasmon resonance spectroscopy. Biochim. Biophys. Acta 1195, 267–275. Salamon, Z., Schmidt, R. A., Tollin, G., and Macleod, H. A. (1996a). Reusable biocompatible interface for immobilization of materials on a solid support. US Patent #: 5,521,702.

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Salamon, Z., Wang, Y., Soulages, J. L., Brown, M. F., and Tollin, G. (1996b). Surface plasmon resonance spectroscopy studies of membrane proteins: Transducin binding and activation by rhodopsin monitored in thin membrane films. Biophys. J. 71, 283–294. Salamon, Z., Macleod, H. A., and Tollin, G. (1997a). Coupled plasmon-waveguide resonators: A new spectroscopic tool for probing proteolipid film structure and properties. Biophys. J. 73, 2791–2797. Salamon, Z., Macleod, H. A., and Tollin, G. (1997b). Surface plasmon resonance spectroscopy as a tool for investigating the biochemical and biophysical properties of membrane protein systems. I. Theoretical principles. Biochim. Biophys. Acta 1331, 117–129. Salamon, Z., Macload, H. A., and Tollin, G. (1997c). Surface plasmon spectroscopy as a tool for investigating the biochemical and biophysical properties of membrane protein systems. II. Applications to biological systems. Biochim. Biophys. Acta 1331, 131–152. Salamon, Z., and Tollin, G. (1999a). Surface plasmon resonance; Theory. In ‘‘Encyclopedia of Spectroscopy and Spectrometry.’’ ( J. C. Lindon, G. E. Tranter, and J. L. Holmes, eds.) Vol. 3, pp. 2311–2319. Academic Press, New York. Salamon, Z., and Tollin, G. (1999b). Surface plasmon resonance; Applications. In ‘‘Encyclopedia of Spectroscopy and Spectrometry.’’ ( J. C. Lindon, G. E. Tranter, and J. L. Holmes, eds.) Vol. 3, pp. 2294–2301. Academic Press, New York. Salamon, Z., Brown, M. F., and Tollin, G. (1999a). Plasmon resonance spectroscopy: Probing interactions within membranes. Trends Biochem. Sci. 24, 213–219. Salamon, Z., Tollin, G., and Macleod, H. A. (1999b). Coupled plasmon-waveguide resonance spectroscopic device and method for measuring film properties. US Patent #: 5,991, 488. Salamon, Z., and Tollin, G. (2000). Surface plasmon resonance spectroscopy in peptide and protein analysis. In ‘‘Encyclopedia of Analytical Chemistry.’’ (R. A. Meyers, ed.), pp. 6050–6061. John Wiley & Sons Ltd., Chichester. Salamon, Z., Cowell, S., Varga, E., Yamammura, H. I., Hruby, V. J., and Tollin, G. (2000). Plasmon resonance studies of agonist/antagonist binding to the human delta-opioid receptor: New structural insights into receptor-ligand interactions. Biophys. J. 79, 2463–2474. Salamon, Z., and Tollin, G. (2001a). Coupled plasmon-waveguide resonance spectroscopic device and method for measuring film properties in the ultraviolet and infrared spectral ranges. US Patent #: 6,330,387 B1. Salamon, Z., and Tollin, G. (2001b). Plasmon resonance spectroscopy: Probing molecular interactions at surfaces and interfaces. Spectroscopy 15, 161–175. Salamon, Z., and Tollin, G. (2001c). Optical anisotropy in lipid bilayer membranes: Coupled plasmon-waveguide resonance measurements of molecular orientation, polarizability, and shape. Biophys. J. 80, 1557–1567. Salamon, Z., and Tollin, G. (2002). Coupled plasmon-waveguide resonance spectroscopic device and method for measuring film properties in the ultraviolet and infrared spectral ranges. US Patent #: 6,421,128 B1. Salamon, Z., and Tollin, G. (2004). Graphical analysis of mass and anisotropy changes observed by plasmon-waveguide resonance spectroscopy can provide useful insights into membrane protein function. Biophys. J. 86, 2508–2516. Salamon, Z., Devanathan, S., Alves, I. D., and Tollin, G. (2005). Plasmon-waveguide resonance studies of lateral segregation of lipids and proteins into microdomains (rafts) in solid-supported bilayers. J. Biol. Chem. 280, 11175–11184.

C H A P T E R

S E V E N

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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Hyeryun Choe and Michael Farzan

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

8

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 (1 mM, E39A ¼ 710 nM, K49A ¼ 96 nM. The mutants inhibited parasite invasion at ligand concentrations that were consistent with their receptor binding affinities for DARC (Hesselgesser et al., 1995). For example, the mutant E6A was almost as effective as CXCL1 with an EC50 of inhibition of invasion of 8.6 nM compared with 7 nM for wild-type CXCL1. The mutant E6A binds to DARC with high affinity and efficiently blocks parasite invasion (Fig. 9.3) but binds to CXCR2 poorly and does not activate neutrophils. This assay demonstrates the feasibility of blocking parasite invasion with inhibitors of chemokines and strongly suggests that screening for small-molecule inhibitors of parasite invasion with the assays described here could constitute a new and novel approach in the fight against P. vivax–induced malaria. Several other low-throughput assays that can measure the potential effectiveness of agents in blocking P. vivax–mediated infection have been described. Most are indirect assays, for example, like the erythrocyte rosetting assay described by Chitnis and Miller (Chitnis and Miller, 1994).

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COS7 cells transfected with PvRII are plated in 3.5-cm diameter wells and used for erythrocyte binding assays 40 to 60 h after transfection; 200 ml of a 10% erythrocyte suspension is added to 2 ml of media in wells containing the transfected cells, the plate is swirled to mix the erythrocytes well, and the erythrocytes are allowed to settle for 2 h at 37 C. The COS7 cells are then washed three times with 2 ml of PBS to remove nonadherent erythrocytes. Transfected COS7 cells with rosettes of adherent erythrocytes are then scored. The number of rosettes is scored in either 10 or 20 fields at a magnification of 40 with an inverted microscope. Binding is scored as negative when no rosettes are seen in the entire well. To study the effect of inhibitors of the interaction between DARC and PvRII the erythrocytes are first resuspended to a hematocrit of 1% in 1 ml of complete DMEM and incubated for 1 h at room temperature with the potential inhibitors at the required concentrations. The erythrocytes are then used in erythrocyte binding assays as described previously. The number of COS7 cells with rosettes of adherent erythrocytes is scored in 20 randomly chosen fields at a magnification of 40 in each well and the percent inhibition is determined as follows: percent binding ¼ 100 (no. of bound COS7 cells in the presence of inhibitors)/(no. of bound COS7 cells in absence of inhibitors); percent inhibition ¼ 100 percent binding; and percent inhibition ¼ 0 if binding (%) i > 100.

3. Methods for the Study of DARC as a Chemokine Sink Shortly after its discovery, DARC was postulated to be a ‘‘sink’’ for chemokines (Darbonne et al., 1991; Neote et al., 1993), and a study to test this concept was carried out in healthy volunteers who were given LPS intravenously to induce chemokines in the circulation (Olszyna et al., 2001). Injection of LPS was associated with increases in the erythrocyte bound levels of the chemokines CXCL1, CXCL8, and CCL2 that all bind to DARC. In contrast CCL4, which was measured as a non-DARC–binding chemokine, remained very low or undetectable in cell fractions after LPS administration. Endotoxemia and gram-negative sepsis, in which LPS plays a major role, are characterized by elevated levels of chemokines in plasma, and a recent study showed that in patients with sepsis, CXCL8 bound to erythrocytes exceeds CXCL8 concentrations in plasma (Marie et al., 1997). In another study, CXCL8 was shown to be released in the plasma of patients with acute myocardial infarction and readily binds to DARC on red blood cells (de Winter et al., 1997), resulting in only a transient rise of plasma CXCL8 and a more prolonged increase of erythrocyte-bound CXCL8. CXCL8 was shown to be transiently elevated in the serum of cancer patients undergoing

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treatment with IL-1 alpha. The erythrocyte-bound CXCL8 levels were higher than those measured in plasma and remained elevated long after the plasma levels had become undetectable (Tilg et al., 1993). Elevated plasma CXCL8 is a diagnostic marker of early-onset bacterial infection in neonates, and a recent study revealed that neonates with suspected bacterial infections had ng/ml levels of CXCL8 bound to DARC on their red blood cells compared with almost undetectable levels in healthy newborns (Orlikowsky et al., 2004). Preeclampsia is characterized by neutrophil activation, and CXCL8 is involved in the pathophysiology. A recent study revealed a correlation between a Duffy-negative phenotype, high plasma levels of CXCL8, and TNF-a in preeclamptic women compared with controls (Velzing-Aarts et al., 2002). The higher CXCL8 levels in preeclampsia may result from increased production and/or reduced clearance, related to a high frequency of a Duffy-negative phenotype. The function of DARC on red blood cells in the long-term maintenance of the levels of free chemokines in plasma has been uncovered by Jima and coworkers ( Jilma-Stohlawetz et al., 2001). Thus DARC functions as a biphasic regulator of chemokines in blood buffering them in acute situation and maintaining their levels on a longer run.

4. Isolation of Erythrocytes and Measurement of Chemokines Whole blood is centrifuged at 2000g, and the plasma is removed and saved. The packed pellet is resuspended in PBS, pH 7.4, and centrifuged over a Ficoll-Hypaque solution adjusted to a density of 1.095 to remove granulocytes, platelets, and mononuclear cells. Erythrocytes prepared in this manner are devoid of contaminating leukocytes and platelets when examined by light microscopy. The red blood cells are then washed three times by centrifugation at 180g in PBS and set aside for extraction of chemokines. To extract chemokines bound to the isolated red blood cells they are lysed by resuspension in 1% Triton X-100 in a volume corresponding to the original blood sample. The lysate is then incubated for 40 min and then stored at 80 C until assayed for chemokines. Chemokines are assayed by ELISA with kits from R and D systems. Typical instructions for measuring CXCL8 levels in plasma and in red cell lysates is taken from the manufacturer’s booklet and is given in the following. Other chemokines such as CCL5 and CCL2 can be measured with the appropriate ELISA kit. 1. Add 100 ml of assay diluent to each well. 2. Add 50 ml of standard, control, or sample per well. Securely cover with a plate sealer and incubate for 2 h at room temperature. Gently tap the plate to ensure thorough mixing.

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3. Aspirate each well and wash, repeating the process three times for a total of four washes. Wash by filling each well with wash buffer (400 ml). Complete removal of liquid at each step is essential to good performance. After the last wash, remove any remaining wash buffer by aspirating or decanting. Invert the plate and blot it against clean paper towels. 4. Add 100 ml of CXCL8 conjugate to all wells. Securely cover with a plate sealer and incubate for 1 h at room temperature. 5. Repeat the aspiration/wash as above. 6. Add 200 ml of substrate solution to each well. Incubate for 30 min at room temperature and protect from light. 7. Add 50 ml of stop solution to each well. The color in the wells should change from blue to yellow. If the color in the wells is green or if color change does not appear uniform, gently tap the plate to ensure thorough mixing. 8. Determine the optical density of each well within 30 min, with a microplate reader set to 450 nm. If wavelength correction is available, set to 540 nm or 570 nm. If wavelength correction is not available, subtract readings at 540 nm or 570 nm from the readings at 450 nm. This subtraction will correct for optical imperfections in the plate. There is very little interference in this assay from either the 1% Triton X-100 in the cell lysate or from plasma samples. The dynamic range of the assay is from 20 to 2000 pg/ml, and all samples should be diluted so that the chemokine concentration lies somewhere in the middle of the standard curve.

5. DARC as a Chemokine Transcytosis Receptor To induce leukocyte emigration, chemokines produced by tissue cells have to negotiate the endothelial cell barrier and associate with the luminal endothelial cell surface (Rot, 1992). The translocation of tissue chemokines is achieved by active endothelial cell transport leading to chemokine immobilization on the tips of the luminal microvilli and their presentation to the adherent leukocytes (Middleton et al., 1997). Until recently, glycosaminoglycans, heparan sulfate in particular, have been postulated to mediate endothelial cell transport and immobilization of chemokines (Handel et al., 2005; Wang et al., 2005). However, early studies of chemokine in situ binding in intact tissues suggested that DARC expressed by the endothelial cells of venules contributes to the chemokine interactions with these cells (Hub and Rot, 1998). Nevertheless the clear demonstration of DARC function in chemokine transcytosis was possible only in vitro. Primary endothelial cells rapidly lose their DARC expression in culture, and in vitro propagated endothelial cell lines do not express this receptor either.


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Therefore, transfected cells were used to uncover in vitro the contribution of DARC to chemokine transcytosis and apical immobilization (Pruenster et al., 2008). Here we detail the transwell-based methods to study DARCmediated chemokine transcytosis and its contribution to chemokineinduced leukocyte transmigration across the cell monolayers expressing this receptor.

6. The Assay of Chemokine Transcytosis by DARC 1. Seed DARC-transfected cells known to form tight polar monolayers (e.g., MDCK or HUVEC) on collagen-coated transwell inserts (pore size 5 mm) and grow in complete medium until confluence. To prevent the cell outgrowth below the filter, it is important to place fluid in the bottom plate only 1 day before the assay. If fluid is placed in the bottom plate simultaneously with seeding the cells, within 3 to 4 days two monolayers will grow in a transwell, one above and one below the filter. 2. Before the assay, measure electrical resistance across the monolayers and select only the wells with comparable monolayers. Anticipate attrition at this point by starting more wells than will be necessary for the assay. Diffusion across the monolayers of FITC-labeled inulin as a tracer (its molecular mass is in the range of chemokines) may confirm the low nonspecific permeability of the monolayers. 3. For transcytosis assay, select 125I-labeled cognate chemokine. Good results were obtained with CXCL8 or CCL2 (specific radioactivity approximately 2000 Ci/mmol). Test for saturability of binding and establish optimal and economical concentration to be used by initially testing different concentrations of chemokine (e.g., from 0.002 to 20 pmol). Use non-cognate chemokine (e.g., CCL19) to test for specificity of binding and transcytosis. Depending on studying basolateral to apical transcytosis or vice versa, place chemokine below or above the monolayer, respectively and incubate for 3 or 4 h as well as overnight at 37 C. Collect fluid from the bottom and top compartments and then recover the cell surface– bound chemokine by the addition of 10 saline for 3 min above the monolayers. The latter fraction is important for studying DARCmediated transcytosis, because a significant part of transported chemokine remains associated with the apical cell membrane. Previously, the fraction of membrane-associated transported chemokine was not accounted for, which explains why the direct contribution of DARC to chemokine transcytosis has not been observed by previous investigators (Lee et al., 2003). Next disrupt the cells with 0.4% Triton-X100 to obtain the intracellular chemokine.

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4. To differentiate between the radioactivity associated with intact and degraded chemokine submit the samples recovered from the different compartments of transwell to precipitation by 12.5% trichloroacetic acid (TCA) at 4 C. 5. Measure in a gamma counter the radioactivity associated with TCAsoluble fractions (degraded chemokine) and TCA-precipitable, insoluble fractions (intact chemokine) for samples obtained from each of the different compartments. The total amount of radioactivity recovered from the transwell system should be close to 100% of the input. With some chemokines that avidly bind to plastic, the amount recovered is less. Chemokine binding to transwell support and bottom well can be tested by measuring the radioactivity associated with these parts. 6. By use of the transcytosis assay as described previously, the contribution of DARC to the unidirectional chemokine transport from basolateral to apical surface of the cell monolayers was uncovered. The functional consequences of DARC-mediated transcytosis to chemokine-induced leukocyte transmigration across the monolayers can be studied as follows. Assemble the transwells as previously, except use inserts with 8-mm pores. On monolayer confluence, add to the bottom plates of transwells either buffer alone or with chemokine at different concentrations. After 1 h of equilibration, label leukocytes with carboxyfluorescein diacetate succinimidyl diester and place 5 105 cells (depending on the chemokine used either neutrophils, monocytes or lymphocytes), into the insert and allow to migrate for 4 h across the cell monolayers either expressing DARC or not. On disassembly, the cells that had migrated across the monolayers and the filters should be counted in the following distinct compartments: (1) in suspension in the bottom well (this is the traditional way of evaluating transmigration); (2) adherent to the bottom plate; as well as (3) adherent to the bottom side of the filter (in both 2 and 3 after being removed with 5 nM EDTA). The enumeration of leukocytes adherent to the bottom side of the filter is important because the expression of DARC by the monolayers disproportionately increases, for not yet clear reasons, the number of the cells in this compartment.

7. Conclusions We have described a number of methods for the experimental study of DARC that should prove to be useful for investigators aiming to analyze the molecular properties of this fascinating protein. Although DARC is an unusual receptor because it does not seem to couple to any of the known intracellular signaling pathways, it has found interest because of its role in malaria pathogenesis together with its functions in regulating chemokine


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levels in blood and their endothelial cell transport. Methods have been presented here that will allow each of these diverse roles of this protein to be further investigated.

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Hetero-Oligomerization of Chemokine Receptors Shirley Appelbe and Graeme Milligan Contents 1. Introduction 2. Coimmunoprecipitation 2.1. Coimmunoprecipitation protocol following heterologous expression of epitope-tagged chemokine receptors 3. Resonance Energy Transfer Techniques 3.1. Bioluminescence resonance energy transfer (BRET) 3.2. Single-point BRET2 protocol 3.3. Data analysis 3.4. Time-resolved FRET 3.5. FRET imaging in living cells 4. Developing Techniques References

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Abstract Although traditionally assumed to be monomeric signaling units, G-protein– coupled receptors (GPCRs) have been shown to exist as dimers/oligomers. Many chemokine receptors have been demonstrated to form homo-oligomers, and hetero-oligomerization between both pairs of chemokine receptors and chemokine receptors and other GPCRs has also been demonstrated. This chapter highlights some of the most common techniques used to investigate chemokine receptor oligomerization.

1. Introduction GPCRs were long assumed to function as monomeric signaling units. However, a large body of evidence now suggests that GPCRs exist as dimers or higher order oligomers (Milligan, 2007). The first conclusive demonstration that receptor oligomers exist in native membranes was Neuroscience and Molecular Pharmacology, University of Glasgow, Glasgow, Scotland, United Kingdom Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05410-X

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Shirley Appelbe and Graeme Milligan

provided by atomic force microscopy images of the organization of rhodopsin in mouse rod outer segment discs (Fotiadis et al., 2003; Liang et al., 2003). These studies clearly demonstrated individual rhodopsin molecules within an oligomeric array of closely packed dimers. As members of the GPCR superfamily, potential oligomerization of various chemokine receptors has been investigated widely. An example of the potential physiologic relevance of chemokine receptor oligomerization was demonstrated following the discovery of a natural genetic mutation of the CCR5 receptor, termed ccr5–32D, which conferred resistance to HIV-1 infection. Individuals homozygous for this mutation were found to be resistant to HIV-1 infection (Liu et al., 1996). Individuals heterozygous for this allele display a delayed progression from infection with HIV-1 to the development of AIDS (Dean et al., 1996; Huang et al., 1996; Micheal et al., 1997; Samson et al., 1996). This delayed progression of infection observed in heterozygous individuals has been hypothesized to result from the homo-oligomerization of ccr5– 32D with wild-type CCR5, resulting in a reduced level of CCR5 expressed at the cell surface (Benkirane et al., 1997). Oligomerization has also been implicated in the delayed progression of HIV-1 infection to AIDS observed in individuals possessing a CCR2 receptor polymorphism CCR2V64I (Smith et al., 1997). Although CCR2 has not been shown to act as a coreceptor for HIV-1, it can form hetero-oligomers with CCR5 and CXCR4 (Mellado et al., 1999), and this is hypothesized to explain the delayed progression of the disease. Demonstration of hetero-oligomerization between both pairs of coexpressed chemokine receptors (Sohy et al., 2007) and the CXCR2 receptor and the DOP opioid receptor (Parenty et al., 2008) have also been instrumental in appreciation of ways in which ligands that lack direct affinity for a GPCR expressed in isolation can produce allosteric effects on that receptor in the presence of a second GPCR for which the ligand does have affinity, if the two GPCRs form a functional hetero-oligomer (Milligan and Smith, 2007; Springael et al., 2007). Several techniques have been developed to investigate GPCR oligomerization. Some of the most useful approaches use energy transfer technology to explore direct protein-protein interactions. Each specific technique in this area yields information on receptor oligomerization that is complementary to the data yielded by the other approaches, and when used in combination, can provide strong evidence of oligomerization.

2. Coimmunoprecipitation Coimmunoprecipitation is a technique that uses antibodies specific for two different forms of a single receptor or a pair of antibodies that identify different receptors that are coexpressed to demonstrate the presence of the

Chemokine Receptor Oligomerization

209

protein targets for the antibodies in the same supramolecular complex. Although unable to distinguish between direct protein-protein interactions involving the target proteins and simply their presence within a larger protein complex, coimmunoprecipitation-based studies have been central features of many studies on GPCR oligomerization and are often the only practical biochemical approach to address this issue in native cells and tissues. The availability of well-characterized anti-chemokine receptor antibodies has allowed more widespread analysis of chemokine receptor heterooligomerization in native cells than for many other GPCRs. Despite this, even in studies on chemokine receptor oligomerization, the use of differentially epitope-tagged forms of the receptors that have been coexpressed in heterologous cell lines, and their coimmunoprecipitation by anti-epitope tag antibodies, has played an important role in exploring the molecular basis and relevance of oligomerization. Coimmunoprecipitation has been applied to demonstrate the homo-oligomerization of a variety of chemokine receptors including CCR2 (Rodriguez-Frade et al., 1999), CCR5 (Vila-Coro et al., 1999), CXCR1 (Wilson et al., 2005), and CXCR2 (Trettel et al., 2003; Wilson et al., 2005). This technique has also been used to demonstrate hetero-oligomerization between pairs of chemokine receptors including CCR2 and CCR5 (Mellado et al., 2001), CCR2 and CXCR4 (Percherancier et al., 2005), and CXCR1 and CXCR2 (Wilson et al., 2005), as well as hetero-interactions between chemokine receptors and other GPCRs, for example CXCR2 and the DOP opioid receptor (Parenty et al., 2008) and CCR5 and the MOP opioid receptor (Chen et al., 2004). Interactions consistent with the presence of chemokine receptor heterooligomers have previously been demonstrated (see Wang and Norcross 2008 for a review). For example, modification of the chemokine receptors CXCR1 and CXCR2 by the addition of Flag and c-Myc epitope tags to the N-terminal of these receptors and their coexpression in HEK293 cells allowed anti-Flag immunoprecipitation. Subsequent separation of the immunoprecipitates and detection with anti-c-Myc revealed immunoreactivity consistent with the presence of a CXCR1/CXCR2 oligomer in the transfected cells (Wilson et al., 2005) as illustrated in Fig. 10.1. An important consideration when coimmunoprecipitation is used is that because of the highly hydrophobic nature of GPCRs, there is a natural tendency for the receptors to aggregate on removal from the plasma membrane. To address this concern it is important to include appropriate controls. These include studies in which lysates from two different cell populations each expressing only one of the GPCRs under examination are mixed before the initial immunoprecipitation step. Figure 10.1 demonstrates that when such controls were performed with cell lysates individually expressing differentially epitope-tagged forms of CXCR1 and CXCR2, there was no indication of interactions between the two receptors, which could only reflect an artefact of the experimental setup.

210


250 160 105 75 a myc

50

IP a flag

35 30

a flag

35 Cell lysates a myc

k

oc

Co

yc

M

C-

Fl

ag

C

M

X C CX R1 -tr an CR sfe 2 ct ed M ix

35

Figure 10.1 Coimmunoprecipitation. HEK293 cells were mock-transfected (mock) or transfected to transiently express Flag-(human) h-CXCR1, c-Myc-h-CXCR2, or both (co-transfected). Samples containing either Flag-h-CXCR1 or c-Myc-h-CXCR2 were also mixed (mix). Confirmation of expression of the appropriate constructs was obtained by immunoblotting cell lysates with either anti-c-Myc or anti-Flag (lower panels). Cell lysates were subsequently immunoprecipitated with anti-Flag. Immunoprecipitated samples were resolved by SDS-PAGE and immunoblotted with anti c-Myc (upper panel) (adapted fromWilson et al. [2005]).

Another important practical consideration in such studies is the centrifugal force used to remove particulate material remaining after cell lysis and membrane solubilization before the immunoprecipitation step. It is vital to centrifuge for a relatively long time and at high centrifugal force. This is done to ensure removal of small membrane fragments that may remain in the ‘‘soluble’’ supernatant fraction that would result in a false-positive result being observed that, rather than oligomerization, might simply reflect the presence of copies of the two monomeric receptors within these membrane fragments. As noted earlier, coimmunoprecipitation may be exploited to investigate chemokine receptor hetero-oligomerization within native tissues if pairs of suitable and well-characterized anti-receptor antibodies are available.

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211

For example, Suzuki et al. (2002) used receptor-specific antibodies to demonstrate the ability of CCR5 to form hetero-oligomers with each of DOP, KOP and MOP opioid receptors in human CEMx174 lymphocytes.

2.1. Coimmunoprecipitation protocol following heterologous expression of epitope-tagged chemokine receptors Seed HEK293T cells into 10 cm2 dishes. When the cells have reached 60 to 70% confluency, transfect the cells with Lipofectamine reagent according to manufacturers’ instructions (Invitrogen, Paisley, U.K.). The differentially epitope-tagged constructs should be both expressed individually and coexpressed and a mock transfection control should also be included. Harvest cells 48 h after transfection and resuspend the cell pellet with 1 ml of 1 RIPA (radioimmune precipitation assay) buffer (100 mM HEPES, pH 7.4, 300 mM sodium chloride, 2% Triton-X 100, 1% sodium deoxycholate, and 0.2% sodium dodecyl sulfate) supplemented with 10 mM NaF, 5 mM EDTA, pH 8, 10 mM NaH2PO4, 5% ethylene glycol, and a Complete EDTA-free protease inhibitor tablet. Place the samples on a rotating wheel at 4 C for 1 h. Centrifuge the samples for 60 min at 100,000g at 4 C and transfer the supernatant to a fresh tube containing 200 ml of 1 RIPA and 50 ml of Protein G (GE Healthcare) to preclear the samples. Incubate the samples at 4 C on a rotating wheel for 1 h. Pellet the Protein G by centrifugation for 10 min at 20,800g, at 4 C. Remove the supernatant into a fresh tube and determine the protein concentration with a bicinhoninic acid (BCA) assay method. This method uses bicinhoninic acid and copper sulphate solutions in which proteins reduce the Cu (II) ions to Cu (I) ions in correlation with protein amount initiating a color change caused by BCA binding to reduced Cu (I). The absorption of the protein samples can be recorded at 562 nm and the concentration calculated by referring to a standard curve. Equalize the protein concentration of the samples to 1 mg/ml with 1 RIPA. At this stage a mixed protein control should be generated in which equal amounts of the samples each expressing a single epitope-tagged chemokine receptor construct should be present. The total protein present in this control should equal the other samples; 600 ml of each sample should be incubated overnight with 40 ml Protein G and an optimal concentration of antibody directed against the epitope of interest at 4 C on a rotating wheel. If the receptors of interest undergo marked N-glycosylation, this may hinder the binding of the antibody and it can be beneficial to include N-glycosidase F to promote deglycosylation in the pulldown step. Reserve 100 ml of the equalized supernatant to investigate protein expression in the cell lysates. Approximately 16 h after incubation, centrifuge samples at 20,800g for 1 min at 4 C and wash the pelleted Protein G

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beads with 500 ml 1 RIPA buffer. Repeat this washing step a further twice before adding 40 ml Laemmli sample buffer. Heat the samples to 85 C for 4 min to elute the proteins. Analyze both immunoprecipitated samples and cell lysates with SDS-PAGE gel electrophoresis.

3. Resonance Energy Transfer Techniques Resonance energy transfer techniques have been used extensively to demonstrate chemokine receptor and other GPCR oligomerization. The main advantage these techniques offer is the ability to demonstrate GPCR interactions in living cells. These techniques exploit the nonradiative transfer of energy between an energy donor and acceptor pair and are based on the Fo¨rster mechanism. This theory states that the energy transfer efficiency is inversely proportional to the distance between donor and acceptor molecules by the sixth power calculated by:

E ¼ 1= 1 þ ½r=Ro 6 ; (Forster, 1948). This calculation demonstrates that the extent of energy transfer between the donor and acceptor moieties is highly dependent on ˚ the proximity of the moieties with the permissive distance being 100 nM. Nonspecific binding or fluorescent ligand dissociation is determined by incubating the cells or beads with a large excess (i.e., 100 to 1000 Kd) of site-saturating nonfluorescent ligand. Alternately, nonspecific binding can be analyzed by incubation of the parental cells, which lack the transfected receptor, or beads, which lack the cognate receptor, in the presence of fluoresceinated peptide alone (Gilbert et al., 1999; Simons et al., 2003c). Affinity measurements can also be quantitatively evaluated by spectrofluorometry, for FPR (Simons et al., 2003a; Sklar et al., 1981, 1985). The determination of FPR ligand affinities can be accomplished through a soluble ligand competition assay as described in Simons et al. (2003a).

6. Modular Molecular Assemblies of GPCR Ternary Complexes on Beads 6.1. Materials Frozen cell membranes (500-ml aliquots), HPSM buffer, syringe with 25-gauge needle, and 25% DOM. The cloning of the FPR, its expression in U937 cells, generation of FPRGai2, and FPR-GFP fusion constructs and membrane preparations have been described elsewhere (Simons et al., 2003a). Wild-type FPR receptors and fusion constructs are over expressed (up to 500,000 receptors/cell) in U937 cells. Expression levels are optimized by sorting on the flow cytometer. The cells are lysed by nitrogen cavitation, and crude postnuclear membrane preparations are stored at 80 C in aliquots of 500 ml, corresponding to 108 cell equivalents.

Subsecond Analysis GPCR Ternary Complex Activation

235

6.2. Solubilization of FPR A typical membrane solubilization preparation involves the following steps: Add 700 ml of HPSM buffer to an aliquot of thawed membranes. Centrifuge sample in a microcentrifuge tube for 15 min and remove supernatant, and resuspend the pellet in 220 ml of HPSM with a syringe attached to a 25-gauge needle. Add 25 ml of 10% dodecyl maltoside, and 2.5 ml of 100 protease inhibitor cocktail (Calbiochem, San Diego, CA) to the membrane suspension, and gently mix for 2 h at 7 C. Remove the unsolubilized material by centrifugation at 135,000g for 15 min, yielding a supernatant of solubilized FPR at 4 108 cell equivalents/ml (5 mg/ml protein), which must be used within 6 h of preparation for best results, otherwise samples can be stored at 80 C. Solubilization may yield up to 100% of receptors harvested from the membranes. It cannot be assumed that DOM allows binding activity to be retained in other GPCR systems, although it has been confirmed for b2-adrenegic receptor. The typical yield of receptors ranges from 100 to 300 nM, depending on receptor expression levels. The initial number of receptors/cell is usually determined by flow cytometry measurements, which are based on the analysis of bound fluorescent ligands used at saturating concentrations or GFP fluorescence, where applicable (cf. section on Standard calibration beads).

6.3. Modular assembly of ternary complexes on beads In cells, the ternary complex of ligand, GPCR, and heterotrimeric G-proteins is expected to have a very short lifetime, which is regulated by the kinetics of nucleotide exchange between the inactive GDP-bound form and the active GTP-bound form (Hamm, 1998, 2001). The exchange is initiated by the contact between the heterotrimeric G-protein and a ligandactivated GPCR. The ternary complex thus survives for the short duration of the dissociative interchange between GDP and GTP under rapid mass transfer considerations where intracellular [GTP] >> [GDP] and can thus more readily replace GDP. Under our experimental conditions, stable LRG assemblies on beads are created in circumstances in which the contact between the GPCR and G beads causes the ejection of GDP from the Ga nucleotide pocket with little to no chance of rebinding, because of the negligible concentrations of soluble nucleotides (GDP or GTPgS), in solution. The nucleotide pocket remains empty until the addition of excess nucleotides, which triggers the disassembly of ternary complexes during the rapid mix experiments.

236

Tione Buranda et al.

7. Preparation of G-Protein Coated Beads (Gabg Beads) 7.1. Materials FLAG tagged subunits (Gb1g2 or Gb4 g 2), Gai2, Gai3, etc, (N.B. the variety of subunits isotypes is only limited by availability), G buffer. We have typically used streptavidin-coated beads (Spherotech), where the typical stock concentration is 40,000 beads/ml (Buranda et al., 2001). The concentration of beads (beads/ml) is determined by counting on a hemocytometer or a flow cytometer. To use a flow cytometer, the rate at which sample is consumed (i.e., flow rate) must be defined. The typical flow rate on our BD flow cytometers is set at 1 ml/sec on ‘‘high-flow’’ setting. Therefore one needs to simply run a sample for a desired time while the cytometers’ ‘‘event count’’ readout function in CellQuest software is turned on. At present, biotin functionalized antiFLAG M2 antibodies (bioM2) purchased from Sigma need to be purified of up to millimolar levels of biotin impurities, by several washes in YM-30 Microcon centrifugal filter devices (Millipore Corp. Bedford, MA) (Buranda et al., 2001). Sample purity can be checked by assaying for biotin in successive filtrate solutions with a simple biotin assay (Wu et al., 2005a). The assay is based on the kinetic analysis of the enhancement of fluorescence of streptavidin/fluorescein biotin complexes in the presence of biotin. The kinetic response of fluorescence enhancement is proportional to the concentration of biotin.

7.2. Preparation of M2 beads We have previously used titration binding curves to show that Spherotech beads are surface-saturated with 4 to 4.5 million bioM2 molecules (Buranda et al., 2001; Wu et al., 2007b). The number of bioM2 antibodies/bead can be assessed by use of a fluorescently labeled FLAG peptide (Kd 8.0 nM ) in a centrifugation assay (Buranda et al., 2001). A centrifugation assay involves the determination of absolute numbers of bead-associated ligands from the analysis of fluorescence of residual supernatant solutions after the removal of beads by centrifugation. Paired samples (ligand binding samples and samples blocked with excess biotin) of beads are allowed to equilibrate before centrifugation. The difference in the fluorescence intensity of residual supernatant solutions of binding samples and blocked samples is used to determine the quantity of bound samples. The solution measurements are, in turn, correlated to a parallel analysis of bead-associated fluorescence by flow cytometry (Buranda et al., 1999). For routine preparations of M2 beads, a simple 50% stoichiometric excess of bioM2 is sufficient to saturate the bead sites because of the very tight multivalent binding of streptavidin and the 3 biotins/bioM2.


237

The M2 beads are washed twice and resuspended in PBS buffer and stored in 25-ml aliquots (40,000 beads/ml) at 4 C until needed.

7.3. Preparation of G (abg) on M2 beads A typical preparation of G-protein–coated beads involves the incubation of an equimolar amount of M2 beads (e.g., a 25-ml aliquot of M2 beads contains 13 pmol FLAG binding sites) and FLAG tagged subunits (b1g2 or b4g2) in 25 ml of G buffer for an hour at 4 C under mild vortex. The beads can then be stored long term at 80 C. Because these experiments allow for the use of various isotypes of Gaij ( j ¼ 1,2,3) subunits, target aij-subunits can be added as needed. It is always useful to determine the surface density of the bg-subunits on each bead to keep track of stability of the beads and for the standardization of measurements over a period of several months. Surface coverage of bg-subunits can be quantitatively determined on the basis of a standard calibration protocol that relies on the fluorescently labeled FLAG peptide described previously (Buranda et al., 2001). Our standard calibration protocols (Wu et al., 2007b) for measuring the number of M2 antibody sites on (Spherotech) 6 mm streptavidin-coated beads has typically yielded 8 to 9 million fluorescent FLAG peptides per bead (Buranda et al., 2001; Simons et al., 2003a) covered by 4 to 4.5 million bivalent anti-FLAG antibodies under surface-saturating conditions. The site density of subunits is then derived from analyzing the difference in the fluorescence intensity of neat anti-FLAG beads (i.e., beads with known maximal surface density of fluorescent FLAG peptide sites) relative to those beads that are partially covered with FLAG tagged bg-subunits with the fluorescent FLAG peptide. In our experience, the functional activity of M2 antibodies drops over time; therefore, over the course of more than 6 months, we have noticed a drop in the surface density of fluorescent FLAG peptide and bg-subunits of 25% (Wu et al., 2007a). The drop in activity of M2 antibodies does not affect the integrity of the ternary complex assembly because it serves as a simple tether of the bg-subunits to the beads. The change in surface coverage only affects the initial amplitude of the signal associated with the ternary complexes at disassembly. Under our normal experimental conditions, the surface coverage of Gbg varied from 45 to 70% of the total available 9 million anti-FLAG–binding sites, depending on the concentration of FLAG tagged Gbg-subunits and the age of the M2 antibody stock (Wu et al., 2007a). The assembly of Ga-subunits onto the Gbg beads can be achieved by mixing Ga with an aliquot of beads in 10 ml G buffer at 4 C for 1 h. The ratio of beads and Ga is governed by the Kd 32 nM (Wu et al., 2007a) of this interaction. The mixture is then centrifuged and resuspended in G buffer at 20,000 beads/ml; this is the empirical standard concentration of beads used for rapid mix flow

238


measurements. It is not necessary to saturate sites with Ga-subunits as the limiting reagent turns out to be GPCR as detailed below.

7.4. Assembly of LRG (abg) on beads The assembly of ligand and receptor (LR) complexes on Gabg-bearing beads is performed as previously described (Simons et al., 2003a). An aliquot of the detergent-solubilized receptor (1% dodecyl maltoside) is mixed with a saturating concentration of the ligand, (Kd 4 nM ) (Bennett et al., 2001; Simons et al., 2003a, 2004; Sklar et al., 2000) and then mixed with the G-protein–complexed beads. For the rapid mix experiments, a typical experimental run uses a volume of 35 ml. The receptor concentration is the limiting reagent, thus one would like to optimize the volume fraction associated with the receptor. The maximum concentration of receptor ever recovered from membrane solubilization preparations was 300 nM. Because the bead component contributes 1 ml (20,000 beads/ml), and the ligand volume can be similarly minimized with a sufficiently high concentration, it is possible to minimize the dilution of the receptor during molecular assembly, steps which are typically done in a minimal volume of 10 ml before increasing the volume to the minimal 35 to 40 ml required for measurement with the rapid mix flow cytometer. As previously noted, the ability of the flow cytometer to discriminate between free and bound fluorophores is limited above 200 nM. It is, therefore, advisable to maintain the concentration of the fluorophore that is in stoichiometric excess (i.e., the fluorescent ligand) below 200 nM. The advantage of using a high-affinity ligand (e.g., Kd of fMLFK-FITC is 4 nM ) (Bennett et al., 2001; Simons et al., 2003a, 2004; Sklar et al., 2000) is realized here. The quantitation of the ternary LRG complexes on beads is carried out with standard calibration beads on the basis of fluorescence readings of fMLFK-FITC or EGFP (vide supra). Although relatively high concentrations of FPR are used, 150 nM, the typical surface coverage (Wu et al., 2005b) on a bead is notably low and depends on the isotypes of the components subunits of a heterotrimer. For example, LRG complexes derived from fMLFK-FITC, wild-type FPR, and Gai3b1g2 yielded 20,000 to 30,000 LRG site occupancies (or 0.7% of 4 106 Gabg sites) on beads compared with >100 nM ) that are cosolubilized with the receptor. A mitigating factor in this assay is that the local concentration of the bead-borne G-proteins is very high relative to the soluble endogenous competitors, thus yielding a practically useful quantity of LRG complexes on beads. Schematics of modularly assembled LRG complexes on beads are summarized in Fig. 11.2 and discussed in the section below.

8. Analysis of Modular Dissassembly of LRG Modules Because LRG complexes can be assembled in modular fashion, it is possible to assess the kinetic and thermodynamic parameters of their interaction at each junction point (Buranda et al., 2007). Disassembly of LRG

240


complexes can be analyzed under circumstances driven by guanine nucleotide activation. Junction points are identified in the schematics displayed in Fig. 11.2. The ternary complex of wild-type FPR is shown in Fig. 11.2A. Guanine nucleotide–mediated disassembly of the ternary complex occurs at three junction points as shown by arrows: (1) ba, (2) aR, and (3) RL. The Gbg heterodimeric subunit is virtually nondissociable under physiologic conditions (Oldham and Hamm, 2007). The junction points are distinguishable by kinetics. To identify the kinetics of each break point we examined the reactivity of isolated component modules of the ternary complex and fusion constructs of FPR-Gai2 (Fig. 11.2C) and FPR-GFP (Fig. 11.2D) as described in the following. Spectrofluorometric and rapid mix flow cytometry can be used to analyze the kinetics of component L, R, and G dissociation from the ternary complexes in solution and beads. To isolate the specific joint associated with a particular kinetic measurement, it is useful to first consider components of the ligand-receptor module separately from the components of the heterotrimeric G-protein unit. With the exception of constitutively active systems, R and G generally remain as separate entities in the absence of ligand. Eqs. (11.3 to 11.7) represent the dissociating modules under consideration. Final results are summarized in Table 11.1.

8.1. Real-time spectrofluorometric measurement of the dissociation of L from R KD

L F þ R Ð LF R ! LF þ R kdiss

ð11:3Þ

The interactions between GPCRs and ligands (agonists or partial agonists) have been quantitatively evaluated (Simons et al., 2003a, 2004) and some Kd values have been tabulated in a review chapter (Buranda et al., 2007). We briefly focus on the measurement of ligand dissociation with a fluorometric anti-FITC antibody assay (Sklar et al., 1981), which has been described in more recent publications (Buranda et al., 1999; Key et al., 2001; Simons et al., 2003a). The polyclonal antibody to fluorescein that is used in our laboratory is not commercially available. However, alternative antifluorescein antibodies such as clone4-4-20, A-6421 (Invitrogen.probes. com), or clone FIT-22 (Biolegend.com) can be purchased. We are not aware of previous characterizations of these antibodies, thus the reader would need to perform dilution series experiments to optimize. To measure kdiss in Eq. (11.3), it is necessary to remove endogenous G-protein from solubilized receptors (Bennett et al., 2001). G-proteins can

241


Table 11.1 Summary of experimental results of LRG dissociation kinetics measured by small-volume rapid-mix flow cytometry Ga

bg

t, fast (sec)

b1g2

18.3 (GTPgS) 33.8 (GDP) 12.4 (GTPgS) 0.8 0.2 (GTPgS) 1.5 0.4 (GDP) 0.7 0.1(GTPgS) 0.8 0.1 (GTPgS) 1.1 (GDP) 0.6 0.05 (GTPgS) 3.1 0.2 (GTPgS) 5.8 (GDP) 2.4 0.2 (GTPgS) 5.2 0.6 (GDP)

Ligand

Receptor

1a

LF

R-Gai2

2a 3b

LF LF

R-Gai2 R

Gai2

b4g2 b1g2

4b 5b

LF L

R RF

Gai2 Gai2

b4g2 b1g2

6b 7b

L LF

RF R

Gai2 Gai3

b4g2 b1g2

8b

L

RF

Gai3

b1g2

Junctionc

LRG LRG LRG LRG LRG LRG LRG L RG

Reproduced from (Wu et al., 2007a) with permission. a Kinetic data analyzed with a single-phase exponential model. b Kinetic data analyzed with a two-phase exponential model. The data only show the results of the analysis of the fast component. The slower component was typically two orders of magnitude or more than the fast component (cf. Figure 11.3 in Wu et al. [2007a]). We have attributed the slow component to receptor misfolding (see text for details). Dissociation produced by GTPgS was always faster and to a greater extent than GDP. c Experimentally measured point of dissociation in the ternary complex.

be removed from solubilized receptors by incubating anti-Gi1,2,3 antibody (Calbiochem or Sigma) for 45 min on ice. The immunocomplex of antibody-substrate is removed by incubating with a slurry of protein A-agarose for 30 min. The sample is centrifuged at 14,000g for 30 sec, and the supernatant is removed. The stoichiometric ratio of anti-Gi1,2,3 antibody, or protein A beads to solubilized receptors is determined by sample size, and typically follows recommended guidelines on product data sheets of the commercial reagents. In a cylindrical cuvette, a fluorescein-labeled ligand (e.g., 10 nM ) to the formyl peptide receptor, fMLFK-FITC (LF) is mixed with a 10-fold stoichiometric excess of detergent-solubilized receptor cleared of endogenous G-proteins (vide supra). At the spectrofluorometer, a baseline reading of the intensity of receptor-bound ligand is taken before a 2-ml aliquot of the antifluorescein antibody is added in situ via a Hamilton syringe through a sample port in thousand fold stoichiometric excess to LFR complexes and free LF in the cuvette. Diffusive encounter between antifluorescein antibodies and LFR or LF selectively quenches 95% of the emission intensity of free LF but not RLF complexes, where the receptor’s steric bulk effectively blocks contact between the fluorescein tag on the short peptide and

242


the antibody (Sklar et al., 1990). Because the antibody can quench dissociating LF, the rate at which LF is quenched represents a direct measure of its dissociation from R: t1/2 20 sec for fMLFK-FITC.

8.2. Real-time spectrofluorometric measurement of the dissociation of L from RG KD

L F R þ G Ð L F RG ! RG þ L F kdiss

ð11:4Þ

To measure the dissociation of LF from FPR in the presence of G-proteins, exogenous G-proteins (e.g., purified Gai3-subunits) are combined with bg in an equimolar ratio (10 mM ) and mixed with solubilized FPR protein (Bennett et al., 2001) and with 10 nM fMLFK-FITC in a volume of 12 ml and incubated for 2 h at 4 C. Before analysis, samples are diluted to 200 ml and equilibrated to room temperature for 2 min. At the spectrofluorometer, anti-FITC-Ab is added to quench the LF as it dissociates from LRG. The dissociation rate of LF from G-protein–coupled R is typically slower than dissociation from uncoupled R: t1/2 100 sec. The decoupling of R and G by in situ addition of 0.1 mM GTPgS yields a rate similar to LR complex in Eq. (11.3) (Bennett et al., 2001).

8.3. Rapid mix measurement of guanine nucleotide–induced disassembly of ternary complexes The basic operation of a rapid mix flow cytometer has been outlined previously, instrumental details, interface with a standard Beckton Dickinson flow cytometer, setup of automation routine, and calibration beads for mixing, have been described elsewhere (Wu et al., 2005b). A typical run consumes a minimum volume of 35 ml. The 1000-ml volume double-barrel syringes of the rapid mix device can, therefore, be loaded with enough sample volume for several sequential runs. For example, for three consecutive runs, 105 ml of LRG beads in DHPSM buffer (600 beads/ml) are loaded into one of the sample syringes of a small-volume rapid mix device (syringe 1 in Fig. 11.1). An equal volume of DHPSM buffer or 0.2 mM guanine nucleotide is loaded into the other sample syringe (syringe 2 in Fig. 11.1). A buffer-only control experiment is used to establish a baseline measurement of the effects of a twofold dilution of the bead suspension. Other controls involve the use of LRG assemblies made in the presence of GTPgS to inhibit the formation of LRG. The experimental run proceeds according to an automated mixing sequence as described elsewhere (Wu et al., 2005b).

243


The time course data are then converted to ASCII format with the FCSQuery program (Bruce Edwards, UNM HSC). Data are analyzed and graphed with commercial software such as Prism software (GraphPad Software, San Diego, CA). To determine the dissociation characteristics, the fluorescence in blocked control samples is subtracted point by point from the time-resolved fluorescence of ligand-binding samples (Wu et al., 2005b, 2007a).

8.4. Real-time rapid mix measurement of the dissociation of Ga from Gbg: disassembly of module in Fig. 11.2B KD

bead gb þ aF Ð bead gb aF

aF

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Numerical Computer Methods, Part E (Methods in Enzymology Vol 384)

Liposomes, Part B (Methods in Enzymology, Volume 372)

Methods in Enzymology, Volume 100: Recombinant DNA, Part B

Methods in Enzymology Vol 440: Nitric Oxide, Part F

Methods in Enzymology Vol 440: Nitric Oxide, Part F

Methods in Enzymology Vol.334, Hyperthermophilic Enzymes (Part C)

Methods in Enzymology Vol 42: Carbohydrate Metabolism, Part C

Methods in Enzymology Vol 441: Nitric Oxide, Part G

Recombinant DNA, Part H, Vol. 217 (Methods in Enzymology)

Methods in Enzymology Vol 131: Enzyme Structure Part L

Methods in Enzymology Vol 284: Lipases, Part A

Methods in Enzymology Vol 39: Hormone Action Part D

Programmed Cell Death Part A (Methods in Enzymology, Vol 442)

Methods in Enzymology 488 - Biothermodynamics, Part 3

Methods in Enzymology 465 - Liposomes, Part G

Methods in Enzymology Vol 387: Liposomes, Part D

Methods in Enzymology Vol 367: Liposomes, Part A

Methods in Enzymology 464 - Liposomes, Part F

Methods in Enzymology 492 - Biothermodynamics, Part 4

Methods in Enzymology 455 - Biothermodynamics, Part 1

Methods in Enzymology Vol.412: Amyloid, prions, and other protein aggregates, Part B

Methods in Enzymology Vol 298: Molecular Motors and the Cytoskeleton, Part B

Methods in Enzymology Vol 307: Confocal Microscopy

Methods in Enzymology Vol 426: Integrins

METHODS IN ENZYMOLOGY - GLYCOBIOLOGY (Vol. 415)

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