Angiogenesis: In Vivo Systems, Part B (Methods in Enzymology, Volume 445)

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 Founding Editors

SIDNEY P. COLOWICK AND NATHAN O. KAPLAN

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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 Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands Copyright # 2008, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2008 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0076-6879/2008 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (+44) 1865 843830, fax: (+44) 1865 853333, E-mail: permissions@elsevier. com. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Support & Contact’’ then ‘‘Copyright and Permission’’ and then “Obtaining Permissions.”

For information on all Elsevier Academic Press publications visit our Web site at elsevierdirect.com ISBN-13: 978-0-12-374314-5 PRINTED IN THE UNITED STATES OF AMERICA 08 09 10 11 9 8 7 6 5 4 3 2 1

CONTRIBUTORS

Wadih Arap The University of Texas M. D. Anderson Cancer Center, Department of Genitourinary Medical Oncology, Houston, Texas Gabriele Bergers Comprehensive Cancer Center, University of California-San Francisco, San Francisco, California, and Department of Neurological Surgery, Brain Tumor Research Center, University of California-San Francisco, San Francisco, California Joyce Bischoff Vascular Biology Program and Department of Surgery, Children’s Hospital, Boston, Harvard Medical School, Boston, Massachusetts Pawel P. Borowicz Center for Nutrition and Pregnancy, and Department of Animal Sciences, North Dakota State University, Fargo, North Dakota Michael Bouvet Department of Surgery, University of California, San Diego, California Weibo Cai Departments of Radiology and Medical Physics, University of Wisconsin, Madison, Wisconsin, and Stanford University School of Medicine, Stanford, California Xiaoyuan Chen Stanford University School of Medicine, Stanford, California Yongping Crawford Genentech, Inc., South San Francisco, California Cheng Cui Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas Michael Detmar Institute of Pharmaceutical Sciences, Swiss Federal Institute of Technology, ETH Zurich, Zurich, Switzerland Xiu Rong Dong Carolina Cardiovascular Biology Center and Department of Medicine, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina xi

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Napoleone Ferrara Genentech, Inc., South San Francisco, California Mike B. Filla Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas Susan J. Fisher Department of Obstetrics, Gynecology and Reproductive Biology, Department of Anatomy, Director, Human Embryonic Stem Cell Program, Faculty Director, Sandler-Moore Mass Spectrometry Core Facility, Institute for Regeneration Medicine, Center for Reproductive Sciences, University of California-San Francisco, San Francisco, California Sanjiv S. Gambhir Stanford University School of Medicine, Stanford, California Noelle M. Griffin Sidney Kimmel Cancer Center, San Diego, California Alexander Groisman Department of Physics, University of California San Diego, La Jolla, California Shireen Hafez Department of Anatomy and Embryology, College of Veterinary Medicine, Alexandria University, Edfina, Elbehera, Egypt Cornelia Halin Institute of Pharmaceutical Sciences, Swiss Federal Institute of Technology, ETH Zurich, Zurich, Switzerland Robert M. Hoffman AntiCancer, Inc., San Diego, California, and Department of Surgery, University of California, San Diego, California Nathan M. Hunkapiller Department of Obstetrics, Gynecology and Reproductive Biology, University of California-San Francisco, San Francisco, California Chrystelle Lamagna Department of Neurological Surgery, Brain Tumor Research Center, University of California-San Francisco, San Francisco, California Klaus Ley La Jolla Institute for Allergy and Immunology, La Jolla, California Charles D. Little Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas

Contributors

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Kan Lu Department of Neurological Surgery, Brain Tumor Research Center, University of California-San Francisco, San Francisco, California Colin T. Maguire Cardiovascular Sciences Graduate Program, Baylor College of Medicine, One Baylor Plaza, Houston, Texas, and Carolina Cardiovascular Biology Center, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina Mark W. Majesky Carolina Cardiovascular Biology Center and Departments of Medicine and Genetics, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina, and Cardiovascular Sciences Graduate Program, Baylor College of Medicine, One Baylor Plaza, Houston Michele McElroy Department of Surgery, University of California, San Diego, California Juan M. Melero-Martin Vascular Biology Program and Department of Surgery, Children’s Hospital, Boston, Harvard Medical School, Boston, Massachusetts Javier Mestas La Jolla Institute for Allergy and Immunology, La Jolla, California Renata Pasqualini The University of Texas M. D. Anderson Cancer Center, Department of Genitourinary Medical Oncology, Houston, Texas Maria K. Pospieszalska La Jolla Institute for Allergy and Immunology, La Jolla, California Dale A. Redmer Center for Nutrition and Pregnancy, and Department of Animal Sciences, North Dakota State University, Fargo, North Dakota Lawrence P. Reynolds Center for Nutrition and Pregnancy, and Department of Animal Sciences, North Dakota State University, Fargo, North Dakota Paul A. Rupp Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas Michael C. Schmid Moores UCSD Cancer Center, University of California-San Diego, La Jolla, California Jan E. Schnitzer Sidney Kimmel Cancer Center, San Diego, California

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Michael Simons Section of Cardiovascular Medicine, Yale University School of Medicine, New Haven, Connecticut Prithu Sundd La Jolla Institute for Allergy and Immunology, La Jolla, California Martin Trepel Department of Oncology and Hematology, University Medical Center HamburgEppendorf, Hamburg, Germany Judith A. Varner Moores UCSD Cancer Center, University of California-San Diego, La Jolla, California San-Pin Wu Department of Molecular and Cellular Biology, Baylor College of Medicine, One Baylor Plaza, Houston, Texas Alexander Zarbock La Jolla Institute for Allergy and Immunology, La Jolla, California, and Department of Anesthesiology and Intensive Care Medicine, University of Mu¨nster, Mu¨nster, Germany

PREFACE A TRIBUTE TO DR. JUDAH FOLKMAN

The field of angiogenesis has recently lost its pioneer and leader, Dr. Judah Folkman. This was a tremendous loss to many of us who knew him and to the field in general. Dr. Folkman inspired a generation of scientists in efforts to translate basic discoveries toward new therapeutics for a wide range of diseases including cancer, blinding eye disease, and inflammatory disease. Due in large part to Dr. Folkman’s efforts and direction, we now have the first generation of therapeutics that disrupt angiogenesis in patients suffering from cancer and macular degeneration. While Dr. Folkman clearly passed away before his time, he did live long enough to observe that many thousands of patients are now better off due to antiangiogenic therapy. I had a rather interesting initiation to the field of antiangiogenesis that was wholly inspired by Dr. Folkman. In the mid-1980s as a junior faculty at the Scripps Research Institute, I was studying what many of us in the field were beginning to appreciate were a family of cell adhesion receptors, later termed ‘‘integrins.’’ I had developed a monoclonal antibody (LM609) to the vitronectin receptor later referred to as integrin anb3. During the course of my work, LM609 was used to stain a variety of diseased and normal tissues. To my surprise, LM609 reacted strongly with blood vessels in tumors and inflammatory sites, but failed to react with blood vessels in normal tissues. After seeing this result, I began to read up on the emerging field of angiogenesis research. It was clear that most of the literature in the field came from Dr. Folkman or one of his disciples. I immediately contacted Dr. Folkman. By the time I finished describing our results, I realized that he was excited as I was about our studies. In fact, before I could ask him any questions, he suggested that I visit his lab to learn the chick chorioallantoic membrane (CAM) assay to determine whether LM609 might have an impact on angiogenesis in a quantitative animal model. Naturally I arranged a trip to the Folkman lab within the next couple of weeks. I had never been to Harvard, and was a bit intimidated by the place. I introduced myself to his administrative assistant, who welcomed me and indicated that Dr. Folkman was expecting me. Within minutes, Dr. Folkman, clad in a lab coat greeted me and suggested that we get started. At this point, I assumed he was going to introduce me to one of his students or technicians who would then proceed to show me the CAM assay step by step. To my surprise, Dr. Folkman led me to a hood, sat down, xv

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and immediately started to instruct me in how to induce angiogenesis on the CAM. In fact, the next thing I knew, I was sitting at the hood next to Dr. Folkman going through the procedure in detail. Therefore, I can say I learned the technique from the master. Ultimately, Dr. Folkman introduced me to several members of the Folkman lab, including Drs. Donald Inber, Pat D’Amore, and Mike Klagsburn. I remember how enthusiastic and communicative all of these folks were. In fact, I am happy to say that I still maintain close contact with them and have had many opportunities over the years to discuss science and reminisce about the past. In fact, Don, Pat, and Mike have all kindly contributed chapters to Methods in Enzymology volumes on angiogenesis. While on the airline flight home from the Folkman lab, I began to realize that my career was about to take a change in course. From that point forward, I began to focus on the role of adhesion receptors in angiogenesis and began to realize that blocking angiogenesis with integrin antagonists could have a very impressive impact on the growth of tumors in mice. Importantly, two of the agents we developed, including humanized LM609, have shown clinical activity in patients with late-stage cancer. Since my initiation to the field, I have since followed Dr. Folkman’s work and have attended dozens of his lectures. Listening to a Folkman lecture is like watching one of your favorite movies—you can watch it over and over again and still find something interesting to think about. It was difficult for anyone to attend his lecture and not come away excited about science in general and angiogenesis in particular. The field of angiogenesis has matured over the past 25 years due in large part to Dr. Folkman’s drive, enthusiasm, perseverance, and kindness. Dr. Folkman’s leadership has helped to recruit many scientists and physicians from the academic and private sectors to focus on new approaches to develop angiogenesis inhibitors. In the early days, there were a limited number of technological approaches to measure and study angiogenesis. The CAM assay was among the first quantitative approaches to measure the growth of newly forming blood vessels. From this humble beginning, the field has exploded and as a result we now have a wide range of techniques, approaches, and animal models designed to monitor and study the growth of new blood vessels in development, tissue remodeling, and disease. These methods are described in detail in this volume by many of the current leaders of the field.

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

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VOLUME 435. Oxygen Biology and Hypoxia Edited by HELMUT SIES AND BERNHARD BRU¨NE VOLUME 436. Globins and Other Nitric Oxide-Reactive Protiens (Part A) Edited by ROBERT K. POOLE VOLUME 437. Globins and Other Nitric Oxide-Reactive Protiens (Part B) Edited by ROBERT K. POOLE VOLUME 438. Small GTPases in Disease (Part A) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 439. Small GTPases in Disease (Part B) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 440. Nitric Oxide, Part F Oxidative and Nitrosative Stress in Redox Regulation of Cell Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 441. Nitric Oxide, Part G Oxidative and Nitrosative Stress in Redox Regulation of Cell Signaling Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 442. Programmed Cell Death, General Principles for Studying Cell Death (Part A) Edited by ROYA KHOSRAVI-FAR, ZAHRA ZAKERI, RICHARD A. LOCKSHIN, AND MAURO PIACENTINI VOLUME 443. Angiogenesis: In Vitro Systems Edited by DAVID A. CHERESH VOLUME 444. Angiogenesis: In Vivo Systems (Part A) Edited by DAVID A. CHERESH VOLUME 445. Angiogenesis: In Vivo Systems (Part B) Edited by DAVID A. CHERESH

C H A P T E R

O N E

Inflammation, Angiogenesis, and Lymphangiogenesis Cornelia Halin and Michael Detmar Contents 1. 2. 3. 4.

Introduction Angiogenesis and Lymphangiogenesis in Inflammation Mouse Models of Inflammation Commonly Used Markers of Blood Vessels and of Lymphatic Vessels 5. Measuring the (Lymph)angiogenic Response by Immunofluorescence 6. Quantitating (Lymph)angiogenesis by Fluorescence-Activated Cell Sorting 7. Measuring Vascular Permeability and Flow 8. Material and Methods 8.1. Mice 8.2. Induction of a DTH response 8.3. Immunofluorescence for MECA-32 and LYVE-1 or CD31 and podoplanin 8.4. Immunofluorescence for Prox1 and CD31 8.5. Immunofluorescent detection of proliferating cells 8.6. Quantification of LEC and BEC numbers by FACS analysis Acknowledgments References

2 3 5 8 10 12 13 14 14 15 15 16 17 18 20 20

Abstract The growth of blood and lymphatic vessels, namely angiogenesis and lymphangiogenesis, is well known to be of importance for tumor growth and metastatic spread. In fact, several therapeutic strategies are currently being pursued in the clinic to interfere with these processes. By contrast, vascular remodeling associated with chronic inflammatory disorders, such as psoriasis, rheumatoid arthritis or inflammatory bowel disease, is only now emerging as a potential new target to treat these conditions. Animal models of inflammation, which

Institute of Pharmaceutical Sciences, Swiss Federal Institute of Technology, ETH Zurich, Zurich, Switzerland Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03001-2

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2008 Elsevier Inc. All rights reserved.

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2

Cornelia Halin and Michael Detmar

mimic the inflammatory and the vascular phenotype of the disease, are important tools for studying inflammation and the accompanying (lymph)angiogenic response in vivo. This review provides a brief summary of our current knowledge of inflammation-induced angiogenesis and lymphangiogenesis and of selected mouse models that can be used to analyze these processes in vivo. It also provides a detailed description of methods, such as immunofluorescence or fluorescence-activated cell sorting (FACS) that can be used to visualize and quantitate vascular remodeling in inflamed tissues.

1. Introduction The importance of angiogenesis for tumor growth, which was first postulated more than 35 years ago, nowadays is a well-established concept (Folkman, 1971, 2006). Angiogenesis, defined as the formation of new blood vessels from pre-existing vessels, also occurs during various physiologic processes, namely during embryogenesis, the female menstrual cycle, the hair growth cycle, and during wound healing (Folkman, 2006). The involvement of angiogenesis in tumor growth has been very well documented, and several antiangiogenic compounds have been investigated as potential therapeutics to treat malignancies (Folkman, 2006; Morabito et al., 2006). The most successful antiangiogenic strategy to date has been the development of a monoclonal antibody directed against vascular endothelial growth factor (VEGF)-A: bevacizumab (Avastin) was approved by the Food and Drug Administration for treatment of colorectal cancer in 2004, and is currently under evaluation for treatment of other types of cancer (Folkman, 2006; Kerr, 2004). Furthermore, many other antiangiogenic strategies, involving small molecules or monoclonal antibodies, are currently in clinical development (Mor et al., 2004; Morabito et al., 2006). It is becoming increasingly recognized that besides angiogenesis, lymphangiogenesis, the formation of new lymphatic vessels, is involved in many pathologic processes (Alitalo et al., 2005; Cao, 2005; Cueni and Detmar, 2006). Lymphatic vessels carry out important physiologic functions: They drain excess bodily fluid from tissues, are involved in intestinal fat absorption, and mediate immune cell migration to draining lymph nodes (LNs). However, in contrast to blood vessels, lymphatic vessel biology is much less well understood: Research in this field has for a long time been hampered by the lack of lymphatic endothelium-specific markers. Only recently have lymphatic endothelium-specific markers (e.g., podoplanin, Prox1 and LYVE-1) been identified and now make it possible to unambiguously detect lymphatic vessels in situ, or to isolate lymphatic endothelial cells for further cultivation and study in vitro (Alitalo et al., 2005; Cao, 2005; Cueni and Detmar, 2006; Hong et al., 2004; Saharinen et al., 2004).

Vascular Remodeling during Inflammation

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Several studies document the association of lymphangiogenesis with tumor progression and metastasis (Alitalo et al., 2005; Cao, 2005; Cueni and Detmar, 2006). In fact, recent results have shown that tumor lymphangiogenesis serves as a prognostic indicator for LN metastasis and survival in human malignancies, including head and neck cancer and melanomas (Beasley et al., 2002; Dadras et al., 2005). In contrast to angiogenesis and lymphangiogenesis associated with tumor growth, vascular remodeling under inflammatory conditions has been less well studied. In the following, we will first briefly review our current knowledge of inflammation-associated angiogenesis and lymphangiogenesis, as well as of animal models that can be used to study these processes in vivo. Furthermore, we will focus on different fluorescencebased methods, which can be used to visualize and quantitate vascular remodeling in vivo in mouse models of inflammation.

2. Angiogenesis and Lymphangiogenesis in Inflammation Inflammation is typically elicited as a response of the body to injury or infection. Furthermore, inflammation may develop and persist in the context of autoimmune diseases, and also during tumor growth. It has for a long time been recognized that acute inflammation induces changes in the vasculature, which significantly contribute to the observed symptoms of the inflammatory response (Pober and Sessa, 2007)—namely, vascular dilation leading to increased blood flow, increase in vascular permeability leading to tissue swelling and edema formation, and an increase in adhesive interactions between endothelial cells and leukocytes in the blood, leading to recruitment of these cells to the site of inflammation. Many inflammatory mediators, such as complement components, histamines, cytokines and chemokines—that are released at the site of inflammation—induce dilation and leakiness of blood vessels. Furthermore, several inflammatory mediators (e.g., TNFa, IL-6 and IL-1b) are known to induce the expression of chemokines and adhesion molecules in endothelial cells ( Jackson et al., 1997; Pober and Sessa, 2007). Besides its involvement in the acute inflammatory response, it is now increasingly recognized that the vasculature also plays an important role in chronic inflammatory disorders ( Jackson et al., 1997; Pober and Sessa, 2007; Szekanecz and Koch, 2007). During chronic inflammation, inflammatory mediators and angiogenic factors remain upregulated in the tissues and are thought to contribute to and exacerbate the inflammatory response. Endothelial cells remain in an activated state, expressing high levels of adhesion molecules and chemokines, thereby facilitating continuous leukocyte

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recruitment. Furthermore, vessels remain enlarged and vascular permeability enhanced, so that blood fluid keeps accumulating at the site of inflammation. Last but not least, in presence of various proangiogenic factors, endothelial cells start to proliferate and initiate a further expansion of the vascular network. However, in contrast to tumor growth, the angiogenic response associated with inflammation mainly consists of a proliferative enlargement of pre-existing vessels, rather than in the sprouting of new vessels ( Jackson et al., 1997). The association of angiogenesis with disease pathology has been well documented for several inflammatory disorders. For example, in rheumatoid arthritis, angiogenesis represents one of the earliest histopathologic findings and is thought to be required for pannus development (Paleolog, 2002). Also in psoriasis, skin lesions are characterized by an angiogenic expansion of the dermal microvasculature (Braverman and Keh-Yen, 1986). Furthermore, recent studies have shown that angiogenesis is crucial during inflammatory bowel disease (IBD) (Chidlow et al., 2007). Increasing evidence also indicates that besides angiogenesis, lymphangiogenesis is involved in certain inflammatory and autoimmune conditions. For example, lymphatic hyperplasia is frequently found in rejected renal transplants (Kerjaschki et al., 2004, 2006) and in psoriatic skin lesions (Kunstfeld et al., 2004). Furthermore, recent studies in mice have shown that bacterial infection of the airways, or inflammation of the skin induce a strong lymphangiogenic response in the affected tissue and in draining LNs (Angeli et al., 2006; Baluk et al., 2005; Halin et al., 2007). The exact role of inflammation-induced lymphangiogenesis is unclear at this point, but it is likely that lymphatic vessel formation and remodeling participate in the regulation of the immune response, by affecting the transport of fluid, antigen and leukocytes to draining LNs (Angeli et al., 2006; Halin and Detmar, 2006). Probably the best-described mediator of inflammation-induced vascular remodeling is vascular endothelial growth factor A (VEGF-A) (Ferrara et al., 2003). Its main receptor, the VEGF receptor (VEGFR)-2, is expressed both on blood vessels as well as on lymphatic vessels. Besides stimulating angiogenesis, VEGF-A exerts several other effects, which directly enhance the inflammatory response in the tissue: Originally identified as vascular permeability factor, VEGF-A increases vascular permeability to allow fluid and plasma proteins, such as fibrinogen and proteins of the complement system, to enter the tissue. Furthermore, VEGF-A is known to be a chemotactic factor for monocytes and macrophages, which both express VEGFR1 (Sawano et al., 2001). Last but not least, VEGA-A may directly induce the expression of adhesion molecules, such as E-selectin, P-selectin, IL-8, VCAM-1 and ICAM-1 in endothelial cells, leading to further recruitment of leukocytes to sites of inflammation (Detmar et al., 1998; Kim et al., 2001; Lee et al., 2002). VEGF-A is produced by various cell types during


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inflammation; these include stromal cells, monocytes and macrophages, keratinocytes, and activated T cells (Mor et al., 2004; Paleolog, 2002). Serum levels of VEGF-A are reportedly elevated in patients with various chronic inflammatory disorders, such as rheumatoid arthritis, IBD and psoriasis (Ballara et al., 2001; Bhushan et al., 1999; Kanazawa et al., 2001). In fact, serum levels of VEGF-A have been shown to correlate with disease severity in patients suffering from various chronic inflammatory conditions (Carvalho et al., 2007). The tight link between the innate immune system, tissue remodeling and angiogenesis is also evidenced by the fact that several inflammatory cytokines have been described to have pro-angiogenic activity. Cytokines, such as TNF-a, TGF-b, IL-1, IL-6, IL-8, and IL-18, may exert a direct proangiogenic effect on the endothelial cells, or else act indirectly by upregulating VEGF-A in the tissue (Cohen et al., 1996; Koch et al., 1992; Leibovich et al., 1987; Park et al., 2001; Pertovaara et al., 1994). Besides VEGF-A, the other major lymphangiogenic factor is VEGF-C. This factor binds to VEGFR-2 and VEGFR-3 expressed on lymphatic endothelial cells and promotes proliferation, migration, and survival of these cells ( Jeltsch et al., 1997; Makinen et al., 2001). VEGF-A and proinflammatory cytokines, such as TNFa, have been shown to induce expression of VEGF-C (Cursiefen et al., 2004; Ristimaki et al., 1998; Skobe and Detmar, 2000). Furthermore, VEGF-C is a chemoattractant for activated macrophages, which express VEGFR-3 (Skobe et al., 2001).

3. Mouse Models of Inflammation Inflammation represents a complex interplay of various cell types secreting a plentitude of factors that may act directly or indirectly on different cellular targets. This complexity makes it difficult to study the effects of the inflammatory environment on endothelial cell proliferation and vascular remodeling in in vitro experiments. Thus, an in-depth analysis of inflammation-induced vascular remodeling either requires the availability of tissue specimens from human patients, or else animal models of chronic inflammation to mimic human disease. Many mouse models of inflammatory disorders have been described, which lend themselves to studying angiogenesis and lymphangiogenesis in vivo. For instance, studies on rheumatoid arthritis-associated angiogenesis have been performed in various mouse models, such as in mice with collagen-induced arthritis (Grosios et al., 2004), in TNFa transgenic mice (Yin et al., 2002) and in transgenic K/BxN mice (De Bandt et al., 2003). Moreover, different mouse models to study the angiogenic response in IBD have been described. Targeted

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mutations in several mouse genes can give rise to IBD ( Jurjus et al., 2004). For example, IL-10–deficient mice spontaneously develop the disease (Kuhn et al., 1993), supporting the hypothesis that a dysregulated immune response to enteric flora can trigger IBD. Furthermore, various methods exist to chemically induce intestinal inflammation in mice (Danese et al., 2007; Jurjus et al., 2004). For example, colitis can be induced by oral administration of dextran sodium sulfate (DSS). The blood vascular network appears expanded in the inflamed intestinal tissue of DSS-treated animals (Fig. 1.1A and B). Furthermore, lymphatics, which typically appear collapsed in the uninflamed tissue, have a wide, dilated lumen in the inflamed tissues, likely due to edema formation and increased fluid drainage (Fig. 1.1A and B). There are several mouse models of the chronic inflammatory skin disorder psoriasis, which mimic the vascular response observed in human psoriasis lesions. Examples include SCID-mice transplanted with human psoriatic skin (Raychaudhuri et al., 2001), conditional JunB/C-Jun knockout mice (Zenz et al., 2005), or K14/VEGF-A transgenic mice (Xia et al., 2003). The latter mouse model was developed by our laboratory and is commonly used by us and many other laboratories. In these mice, murine VEGF-A is continuously expressed in epidermal keratinocytes under the control of the K14 promoter. Homozygous animals spontaneously develop inflammatory skin lesions at about 3 to 6 months of age. Such lesions display the typical disease pathology of psoriasis; massive leukocyte infiltration, hyperproliferation and abnormal differentiation of the epidermal keratinocytes, as well as a dramatic increase in size and number of blood vessels and of lymphatic vessels (Kunstfeld et al., 2004; Xia et al., 2003). In hemizygous animals, which do not spontaneously develop skin lesions, the inflammatory response can be experimentally induced and therefore synchronized between the animals. To elicit chronic inflammation, hemizygous K14/ VEGF-A mice are made allergic toward a contact sensitizer, such as oxazolone, by inducing a delayed-type hypersensitivity (DTH) reaction in the skin. Unlike wildtype mice, hemizygous VEGF-A transgenic mice are unable to downregulate DTH-induced skin inflammation and develop chronic, psoriasis-like inflammatory skin lesions characterized by epidermal hyperproliferation, leukocyte infiltration, and vascular remodeling (Kunstfeld et al., 2004; Xia et al., 2003). Immunofluorescent images of blood and lymphatic vessels found in inflamed and control ears of K14/VEGF-A transgenic mice are shown in Fig. 1.1C and D. An experimental method to induce chronic skin inflammation in wildtype mice consists in eliciting a DTH response toward oxazolone and repeatedly exposing the inflamed skin of the animal to the contact sensitizer (e.g., every 2 to 5 days), in order to maintain an inflammatory response over a longer time (Halin et al., 2007; Kitagaki et al., 1997; Sugiura et al., 2004). If wildtype mice are only challenged once, DTH-induced skin

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B

LYVE-1 & MECA-32 & DAPI

A

CD31 & podoplanin & DAPI


Control

Inflamed

Control

Inflamed

Control

Inflamed

Control

Inflamed

CD31 & podoplanin

C

LYVE-1 & MECA-32

D

CD31 & prox-1 & DAPI

E

Figure 1.1 Analyzing blood vessels and lymphatic vessels in tissues by immunofluorescence. (A and B) Immunofluorescence performed on the colon of control-treated and DSS-treated (inflamed) mice. Oral administration of DSS is a commonly used method for inducing colitis in mice. (C and D) Immunofluorescence performed on sections from control and inflamed ear tissue of K14/VEGF-A transgenic mice (9 days after induction of a DTH response). Chronic skin inflammation induced in these mice strongly resembles human psoriasis. Sections are stained for (A and C) CD31 (pan-endothelial marker, red)

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inflammation is typically cleared within 7 to 10 days. Using such a protocol of repeated DTH challenges, it is possible to elicit a strong inflammatory response in the skin, which is characterized by a profound angiogenic and lymphangiogenic response (Halin et al., 2007).

4. Commonly Used Markers of Blood Vessels and of Lymphatic Vessels One of the most widely used experimental approaches to analyzing (lymph)angiogenesis in inflamed tissue is immunohistology. In particular, immunofluorescence is a very useful technique, since it allows simultaneous analysis and comparison of different markers (i.e., blood vessels and lymphatic vessels) in a single tissue section. Several recent publications provide a good overview of molecules that are specifically expressed by either lymphatic or blood vascular endothelial cells in vitro and in vivo (Cueni and Detmar, 2006; Hirakawa et al., 2003; Saharinen et al., 2004). However, not all of the specific markers lend themselves for routine stainings; for example, receptor tyrosine kinases (e.g., VEGFRs) or soluble factors expressed by a certain cell type typically represent rather difficult targets for immunofluorescence. Even so, the following vascular markers—mainly abundantly expressed adhesion molecules—are available, which produce strong and specific signals in immunofluorescence on mouse tissue sections (Table 1.1). CD31 (PECAM-1, platelet endothelial cell adhesion molecule): CD31, is a 130-kDa integral membrane glycoprotein (member of the immunoglobulin superfamily), which mediates homophilic and heterophilic cell– cell adhesion. It is constitutively expressed on the surface of endothelial cells (panendothelial marker; Fig. 1.1A, C, and E), but is also weakly expressed on many peripheral leukocytes and platelets (Baluk et al., 2007; Ilan and Madri, 2003). Blood vessels stain strongly for CD31, whereas its expression levels are somewhat lower on lymphatic vessels (Ebata et al., 2001). VE-cadherin (CD144): VE-cadherin is an endothelial cell-specific, homophilic adhesion molecule. It is concentrated in interendothelial adherens junctions and is thought to be involved in the maintenance of vascular integrity (Baluk et al., 2007; Breier et al., 1996). It stains blood vessels as well as lymphatic vessels (Baluk et al., 2007). and podoplanin (lymphatic vessel marker, green), or for (B and D) MECA-32 (blood vessel marker, red) and LYVE-1 (lymphatic vessel marker, green). (E) A combination of CD31 (pan-endothelial marker) and Prox1 (nuclear marker of lymphatic endothelial cells) can also be used to differentiate between blood and lymphatic vessels. Scale bars ¼ 50 mm.

Table 1.1 Summary of vascular markers commonly used to identify blood vessels and lymphatic vessels in tissue by immunofluorescence Marker

Name

Panendothelial CD31 (platelet endothelial cell adhesion molecule; PECAM-1) VE-cadherin (vascular endothelial cadherin) Blood vascular MECA-32

CD34

Lymphatic

Prox1 (prospero-related homeobox gene) LYVE-1 Podoplanin

Expression

References

Strongly expressed by blood vessels— weaker staining of lymphatic vessels; also expressed on platelets and some other leukocytes

(Baluk et al., 2007; Ilan and Madri, 2003)

Adhesion molecule in endothelial adherens junctions

(Baluk et al., 2007; Breier et al., 1996)

Marker expressed on most blood vascular endothelial cells in the mouse; named after a monoclonal antibody MECA-32 Sialomucin found in blood vascular endothelial cell membranes— also expressed by hemotopoietic precursor cells

(Halin et al., 2007; Hallmann et al., 1995)

Transcription factor expressed by lymphatic endothelial cells; also present at low levels in other tissues Hyaluronan receptor expressed by lymphatic endothelial cells and activated macrophages Mucin-type transmembrane glycoprotein expressed by lymphatic endothelial cells and various epithelial cells

(Oliver et al., 1993; Wigle and Oliver, 1999)

(Baumheter et al., 1993; Baumhueter et al., 1994)

(Banerji et al., 1999) (Breiteneder-Geleff et al., 1999; Schacht et al., 2003)

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MECA-32: MECA-32 is an antigen recognized by a monoclonal antibody (named MECA-32), which binds to a dimeric transmembrane protein (100 to 110 kD). It is expressed by most blood vascular endothelial cells (Halin et al., 2007; Hallmann et al., 1995) (Fig. 1.1B and D). CD34: The leukocyte antigen CD34 (expressed on hematopoietic progenitor cells) is a sialomucin transmembrane protein, which is also expressed on blood vascular endothelial cells. Depending on its glycosylation, it can serve as a ligand for L-selectin (Baumhueter et al., 1993, 1994). Lymphatic vascular endothelial hyaluronan receptor-1 (LYVE-1): LYVE-1 was identified as a specific cell-surface marker on lymphatic endothelial cells and activated macrophages (Fig. 1.1B and D). As a homologue of the blood vascular endothelium-specific hyaluronan receptor CD44 (Banerji et al., 1999), LYVE-1 is presumably involved in binding hyaluronan metabolites, but its exact function remains unclear to date. A recent report states that inflammatory conditions downregulate LYVE-1 expression in vitro and in vivo ( Johnson et al., 2007). However, in various other studies of chronically inflamed tissue, we and others could not observe such a dramatic loss of LYVE-1 expression (Angeli et al., 2006; Halin et al., 2007; Kajiya et al., 2006; Kunstfeld et al., 2004). Podoplanin: Podoplanin is a mucin-type transmembrane glycoprotein, which is expressed by lymphatic vessels, but not by blood vessels (Breiteneder-Geleff et al., 1999; Schacht et al., 2003) (Fig. 1.1A and C). Furthermore, podoplanin is expressed by various epithelial cells and some tumor cells. It is stably expressed both under uninflamed and inflamed conditions ( Johnson et al., 2007). Interestingly, it has recently been reported that tissue inflammation might induce low-level expression of both podoplanin and LYVE-1 on a subset of dermal blood vessels (Groger et al., 2007). Prox1: The transcription factor Prox1 was discovered as a homolog of the Drosophila homeobox gene prospero (Oliver et al., 1993; Wigle and Oliver, 1999). It is currently thought to be the most specific lineage marker for lymphatic endothelium (Fig. 1.1E).

5. Measuring the (Lymph)angiogenic Response by Immunofluorescence Besides providing a qualitative readout of the inflammatory response, immunostained sections can also be used to quantitate parameters such as vascular density or tissue area covered by vessels. Such a morphometric analysis is typically performed using image analysis software. Commonly used softwares are, for example, IP-LAB (Scanalytics, Fairfax, VA), ImagePro Plus (Leeds Precision Instruments, Minneapolis, MN), or Adobe


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B

LYVE-1 & Ki67 & DAPI

A

LYVE-1 & BrdU & DAPI

Photoshop CS3 (Adobe Systems, San Jose, CA). Typically, several different fields are arbitrarily chosen in each section, and the blood or lymphatic vessel density (i.e., number of vessels per square millimeter), the average vessel size, or the relative tissue area occupied by vessels, is measured with the help of the software. Sometimes fields of views may not be assigned arbitrarily, but hot spots with high vessel density may be chosen in the sections taken from each experimental condition. By combining immunofluorescent analysis of MECA-32 (blood vessels) with LYVE-1 (lymphatic vessels), or else simultaneously staining for CD31 (all vessels) and podoplanin (lymphatic vessels), it is possible to analyze inflammation-induced changes in both vessel types in the same section (Fig. 1.1 A through D). Besides quantification of vessel size and number, another parameter, which can easily be determined by immunofluorescence, is the number of proliferating endothelial cells in the tissue. Two different methods are generally used to detect proliferating cells in tissue sections, namely, staining for BrdU or for Ki67. Bromodeoxyuridine (5-bromo-2-deoxyuridine, BrdU) is a synthetic analogue of thymidine. Typically, BrdU is administered systemically to the experimental animal several hours prior to sacrifice. During this time, BrdU will be incorporated into newly synthesized DNA in replicating cells, such as proliferating endothelial cells, where it can later be detected by staining tissues sections with a BrdU-specific antibody (Fig. 1.2A). Another

Figure 1.2 Analyzing endothelial cell proliferation in tissues by immunofluorescence. Immunofluorescent analysis of proliferating cells in control and inflamed ear tissue of K14/VEGF-A transgenic mice. Immunofluorescence of LYVE-1 (lymphatic vessel marker, green) is shown in combination with (A) a BrdU staining (red) or (B) a Ki67 staining (red), two commonly used stainings to identify proliferating cells. Scale bars ¼ 25 mm.

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widely used technique is to detect proliferating cells by Ki67. The Ki67 antigen is a nuclear protein that is expressed by proliferating cells in all phases of the active cell cycle, and therefore can be used as an endogenous marker to detect proliferating cells in the tissue (Fig. 1.2B).

6. Quantitating (Lymph)angiogenesis by Fluorescence-Activated Cell Sorting Morphometric analysis represents a good method for analyzing the overall, inflammation-induced changes in the vasculature. However, it is difficult to asses by this method, to which extent the expansion of the vascular network is due to actual endothelial cell proliferation, or rather due to dilatation and enlargement of existing vessels. Furthermore, the result of the analysis may vary between different tissue regions from which sections were prepared. We have recently described a method involving quantitative fluorescence-activated cell sorting (FACS) analysis, which bypasses some of these difficulties, by analyzing total endothelial cell numbers in tissue single cell suspensions (Halin et al., 2007). Using this method, it is possible to quantify the (lymph)angiogenic response in the entire tissue. For example, we have successfully used this method to quantify the (lymph)angiogenic response in DTH-inflamed ears and ear-draining auricular LNs of either K14/VEGF-A transgenic or wildtype mice. To this end, inflamed and control ears and LNs are first digested enzymatically and passed through a cell strainer to generate tissue single cell suspensions (Fig. 1.3). FACS analysis is then performed on single cell suspensions, staining for the leukocyte marker CD45, the panendothelial cell marker CD31 and the lymphatic endothelial cell marker podoplanin. This combination of markers makes it possible to differentiate between leukocytes (CD45þCD31–podoplanin–), blood vascular endothelial cells (BECs) (CD45–CD31þpodoplanin–) and lymphatic endothelial cells (LECs) (CD45–CD31þpodoplaninþ) (Fig. 1.3). Using this approach, we found that chronic inflammation induced a significant increase in the number of BECs and LECs in inflamed ears. Interestingly, tissue inflammation specifically induced lymphangiogenesis, but not angiogenesis in the ear-draining auricular LN (Fig. 1.4), an observation that could also be confirmed by immunofluorescent analysis of the tissue sections (Halin et al., 2007). Furthermore, our studies demonstrated that these processes are mediated by VEGF-A, which becomes expressed in the inflamed tissue, from where it drains via the afferent lymphatics into the draining LN (Halin et al., 2007).

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(I) Harvest ears and eardraining lymph nodes

(II) Enzymatically digest the tissue

(III) Pass through cell strainer to generate tissue single-cell suspension

(IV) Stain cells for FACS analysis: BECs 1.48

104

CD31

CD31

LECs

105 75.5

105

103 14.3

102 0 0 102

103

CD45

104

104 103 102 0

81.8

105

24.7

0

0 0 102

103

104

105

Podoplanin

Figure 1.3 Measuring the (lymph)angiogenic response in ear skin and ear-draining lymph nodes by quantitative FACS analysis. Schematic representation of the experimental work flow. (I) The tissue of interest is harvested and digested in PBS containing 0.4% collagenase IV. (II) Tissue fragments are passed through a cell strainer, to generate tissue single cell suspensions. (III) Cells are stained for CD45, CD31 and podoplanin.This combination of markers makes it possible to differentiate between leukocytes (CD45þCD31^podoplanin^), blood vascular endothelial cells (CD45þCD31þpodoplanin^) and lymphatic endothelial cells (CD45þCD31þpodoplaninþ) by FACS analysis.

7. Measuring Vascular Permeability and Flow One prominent sign of inflammation is increased vascular permeability, leading to edema formation (Pober and Sessa, 2007). A commonly used method for measuring leakage of fluid from blood vessels is to intravenously inject the dye Evans blue into anesthetized mice and to measure its extravasation into the inflamed or control tissue, either by visual inspection or

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Inflamed skin

Normal skin

Lymphatic vessels Blood vessels

Afferent lymphatics

Lymph node B cell follicle Subcapsular sinus

Efferent lymphatics

Inflammation-induced vascular remodeling: Skin:

Angiogenesis and lymphangiogenesis

Skin-draining lymph node:

Lymphangiogenesis only

Figure 1.4 Schematic representation of inflammation-induced tissue (lymph)angiogenesis and LN lymphangiogenesis. We have recently observed that DTH-induced chronic skin inflammation induces both angiogenesis as well as lymphangiogenesis in the inflamed tissue. Furthermore, a profound lymphangiogenic response occurs in the LN, which drains the inflamed tissue.

photometrically (Lange-Asschenfeldt et al., 2002; Thurston et al., 1998). For photometrical quantification, the dye-injected and anesthetized animals typically are perfused prior to sacrifice, to remove excess dye from within the vasculature. Evans blue is subsequently extracted from the excised tissue and its concentration in the tissue determined with a spectrophotometer. Several protocols of this procedure can be found in the literature (LangeAsschenfeldt et al., 2002; Thurston et al., 1998). Besides measuring permeability of blood vessels, several techniques are currently pursued to study lymphatic flow by intravital microscopy. Most of these methods rely on injecting ink or fluorescent probes into the tissue, to visualize its flow through lymphatic vessels (Harrell et al., 2007; Nagy et al., 2002; Padera et al., 2002).

8. Material and Methods 8.1. Mice K14/VEGF-A transgenic mice overexpress mouse VEGF-A164 in the epidermis under control of the human keratin 14 promoter (Detmar et al., 1998). Homozygous K-14-VEGF-A transgenic mice spontaneously


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develop psoriasis-like inflammatory skin lesions at around 3 to 6 months of age (Xia et al., 2003). For experiments, typically hemizygous mice are used, in which chronic skin inflammation first has to be experimentally induced, by eliciting a DTH response to oxazolone (Halin et al., 2007; Kunstfeld et al., 2004). Alternatively, chronic skin inflammation may be established in wildtype mice by inducing a DTH response toward oxazolone in the ear skin and maintaining the inflammatory response by continuously reapplying oxazolone onto the inflamed ear skin (e.g., every 2 to 5 days) (see protocol in following section). Several protocols for inducing DSS-colitis in mice can be found in the literature (e.g., Wirtz et al., 2007).

8.2. Induction of a DTH response To induce a DTH response in the ear skin of a mouse, the animal is first anesthetized by intraperitoneal injection of saline-containing medetomidine (0.2 mg/kg) and ketamine (80 mg/kg). Once anesthetized, the abdomen of the animal is shaved, using a small electric shaver (Provet, Lyssach, Switzerland). A 2% oxazolone (4-ethoxymethylene-2 phenyl-2-oxazoline-5-one; Sigma, St. Louis, MO) solution in acetone:olive oil (4:1 ratio of volumes) is freshly prepared and applied to the shaved abdomen (50 ml dispersed on an area of approximately 2 3 cm) and to the inner side of each paw (5 ml). Five days after sensitization (day 0), the animal is anesthetized and the ears are challenged by topical application of 10 ml of a 1% oxazolone solution in acetone:olive oil (4:1 ratio of volumes) to the inner and the outer side of the ear. In experiments with wildtype mice, the animals are subjected to repeated challenges with 1% oxazolone on the ears every 2 to 3 days. Inflamed and control ears are typically analyzed 1 or 2 days (to analyze acute inflammation) or more than 9 days (to analyze chronic inflammation) after the first challenge.

8.3. Immunofluorescence for MECA-32 and LYVE-1 or CD31 and podoplanin The mouse is sacrificed and the ears and ear-draining auricular LNs are harvested. Tissues are embedded in optimal cutting temperature (OCT) compound (Sakkura Finetek, Torrance, CA) and frozen on dry ice. Eightmicrometer cryostat sections are cut, transferred to SuperfrostPlus glass slides (Menzel, Braunschweig, Germany), and stored at –80 C until needed. For immunofluorescent staining, slides are taken out of the freezer and, once thawed, are fixed by immersion into –20 C cold acetone (2 min), followed by incubation at 4 C in 80% MeOH (5 min). Next, sections are washed for 5 min in PBS at room temperature (RT), and the

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tissue on the glass slide is surrounded with a thick line from a waterrepellant pen. Sections are then blocked for 1 h in phosphate buffered saline (PBS) containing 12% bovine serum albumin (12% BSA-PBS; 50–150 ml added to each encircled tissue, at RT). All incubation steps are carried out in a light-tight staining chamber that is humidified with wet paper towels. After the blocking step, the sections are briefly washed in PBS and then incubated overnight with the primary antibodies in 12% BSA-PBS at 4 C. The next day, the sections are washed three times with PBS (at RT, 5 to 10 min each) on a rotating table. Sections are then incubated for 1 h at RT with fluorescent Alexa488- or Alexa594-coupled secondary antibodies in 12% BSA-PBS. Subsequently, sections are washed three times for 5 to 10 min in PBS, followed by a 10-min incubation in 10 mM Tris-HCl, pH 8.8. (The latter step is optional, but may help to reduce background fluorescence.) Finally, sections are covered with Vectashield containing DAPI (for visualizing nuclei, Vector Laboratories, Burlingame, CA) and coverslipped. Images are acquired in the individual fluorescent channels. Adobe Photoshop (Adobe Systems) is used to adjust image levels and for merging the images acquired in the different fluorescent channels (image overlay). The following antibodies have been tested with this protocol: Primary antibodies

Rabbit–anti-mouse LYVE-1 (Angiobio, Del Mar), 2 mg/ml Rat–anti-mouse MECA-32 (BD Biosciences, San Jose, CA), 0.15 mg/ml Rat–anti-mouse CD31 (clone MEC13.3, BD Biosciences), 0.25 mg/ml Hamster–anti-mouse podoplanin (clone 8.1.1, Developmental Studies Hybridoma Bank, University of Iowa), 1:600 dilution of hybridoma concentrate, approximately 1.6 mg/ml

Secondary antibodies

Alexa488-conjugated anti-rabbit IgG (Molecular Probes/Invitrogen), 10 mg/ml Alexa488-conjugated anti-hamster IgG (Molecular Probes/Invitrogen), 10 mg/ml Alexa594- conjugated anti-rat IgG (Molecular Probes/Invitrogen), 10 mg/ml

8.4. Immunofluorescence for Prox1 and CD31 Eight-micrometer cryostat sections of the tissue are prepared and stored as described above. For immunofluorescent staining, slides are taken out of the freezer and are briefly rehydrated in PBS (2 min). Sections are surrounded with a thick line from a water-repellant pen and are then fixed by incubating for 20 min at RT in 4% paraformaldehyde (PFA). Slides are washed during


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5 min in PBS and blocked for 1 h at RT in PBS containing 10% goat serum and 0.01% triton-X100 (blocking solution). After blocking, sections are briefly washed in PBS and then incubated overnight in blocking solution, in presence of the following primary antibodies:

Rabbit anti-human/mouse Prox1 (Covance, Princeton, NJ) 1:200 dilution of antiserum Rat–anti-mouse CD31 (clone MEC13.3, BD Biosciences), 0.25 mg/ml

The next day, the sections are washed three times with PBS (at RT, 5 min each), and then incubated for 1 h at RT with goat anti-rabbit Alexa594 and goat anti-rat Alexa488 (both from Molecular Probes/Invitrogen, 10 mg/ml) in blocking solution. Subsequent steps are performed as described above.

8.5. Immunofluorescent detection of proliferating cells 8.5.1. BrdU staining For BrdU incorporation, mice are injected intraperitoneally with BrdU (Sigma, St. Louis, MO, 50 mg/kg) and sacrificed 2.5 h later. Tissues are harvested and processed as described above. After fixation in acetone and MeOH (as described above), sections are hydrated in PBS (5 min), and then incubated for 20 min with 2N HCl, a crucial step to make BrdU-labeled DNA in the nucleus accessible for antibody staining (all steps at RT). After another wash with PBS, sections are blocked for 1 h in 12% BSA-PBS at RT. Incubation with the primary antibodies—rabbit–anti-mouse LYVE-1 (Angiobio, Del Mar, 2 mg/ml); rat–anti-mouse MECA-32 (BD Biosciences: 0.15 mg/ml); rat–anti-mouse CD31 (clone MEC13.3, BD Biosciences, 0.25 mg/ml)—is performed for 2 h at RT. After the washing steps (described above), the secondary, Alexa488-conjugated secondary antibody is added in 12% BSA-PBS, together with the Alexa594-labeled mouse–antiBrdU antibody (Molecular Probes/Invitrogen, cat. A21304, diluted 1:50). Incubation is carried out at RT for 2 h. Subsequent steps are again as described above. 8.5.2. Ki67 staining Ki67 staining requires a stronger fixation of the tissue than the previously described staining procedures. Thus, once the animal is sacrificed, tissues are harvested and are fixed for 4 h in 4% PFA at 4 C. Subsequently, the tissue is frozen in OCT and 8-mm cryosections are prepared (which can be stored at –80 C until needed). For immunostaining, sections are thawed and briefly immersed in PBS to remove OCT. Then, an antigen-retrieval step is performed, using the following buffer: 4.5 ml of citric acid (0.1 M),

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20.5 ml trisodium citrate (0.1 M), and 225 ml H2O. Sections are microwaved in this buffer, first 5 min at high and then 5 min at low energy. Subsequently, sections are washed for 5 min in PBS, and then blocked for 1 h at RT in 12% BSA-PBS. Staining with primary antibodies is carried out overnight at 4 C, using rat anti-mouse Ki67 (TEC-3 clone, from DAKO Cytomation, 1:75 dilution of culture supernatant), and rabbit–anti-mouse LYVE-1 (Upstate, Del Mar, 1:500 dilution) in 12% BSA-PBS. Since most antibodies against blood vascular markers are also rat antibodies, we have not attempted to combine such stainings with the one for Ki67. However, at least for the panendothelial marker CD31, also antibodies raised in other species (e.g., goat) are commercially available and reportedly work in immunofluorescence (Magnusson et al., 2004). After overnight incubation with the primary antibodies, sections are washed as described above. Incubation with Alexa488 and Alexa594 conjugated secondary antibodies (in 12% BSA-PBS) are carried out at RT for 1 h. Subsequent steps are again as described above.

8.6. Quantification of LEC and BEC numbers by FACS analysis The mouse is sacrificed and its two ears and two ear-draining auricular LNs are collected (see also Fig. 1.3). The ears are split into two halves by tearing the tissue along the central cartilage backbone. Ear parts and LNs are then separately digested in 4 ml PBS containing 0.4% collagenase IV (Invitrogen, Basel, Switzerland). Digestions are carried out for 45 min at 37 C, in 15-ml falcon tubes that are attached to an overhead rotator. All subsequent processing steps are performed on ice and using ice-cold solutions, to maintain cell viability. 8.6.1. Preparation of LN single-cell suspensions A 40-mm cell strainer (BD Biosciences) is placed onto a 50-ml falcon tube and the digested LNs (in 4 ml digestion solution) are passed through the strainer, using the plunger of a 1-ml syringe for grinding. This process is repeated twice by transferring the 4-ml digestion solution from the falcon tube back onto the cell strainer. LN single-cell suspensions are subsequently transferred back into the 15-ml falcon tube, and cell numbers are counted, using a hematocytometer and trypan blue, to determine the absolute numbers of viable cells. Finally, the falcon tube is filled with PBS containing 1% FBS (1% FBS-PBS) and is centrifuged for 5 min at 260 g. The supernatant is aspirated and the cell pellet is resuspended in 0.3 to 1 ml of 1% FBS-PBS for FACS staining (small volume for control LNs, larger volume for inflamed LNs).


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8.6.2. Preparation of ear single-cell suspensions After digestion with collagenase IV, the ear halves and the digestion solution are transferred to a 6-cm tissue culture plate, where the ear tissue is disaggregated into small pieces, using forceps and a scalpel for scraping. Subsequently, a fresh 40-mm cell strainer is placed onto a 50-ml falcon tube. The tissue homogenate (4 ml) is passed through the strainer, again using the plunger of a 1-ml syringe for grinding. Subsequently, the filter is rinsed with approximately 10 ml of 1% FBS-PBS, while continuing grinding the residual tissue on the strainer with the syringe plunger. The entire volume is transferred to a 15-ml tube, and centrifuged for 5 min at 260 g. The pellet is resuspended in 1% FBS-PBS, and the cell solution passed once again through a fresh cell strainer. After another centrifugation step, the pellet is resuspended in 1% FBS-PBS (approximately 0.5 ml for control ears and 1.0 ml for inflamed ears). 8.6.3. FACS staining For FACS analysis, 100 ml of ear and LN cell suspensions are transferred to individual wells of a U-bottom–shaped, 96-well plate. All antibody-staining steps are performed on ice in a total volume of 200 ml of 1% FBS-PBS per well. A 20-min incubation is performed using the following primary antibodies:

Rat–anti-mouse CD31 (clone MEC13.3), 0.25 mg/ml Hamster–anti-mouse podoplanin (clone 8.1.1), 1:300 dilution of hybridoma concentrate, approximately 3.3 mg/ml

The plate is then centrifuged for 2 min at 260 g and the supernatant discarded (by rapidly emptying the plate over a bucket—the cells will remain in the well). Wells are washed by adding 200 ml of 1% PBS-FBS and are re-centrifuged. Subsequently, the following fluorescently labeled antibodies are added:

Goat–anti-hamster-PE (Caltag, Invitrogen), 0.8 mg/ml Goat–anti-rat-APC (Caltag, Invitrogen), 0.5 mg/ml Mouse–anti-mouse CD45.1-FITC or mouse–anti-mouse CD45.2-FITC (clone A20 and clone 104, respectively; BD Biosciences), 2.5 mg/ml

Cells are incubated for 20 min with secondary antibodies on ice in the dark. CD45, the common leukocyte antigen, is expressed in different isoforms, which differ in their expression between different mouse strains. Thus, the choice of antibody will depend on the mouse strain used: e.g., FVB mice are CD45.1, whereas C57BL/6 mice are CD45.2.

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The abovementioned mouse–anti-mouse CD45.1/2 antibodies are an alternative to other anti-CD45 antibodies, which typically are generated in rat and therefore would not be compatible with a triple staining using rat– anti-mouse CD31. After 20 min of incubation with the secondary, fluorescently labeled antibodies, cells are spun down, washed once with 1% FBS-PBS and resuspended in the same buffer for FACS analysis (300 to 500 ml). In our experiments, FACS analysis is performed on a BD FACSCanto (Beckton Dickinson, Basel, Switzerland) using FACSDiva software. Data are analyzed with Flowjo software (Treestar, Ashland, TN). For quantification of total BEC (CD45–CD31þpodoplanin–) and LEC (CD45–CD31þpodoplaninþ) number in LN samples (Fig. 1.3), the fraction of LECs and BECs among all live cells (gated) is determined and multiplied by the total number of LN cells that were counted prior to staining (see above). For quantification of LECs and BECs in ear samples, stained samples are acquired for a fixed time interval (e.g., 45 or 90 s). By knowing the acquisition parameters of the FACS machine, notably the sample volume analyzed per time interval (e.g., 100 ml/60 s), it is possible to calculate back the total number of LECs and BECs present in the entire ear single-cell suspension.

ACKNOWLEDGMENTS We would like to thank Nicola Harris and Ben Marsand, ETH Zurich, Switzerland, for providing OCT-embedded intestinal tissue blocks from DSS-treated and control-treated mice. Furthermore, we thank Jana Zielinski and Ingrid van Mier for excellent technical assistance. Research in the authors’ laboratories is supported by the Swiss National Fund (grant 310000-116128 to C.H. and 3100A0-108207 to M.D.), the Prof. Dr. Max Cloe¨tta Foundation (C.H.), the National Institutes of Health grant CA69184 (M.D.) and Commission of the European Communities grant LSHC-CT-2005-518178 (M.D.).

REFERENCES Alitalo, K., Tammela, T., and Petrova, T. V. (2005). Lymphangiogenesis in development and human disease. Nature 438, 946–953. Angeli, V., Ginhoux, F., Llodra, J., Quemeneur, L., Frenette, P. S., Skobe, M., Jessberger, R., Merad, M., and Randolph, G. J. (2006). B cell–driven lymphangiogenesis in inflamed lymph nodes enhances dendritic cell mobilization. Immunity 24, 203–215. Ballara, S., Taylor, P. C., Reusch, P., Marme, D., Feldmann, M., Maini, R. N., and Paleolog, E. M. (2001). Raised serum vascular endothelial growth factor levels are associated with destructive change in inflammatory arthritis. Arthritis Rheum. 44, 2055–2064. Baluk, P., Fuxe, J., Hashizume, H., Romano, T., Lashnits, E., Butz, S., Vestweber, D., Corada, M., Molendini, C., Dejana, E., and McDonald, D. M. (2007). Functionally specialized junctions between endothelial cells of lymphatic vessels. J. Exp. Med. 204, 2349–2362.


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Baluk, P., Tammela, T., Ator, E., Lyubynska, N., Achen, M. G., Hicklin, D. J., Jeltsch, M., Petrova, T. V., Pytowski, B., Stacker, S. A., Yla-Herttuala, S., Jackson, D. G., Alitalo, K., and McDonald, D. M. (2005). Pathogenesis of persistent lymphatic vessel hyperplasia in chronic airway inflammation. J. Clin. Invest. 115, 247–257. Banerji, S., Ni, J., Wang, S. X., Clasper, S., Su, J., Tammi, R., Jones, M., and Jackson, D. G. (1999). LYVE-1, a new homologue of the CD44 glycoprotein, is a lymph-specific receptor for hyaluronan. J. Cell Biol. 144, 789–801. Baumhueter, S., Singer, M. S., Henzel, W., Hemmerich, S., Renz, M., Rosen, S. D., and Lasky, L. A. (1993). Binding of L-selectin to the vascular sialomucin CD34. Science 262, 436–438. Baumhueter, S., Dybdal, N., Kyle, C., and Lasky, L. A. (1994). Global vascular expression of murine CD34, a sialomucin-like endothelial ligand for L-selectin. Blood 84, 2554–2565. Beasley, N. J., Prevo, R., Banerji, S., Leek, R. D., Moore, J., van Trappen, P., Cox, G., Harris, A. L., and Jackson, D. G. (2002). Intratumoral lymphangiogenesis and lymph node metastasis in head and neck cancer. Cancer Res. 62, 1315–1320. Bhushan, M., McLaughlin, B., Weiss, J. B., and Griffiths, C. E. (1999). Levels of endothelial cell stimulating angiogenesis factor and vascular endothelial growth factor are elevated in psoriasis. Br. J. Dermatol. 141, 1054–1060. Braverman, I. M., and Keh-Yen, A. (1986). Three-dimensional reconstruction of endothelial cell gaps in psoriatic vessels and their morphologic identity with gaps produced by the intradermal injection of histamine. J. Invest. Dermatol. 86, 577–581. Breier, G., Breviario, F., Caveda, L., Berthier, R., Schnurch, H., Gotsch, U., Vestweber, D., Risau, W., and Dejana, E. (1996). Molecular cloning and expression of murine vascular endothelial-cadherin in early stage development of cardiovascular system. Blood 87, 630–641. Breiteneder-Geleff, S., Soleiman, A., Kowalski, H., Horvat, R., Amann, G., Kriehuber, E., Diem, K., Weninger, W., Tschachler, E., Alitalo, K., and Kerjaschki, D. (1999). Angiosarcomas express mixed endothelial phenotypes of blood and lymphatic capillaries: Podoplanin as a specific marker for lymphatic endothelium. Am. J. Pathol. 154, 385–394. Cao, Y. (2005). Opinion: Emerging mechanisms of tumour lymphangiogenesis and lymphatic metastasis. Nat. Rev. Cancer 5, 735–743. Carvalho, J. F., Blank, M., and Shoenfeld, Y. (2007). Vascular endothelial growth factor (VEGF) in autoimmune diseases. J. Clin. Immunol. 27, 246–256. Chidlow, J. H., Shukla, D., Grisham, M. B., and Kevil, C. G. (2007). Pathogenic angiogenesis in IBD and experimental colitis: New ideas and therapeutic avenues. Am. J. Physiol. Am. J. Physiol. Gastrointest. Liver Physiol. 293, G5–G18. Cohen, T., Nahari, D., Cerem, L. W., Neufeld, G., and Levi, B. Z. (1996). Interleukin 6 induces the expression of vascular endothelial growth factor. J. Biol. Chem. 271, 736–741. Cueni, L. N., and Detmar, M. (2006). New insights into the molecular control of the lymphatic vascular system and its role in disease. J. Invest. Dermatol. 126, 2167–2177. Cursiefen, C., Chen, L., Borges, L. P., Jackson, D., Cao, J., Radziejewski, C., D’Amore, P. A., Dana, M. R., Wiegand, S. J., and Streilein, J. W. (2004). VEGF-A stimulates lymphangiogenesis and hemangiogenesis in inflammatory neovascularization via macrophage recruitment. J. Clin. Invest. 113, 1040–1050. Dadras, S. S., Lange-Asschenfeldt, B., Velasco, P., Nguyen, L., Vora, A., Muzikansky, A., Jahnke, K., Hauschild, A., Hirakawa, S., Mihm, M. C., and Detmar, M. (2005). Tumor lymphangiogenesis predicts melanoma metastasis to sentinel lymph nodes. Mod. Pathol. 18, 1232–1242. Danese, S., Sans, M., Spencer, D. M., Beck, I., Donate, F., Plunkett, M. L., de la Motte, C., Redline, R., Shaw, D. E., Levine, A. D., Mazar, A. P., and Fiocchi, C. (2007). Angiogenesis blockade as a new therapeutic approach to experimental colitis. Gut 56, 855–862.

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De Bandt, M., Ben Mahdi, M. H., Ollivier, V., Grossin, M., Dupuis, M., Gaudry, M., Bohlen, P., Lipson, K. E., Rice, A., Wu, Y., Gougerot-Pocidalo, M. A., and Pasquier, C. (2003). Blockade of vascular endothelial growth factor receptor I (VEGFRI), but not VEGF-RII, suppresses joint destruction in the K/BxN model of rheumatoid arthritis. J. Immunol. 171, 4853–4859. Detmar, M., Brown, L. F., Schon, M. P., Elicker, B. M., Velasco, P., Richard, L., Fukumura, D., Monsky, W., Claffey, K. P., and Jain, R. K. (1998). Increased microvascular density and enhanced leukocyte rolling and adhesion in the skin of VEGF transgenic mice. J. Invest. Dermatol. 111, 1–6. Ebata, N., Sawa, Y., Nodasaka, Y., Yamaoka, Y., Yoshida, S., and Totsuka, Y. (2001). Immunoelectron microscopic study of PECAM-1 expression on lymphatic endothelium of the human tongue. Tissue Cell 33, 211–218. Ferrara, N., Gerber, H. P., and LeCouter, J. (2003). The biology of VEGF and its receptors. Nat. Med. 9, 669–676. Folkman, J. (1971). Tumor angiogenesis: Therapeutic implications. N. Engl. J. Med. 285, 1182–1186. Folkman, J. (2006). Angiogenesis. Annu. Rev. Med. 57, 1–18. Groger, M., Niederleithner, H., Kerjaschki, D., and Petzelbauer, P. (2007). A previously unknown dermal blood vessel phenotype in skin inflammation. J. Invest. Dermatol. 127, 2893–2900. Grosios, K., Wood, J., Esser, R., Raychaudhuri, A., and Dawson, J. (2004). Angiogenesis inhibition by the novel VEGF receptor tyrosine kinase inhibitor, PTK787/ZK222584, causes significant anti-arthritic effects in models of rheumatoid arthritis. Inflamm. Res. 53, 133–142. Halin, C., and Detmar, M. (2006). An unexpected connection: Lymph node lymphangiogenesis and dendritic cell migration. Immunity 24, 129–131. Halin, C., Tobler, N. E., Vigl, B., Brown, L. F., and Detmar, M. (2007). VEGF-A produced by chronically inflamed tissue induces lymphangiogenesis in draining lymph nodes. Blood 110, 3158–3167. Hallmann, R., Mayer, D. N., Berg, E. L., Broermann, R., and Butcher, E. C. (1995). Novel mouse endothelial cell surface marker is suppressed during differentiation of the blood– brain barrier. Dev. Dyn. 202, 325–332. Harrell, M. I., Iritani, B. M., and Ruddell, A. (2007). Tumor-induced sentinel lymph node lymphangiogenesis and increased lymph flow precede melanoma metastasis. Am. J. Pathol. 170, 774–786. Hirakawa, S., Hong, Y. K., Harvey, N., Schacht, V., Matsuda, K., Libermann, T., and Detmar, M. (2003). Identification of vascular lineage–specific genes by transcriptional profiling of isolated blood vascular and lymphatic endothelial cells. Am. J. Pathol. 162, 575–586. Hong, Y. K., Shin, J. W., and Detmar, M. (2004). Development of the lymphatic vascular system: A mystery unravels. Dev. Dyn. 231, 462–473. Ilan, N., and Madri, J. A. (2003). PECAM-1: Old friend, new partners. Curr. Opin. Cell Biol. 15, 515–524. Jackson, J. R., Seed, M. P., Kircher, C. H., Willoughby, D. A., and Winkler, J. D. (1997). The codependence of angiogenesis and chronic inflammation. FASEB J. 11, 457–465. Jeltsch, M., Kaipainen, A., Joukov, V., Meng, X., Lakso, M., Rauvala, H., Swartz, M., Fukumura, D., Jain, R. K., and Alitalo, K. (1997). Hyperplasia of lymphatic vessels in VEGF-C transgenic mice. Science 276, 1423–1425. Johnson, L. A., Prevo, R., Clasper, S., and Jackson, D. G. (2007). Inflammation-induced uptake and degradation of the lymphatic endothelial hyaluronan receptor LYVE-1. J. Biol. Chem. 282, 33671–33680.


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Jurjus, A. R., Khoury, N. N., and Reimund, J. M. (2004). Animal models of inflammatory bowel disease. J. Pharmacol. Toxicol. Methods 50, 81–92. Kajiya, K., Hirakawa, S., and Detmar, M. (2006). Vascular endothelial growth factor-A mediates ultraviolet B–induced impairment of lymphatic vessel function. Am. J. Pathol. 169, 1496–1503. Kanazawa, S., Tsunoda, T., Onuma, E., Majima, T., Kagiyama, M., and Kikuchi, K. (2001). VEGF, basic-FGF, and TGF-beta in Crohn’s disease and ulcerative colitis: A novel mechanism of chronic intestinal inflammation. Am. J. Gastroenterol. 96, 822–828. Kerjaschki, D., Huttary, N., Raab, I., Regele, H., Bojarski-Nagy, K., Bartel, G., Krober, S. M., Greinix, H., Rosenmaier, A., Karlhofer, F., Wick, N., and Mazal, P. R. (2006). Lymphatic endothelial progenitor cells contribute to de novo lymphangiogenesis in human renal transplants. Nat. Med. 12, 230–234. Kerjaschki, D., Regele, H. M., Moosberger, I., Nagy-Bojarski, K., Watschinger, B., Soleiman, A., Birner, P., Krieger, S., Hovorka, A., Silberhumer, G., Laakkonen, P., Petrova, T., et al. (2004). Lymphatic neoangiogenesis in human kidney transplants is associated with immunologically active lymphocytic infiltrates. J. Am Soc. Nephrol. 15, 603–612. Kerr, D. J. (2004). Targeting angiogenesis in cancer: Clinical development of bevacizumab. Nat. Clin. Pract. Oncol. 1, 39–43. Kim, I., Moon, S. O., Kim, S. H., Kim, H. J., Koh, Y. S., and Koh, G. Y. (2001). Vascular endothelial growth factor expression of intercellular adhesion molecule 1 (ICAM-1), vascular cell adhesion molecule 1 (VCAM-1), and E-selectin through nuclear factorkappa B activation in endothelial cells. J. Biol. Chem. 276, 7614–7620. Kitagaki, H., Ono, N., Hayakawa, K., Kitazawa, T., Watanabe, K., and Shiohara, T. (1997). Repeated elicitation of contact hypersensitivity induces a shift in cutaneous cytokine milieu from a T helper cell type 1 to a T helper cell type 2 profile. J. Immunol. 159, 2484–2491. Koch, A. E., Polverini, P. J., Kunkel, S. L., Harlow, L. A., DiPietro, L. A., Elner, V. M., Elner, S. G., and Strieter, R. M. (1992). Interleukin-8 as a macrophage-derived mediator of angiogenesis. Science 258, 1798–1801. Kuhn, R., Lohler, J., Rennick, D., Rajewsky, K., and Muller, W. (1993). Interleukin-10– deficient mice develop chronic enterocolitis. Cell 75, 263–274. Kunstfeld, R., Hirakawa, S., Hong, Y. K., Schacht, V., Lange-Asschenfeldt, B., Velasco, P., Lin, C., Fiebiger, E., Wei, X., Wu, Y., Hicklin, D., Bohlen, P., and Detmar, M. (2004). Induction of cutaneous delayed-type hypersensitivity reactions in VEGF-A transgenic mice results in chronic skin inflammation associated with persistent lymphatic hyperplasia. Blood 104, 1048–1057. Lange-Asschenfeldt, B., Weninger, W., Velasco, P., Kyriakides, T. R., von Andrian, U. H., Bornstein, P., and Detmar, M. (2002). Increased and prolonged inflammation and angiogenesis in delayed-type hypersensitivity reactions elicited in the skin of thrombospondin-2–deficient mice. Blood 99, 538–545. Lee, T. H., Avraham, H., Lee, S. H., and Avraham, S. (2002). Vascular endothelial growth factor modulates neutrophil transendothelial migration via up-regulation of interleukin8 in human brain microvascular endothelial cells. J. Biol. Chem. 277, 10445–10451. Leibovich, S. J., Polverini, P. J., Shepard, H. M., Wiseman, D. M., Shively, V., and Nuseir, N. (1987). Macrophage-induced angiogenesis is mediated by tumour necrosis factor-alpha. Nature 329, 630–632. Magnusson, P., Rolny, C., Jakobsson, L., Wikner, C., Wu, Y., Hicklin, D. J., and ClaessonWelsh, L. (2004). Deregulation of Flk-1/vascular endothelial growth factor receptor-2 in fibroblast growth factor receptor-1–deficient vascular stem cell development. J. Cell Sci. 117, 1513–1523. Makinen, T., Veikkola, T., Mustjoki, S., Karpanen, T., Catimel, B., Nice, E. C., Wise, L., Mercer, A., Kowalski, H., Kerjaschki, D., Stacker, S. A., Achen, M. G., and Alitalo, K.

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(2001). Isolated lymphatic endothelial cells transduce growth, survival and migratory signals via the VEGF-C/D receptor VEGFR-3. EMBO J. 20, 4762–4773. Mor, F., Quintana, F. J., and Cohen, I. R. (2004). Angiogenesis-inflammation cross-talk: vascular endothelial growth factor is secreted by activated T cells and induces Th1 polarization. J. Immunol. 172, 4618–4623. Morabito, A., De Maio, E., Di Maio, M., Normanno, N., and Perrone, F. (2006). Tyrosine kinase inhibitors of vascular endothelial growth factor receptors in clinical trials: Current status and future directions. Oncologist 11, 753–764. Nagy, J. A., Vasile, E., Feng, D., Sundberg, C., Brown, L. F., Detmar, M. J., Lawitts, J. A., Benjamin, L., Tan, X., Manseau, E. J., Dvorak, A. M., and Dvorak, H. F. (2002). Vascular permeability factor/vascular endothelial growth factor induces lymphangiogenesis as well as angiogenesis. J. Exp. Med. 196, 1497–1506. Oliver, G., Sosa-Pineda, B., Geisendorf, S., Spana, E. P., Doe, C. Q., and Gruss, P. (1993). Prox 1, a prospero-related homeobox gene expressed during mouse development. Mech. Dev. 44, 3–16. Padera, T. P., Stoll, B. R., So, P. T., and Jain, R. K. (2002). Conventional and high-speed intravital multiphoton laser scanning microscopy of microvasculature, lymphatics, and leukocyte-endothelial interactions. Mol. Imaging 1, 9–15. Paleolog, E. M. (2002). Angiogenesis in rheumatoid arthritis. Arthritis Res. 4(Suppl 3), S81–S90. Park, C. C., Morel, J. C., Amin, M. A., Connors, M. A., Harlow, L. A., and Koch, A. E. (2001). Evidence of IL-18 as a novel angiogenic mediator. J. Immunol. 167, 1644–1653. Pertovaara, L., Kaipainen, A., Mustonen, T., Orpana, A., Ferrara, N., Saksela, O., and Alitalo, K. (1994). Vascular endothelial growth factor is induced in response to transforming growth factor-beta in fibroblastic and epithelial cells. J. Biol. Chem. 269, 6271–6274. Pober, J. S., and Sessa, W. C. (2007). Evolving functions of endothelial cells in inflammation. Nat. Rev. Immunol. 7, 803–815. Raychaudhuri, S. P., Dutt, S., Raychaudhuri, S. K., Sanyal, M., and Farber, E. M. (2001). Severe combined immunodeficiency mouse-human skin chimeras: A unique animal model for the study of psoriasis and cutaneous inflammation. Br. J. Dermatol. 144, 931–939. Ristimaki, A., Narko, K., Enholm, B., Joukov, V., and Alitalo, K. (1998). Proinflammatory cytokines regulate expression of the lymphatic endothelial mitogen vascular endothelial growth factor-C. J. Biol. Chem. 273, 8413–8418. Saharinen, P., Tammela, T., Karkkainen, M. J., and Alitalo, K. (2004). Lymphatic vasculature: development, molecular regulation and role in tumor metastasis and inflammation. Trends Immunol. 25, 387–395. Sawano, A., Iwai, S., Sakurai, Y., Ito, M., Shitara, K., Nakahata, T., and Shibuya, M. (2001). Flt-1, vascular endothelial growth factor receptor 1, is a novel cell surface marker for the lineage of monocyte-macrophages in humans. Blood 97, 785–791. Schacht, V., Ramirez, M. I., Hong, Y. K., Hirakawa, S., Feng, D., Harvey, N., Williams, M., Dvorak, A. M., Dvorak, H. F., Oliver, G., and Detmar, M. (2003). T1alpha/podoplanin deficiency disrupts normal lymphatic vasculature formation and causes lymphedema. EMBO J. 22, 3546–3556. Skobe, M., and Detmar, M. (2000). Structure, function, and molecular control of the skin lymphatic system. J. Invest. Dermatol. Symp. Proc. 5, 14–19. Skobe, M., Hamberg, L. M., Hawighorst, T., Schirner, M., Wolf, G. L., Alitalo, K., and Detmar, M. (2001). Concurrent induction of lymphangiogenesis, angiogenesis, and macrophage recruitment by vascular endothelial growth factor-C in melanoma. Am. J. Pathol. 159, 893–903.


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Sugiura, T., Oka, S., Gokoh, M., Kishimoto, S., and Waku, K. (2004). New perspectives in the studies on endocannabinoid and cannabis: 2-arachidonoylglycerol as a possible novel mediator of inflammation. J. Pharmacol. Sci. 96, 367–375. Szekanecz, Z., and Koch, A. E. (2007). Mechanisms of disease: Angiogenesis in inflammatory diseases. Nat. Clin. Pract. Rheumatol. 3, 635–643. Thurston, G., Murphy, T. J., Baluk, P., Lindsey, J. R., and McDonald, D. M. (1998). Angiogenesis in mice with chronic airway inflammation: Strain-dependent differences. Am. J. Pathol. 153, 1099–1112. Wigle, J. T., and Oliver, G. (1999). Prox1 function is required for the development of the murine lymphatic system. Cell 98, 769–778. Wirtz, S., Neufert, C., Weigmann, B., and Neurath, M. F. (2007). Chemically induced mouse models of intestinal inflammation. Nat. Protoc. 2, 541–546. Xia, Y. P., Li, B., Hylton, D., Detmar, M., Yancopoulos, G. D., and Rudge, J. S. (2003). Transgenic delivery of VEGF to mouse skin leads to an inflammatory condition resembling human psoriasis. Blood 102, 161–168. Yin, G., Liu, W., An, P., Li, P., Ding, I., Planelles, V., Schwarz, E. M., and Min, W. (2002). Endostatin gene transfer inhibits joint angiogenesis and pannus formation in inflammatory arthritis. Mol. Ther. 5, 547–554. Zenz, R., Eferl, R., Kenner, L., Florin, L., Hummerich, L., Mehic, D., Scheuch, H., Angel, P., Tschachler, E., and Wagner, E. F. (2005). Psoriasis-like skin disease and arthritis caused by inducible epidermal deletion of Jun proteins. Nature 437, 369–375.

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Color-Coded Fluorescent Mouse Models of Cancer Cell Interactions with Blood Vessels and Lymphatics Michele McElroy,* Michael Bouvet,* and Robert M. Hoffman*,† Contents 1. 2. 3. 4. 5.

Introduction Mouse Models: Fluorescent Tumors to Image Angiogenesis Color-Coded Tumor-Host Models: GFP Nude Mouse Color-Coded Tumor-Host Models: Nestin-Driven–GFP Nude Mouse Imaging Cancer Cell Trafficking in Lymphatics: Experimental Metastasis Model 6. Imaging Cancer Cell Trafficking in Lymphatics: Spontaneous Metastasis Model 7. Protocol: Production of Fluorescent-Protein–Expressing Cancer Cell Lines 7.1. GFP retrovirus production 7.2. RFP retrovirus production 7.3. Production of the histone H2B-GFP vector 7.4. GFP or RFP transduction of tumor cell lines 7.5. Double RFP and histone H2B-GFP gene transduction of cancer cells 7.6. Protocol: Conjugation of anti LYVE-1 antibody 7.7. Protocol: Establishment of imageable tumor models—cell injection 7.8. Surgical orthotopic implantation 7.9. Experimental lymphatic metastasis model 7.10. Spontaneous lymphatic metastasis model 8. Protocol: Imaging of Cancer Cell Interactions with Blood Vessels and Lymphatics 8.1. Whole-body imaging of tumor vasculature 8.2. Skin flap elevation for improved imaging of superficial tumor vasculature for lymphatic migration

* {

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Department of Surgery, University of California, San Diego, California AntiCancer, Inc., San Diego, California

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03002-4

#


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Michele McElroy et al.

8.3. Intravital imaging for deep tumors 8.4. Tumor tissue sampling for highest-resolution imaging of tumor vasculature 9. Protocol: Imaging Methods for Mice Expressing Fluorescent Proteins 9.1. Imaging with a handheld flashlight 9.2. Imaging with fluorescence microscopy 9.3. Imaging with the Olympus OV-100 small-animal imaging system 9.4. Imaging with the Olympus IV-100 scanning laser microscope system 9.5. Imaging using spectral separation 10. Summary and Conclusions References

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Abstract Several new strategies now exist for imaging cancer cell interactions with both blood vessels and lymphatics in living animals. Tumors labeled with fluorescent proteins allow the nonluminous capillaries and larger blood vessels to be clearly visualized against the bright tumor fluorescence via either intravital or wholebody imaging. Signal attenuation by overlying tissue can be markedly reduced by opening a reversible skin flap in the light path, increasing detection sensitivity. With this increase in observable depth of tissue, many previously obscured small tumor vessels can be imaged. In addition, dual-color fluorescence imaging, effected by using red fluorescent protein (RFP)–expressing tumors growing in green fluorescent protein (GFP)–expressing transgenic mice, can show with great clarity tumor–stroma interactions, including the developing tumor vasculature. The GFP-expressing host vasculature, both mature and nascent, can be distinguished from the RFP-expressing tumor itself in this model. Transgenic mice with GFP gene expression driven by the nestin promoter offer another way to image the developing tumor vasculature. In this model system, only nascent blood vessels express GFP, allowing newly developing blood vessels to be imaged against a background of RFP-expressing tumor cells. Finally, dual-color imaging technology can facilitate the imaging of cancer cell interactions with lymphatics. Delivery of FITC-dextran or fluorescent antibodies specific for lymphatic endothelium to the lymphatics around an RFP-expressing tumor allows imaging of tumor cell shedding into the lymphatic system. This imaging technology has the potential to visualize each step of tumor progress.

1. Introduction Traditional models for the visualization of angiogenesis have included the chorioallantoic membrane assay (Auerbach et al., 1974; Crum et al., 1985), the iris neovascularization model (Miller et al., 1993), the disc

Fluorescent Mouse Models of Cancer Cell Interactions

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angiogenesis assay (Passaniti et al., 1992), and corneal models (Alessandri et al., 1983; Deutsch and Hughes, 1979; Epstein et al., 1990; Korey et al., 1977; Mahoney and Waterbury, 1985). While these models have facilitated the understanding of some of the mechanisms of blood vessel induction, they are cumbersome and are typically not used for the evaluation of tumor neovascularization. As such, they are poorly suited to drug discovery research in the arena of anticancer therapeutics. More appropriate models for studying tumor angiogenesis have been developed in rodents, including subcutaneous tumor xenograft models. In the past, subcutaneous tumor xenograft models in mice involved cumbersome histologic and immunohistochemical analysis. Measurements in this type of model system are hampered by sampling bias and require sacrifice of the animal at the time of tumor evaluation. This system cannot therefore be used for ongoing angiogenesis studies in a live tumor-bearing animal, and ultimately the subcutaneous location of the tumor precludes accurate recapitulation of human disease. The rat corneal tumor (Fournier et al., 1981; Gimbrone et al., 1974; Muthukkaruppan and Auerbach, 1979) and rodent skin-fold, windowchamber models (Al-Mehdi et al., 2000; Dewhirst et al., 1984; Fukumura et al., 1998; Papenfuss et al., 1979) have also been used to evaluate tumor angiogenesis. Both of these models allow the study of tumor vasculature in living animals, but quantification of tumor blood vessels requires very specialized procedures and the location of the tumors is quite different from their natural environment. A key limitation of both of these models, including the subcutaneous-transplant model, is that the tumors do not metastasize (Cowen et al., 1995). This could be due to a number of reasons, one of which may be differences in tumor angiogenesis in an ectopic site. Newer techniques for evaluating the interaction between tumor and developing or established blood vessels in living animals rely on fluorescence imaging to visualize cancer cells and host blood vessels. Several such models now exist and allow for longitudinal imaging and evaluation of tumor blood vessels and lymphatics in the same animal. These models allow for imaging of tumors in both orthotopic and ectopic sites, and do not necessarily require sacrifice of the animal at the time of tumor evaluation.

2. Mouse Models: Fluorescent Tumors to Image Angiogenesis Fluorescent proteins have been very useful for imaging of tumors in living animals, and can also be used to allow visualization of tumor blood vessels (Hoffman, 2002, 2005). GFP fluorescence in tumor tissue growing in a mouse model can be detected by either whole-body or intravital

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fluorescence imaging. Using these fluorescence imaging techniques, functional blood vessels appear dark or nonluminous against the background of the fluorescent tumor cells. The high-resolution imaging now available for many fluorescence imaging systems allows measurement of total vessel length, allowing quantitative optical imaging of in vivo angiogenesis. Tumor growth, vascularization and metastasis can be imaged in real time using this strategy (Yang et al., 2001). Surgical orthotopic implantation (SOI) allows for development of tumor models in the most appropriate microenvironment and most closely replicates true human disease (Hoffman, 1999). These orthotopically-growing tumors can metastasize, giving rise to metastatic implants that resemble the clinical behavior of the original human tumor both in sites of metastasis and frequency of occurrence (Hoffman, 1999). Tumor cells transduced and selected for high expression of fluorescent proteins implanted orthotopically can thus be used to visualize both primary and metastatic tumor blood vessels (Yang et al., 2001). It has been suggested that metastatic tumor angiogenesis may be very different than that of the primary tumor, and may require very different interventions (Li et al., 2000). Fluorescent orthotopic tumor models allow testing of antiangiogenic therapeutics on both primary tumor and spontaneous metastases. Whole body imaging of angiogenesis has been demonstrated in mouse models of breast cancer. The MDA-MB-435 human breast cancer cell line was stably transduced to express green fluorescent protein (GFP). MDA-MB435-GFP tumor cells were then implanted orthotopically into the mammary fat pad of nude mice. Tumor growth and developing tumor vasculature could be imaged noninvasively via whole-body fluorescence imaging. Quantitative analysis of developing tumor blood vessels demonstrated increasing microvessel density over 20 weeks (Yang et al., 2001). While this technique allows noninvasive longitudinal imaging of the same tumor in the same animal over time, the overlying skin attenuates and scatters the fluorescence signal, making very small capillaries difficult to image. The issue of skin attenuation of fluorescence signal can be addressed by using a surgically elevated reversible skin flap. By removing the skin from the light path, the observable depth of tissue and the detection sensitivity for small blood vessels can be increased many fold (Yang et al., 2002). The orthotopically-implanted human pancreatic tumor BxPC3-GFP was externally visualized after elevation of a skin flap via fluorescence microscopy and functional microvessels were noted based on their dark contrast against the brilliant green of the pancreatic tumor (Fig. 2.1). Intravital imaging can also be used to improve detection sensitivity for small functional capillaries in fluorescent orthotopic models of pancreatic cancer. Angiogenesis associated with either primary tumor or metastatic growths can be easily imaged through its nonluminous contrast to the surrounding fluorescent tumor tissue. The BxPC3-GFP human pancreatic

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tumor has been imaged in living animals using this strategy (Yang et al., 2001). Because this intravital imaging does not require animal sacrifice, the same animal can be imaged over time. Sequential imaging of the human prostate cancer cell line PC-3-GFP via fluorescence intravital imaging showed progression of blood vessel ingrowth into the developing tumor from days 7 to 20 (Yang et al., 2001). Intravital imaging of primary and metastatic tumor tissue acquired at day 50 after tumor implantation reveal clear delineation of the superficial vasculature (Fig. 2.2). A

B

200 µm

Figure 2.1 Direct view of microvessels of orthotopically growing pancreatic cancer. (A) The human BxPC-3-GFP pancreatic tumor, microvessels, and metastasis to the spleen were directly viewed via a skin-flap window over the abdominal wall of a nude mouse at day 58 after SOI. (B) Microvessels were directly viewed and highly resolved through the skin-flap window at higher magnification. Bar ¼ 200 mm. (FromYang, M., Baranov, E., Wang, J. W., Jiang, P., Wang, X., Sun, F. X., Bouvet, M., Moossa, A. R., Penman, S., and Hoffman, R. M. (2002). Direct external imaging of nascent cancer, tumor progression, angiogenesis, and metastasis on internal organs in the fluorescent orthotopic model. Proc. Natl. Acad. Sci. USA 99, 3824^3829.)

A

B

Day 50

Day 50

Figure 2.2 Intravital fluorescence imaging of Bx-PC-3-GFP human pancreas cancer angiogenesis. (A) Orthotopic tumor. (B) Metastatic lesion in spleen. The GFPexpressing human tumor was transplanted to nude mice by SOI and intravitally imaged 50 days later. Bar ¼ 200 mm. (FromYang, M., Baranov, E., Li, X. M.,Wang, J.W., Jiang, P., Li, L., Moossa, A. R., and Penman, S., Hoffman, R. M. (2001). Whole-body and intravital optical imaging of angiogenesis in orthotopically implanted tumors. Proc. Natl. Acad. Sci. USA 98, 2616^2621.)

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3. Color-Coded Tumor-Host Models: GFP Nude Mouse Mice expressing GFP under the control of the chicken beta-actin promoter and cytomegalovirus promoter were first described by Okabe and colleagues in 1997. In these animals, all tissues with the exception of hair and erythrocytes fluoresce green. Tumor cells stably transduced to express red fluorescent protein (RFP) can be transplanted into these GFPexpressing animals and be reliably distinguished from host tissues using dualcolor fluorescence imaging (Hoffman, 2002; Yang et al., 2003). The GFP mouse was crossed with nu/nu athymic mouse to generate athymic GFP offspring. These mice are bright green under fluorescence imaging and express GFP in essentially all tissues with the exception of erythrocytes. In the adult animal, all organ systems evaluated, including the entire respiratory, digestive and reproductive tracts, express GFP. These animals have a similar lifespan when compared to non-GFP athymic mice (Yang et al., 2004). Previous work by Duda et al. (2004) has described the implantation of nonfluorescent tumors into GFP mice. The fluorescent host stromal and endothelial cells were able to be distinguished from tumor cells in this model by the presence or absence of green fluorescence (Fig. 2.3). Dualcolor fluorescence imaging of RFP-expressing tumors in GFP-expressing transgenic mice also offers the advantage of allowing the measurement of tumor growth over time via whole-body imaging. Tumors can be followed through time in a living animal with whole-body imaging, allowing longitudinal evaluation of tumor growth and metastasis in addition to facilitating the evaluation of tumor–stromal interactions through dual-color fluorescence imaging (Yang et al., 2003). A very high-resolution approach to noninvasive imaging of tumor blood vessels involves implanting cancer cells expressing GFP in the nucleus and RFP in the cytoplasm into the footpad of the GFP nude mouse. This model, along with imaging using a scanning laser microscope especially designed for mice (the Olympus IV100), enables noninvasive imaging of tumor blood vessels, blood flow within the vessels, as well as individual color-coded cancer and stromal cells (Fig. 2.4) (Yang et al., 2007). Several different types of human cancer can be studied in dual-color orthotopic models, including breast (using the MD-435-RFP cell line), prostate (PC-3-RFP), colon (HCT-116-RFP), and fibrosarcoma (HT1080RFP) (Yang et al., 2004). Fresh tissue from GFP-expressing mice bearing B16F10-RFP mouse melanoma was used to image early events in tumor angiogenesis (Yang et al., 2003). The contrast between the green fluorescent host cells and the red fluorescent cancer cells allowed the two populations to

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A

50 µM B

50 µM C

50 µM

Figure 2.3 Visualization of angiogenesis in live tumor tissue 3 weeks after subcutaneous injection of B16F10-RFP melanoma cells in the transgenic GFP mouse. (A) Visualization of angiogenesis onset and development imaged in live tumor tissue. Host-derived GFP-expressing fibroblast cells (arrows) and endothelial cells (arrowheads) are shown forming new blood vessels in the RFP-expressing B16F10 melanoma. (B) Well-developed, host-derived, GFP-expressing blood vessels are visualized in the

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Figure 2.4 Whole-body, noninvasive, subcellular imaging of dual-color mouse mammary cancer cells and GFP stromal cells in the live GFP nude mouse. Dual-color MMT cells were injected in the footpad of GFP transgenic nude mice. Whole-body image of dual-color MMT tumor. Numerous dual-color, spindle-shaped MMT cells interacted with GFP-expressing host cells.Well-developed tumor blood vessels and real-time blood flow were visualized by whole-body imaging (arrows). (From Yang, M., Jiang, P., and Hoffman, R. M. (2007).Whole-body subcellular multicolor imaging of tumor-host interaction and drug response in real time. Cancer Res. 67, 5195^5200.) Bar ¼ 20 micrometers.

be clearly distinguished. Host-derived fibroblasts and endothelial cells as well as mature blood vessels could be seen easily against the red background of the tumor when imaged by dual-color fluorescence microscopy 3 weeks after implantation of tumor cells (Yang et al., 2003). This strategy offers the advantage of allowing the imaging of very small blood vessels, although removal of the tissue in question from the animal was performed. RFP-expressing mouse melanoma. (C) Tumor vasculature in viable tumor tissue and necrotic tumor tissue in the same tumor mass are visualized. GFP-expressing tumor vasculature can be readily identified in the area where the tumor tissue maintained good viability; however, only remnants of GFP-expressing vasculature can be visualized in the necrotic area. Bars ¼ 50 mm. (FromYang, M., Li, L., Jiang, P., Moossa, A. R., Penman, S., and Hoffman, R. M. (2003). Dual-color fluorescence imaging distinguishes tumor cells from induced host angiogenic vessels and stromal cells. Proc. Natl. Acad. Sci. USA 100,14259^14262.)


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100 µM D

100 µM

100 µM E

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100 µM G

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100 µM

Figure 2.5 Fluorescence imaging of tumor angiogenesis in transgenic ND-GFP nude mice. (A) RFP-expressing mouse B16F10 melanoma growing in a nestin-GFP transgenic nude mouse. Host-derived ND-GFPêxpressing blood vessels were visualized in the RFP-expressing mouse melanoma on day 10 after subcutaneous injection of B16F10RFP cells in the transgenic ND-GFP nude mouse. (B) Numerous host-derived ND-GFPêxpressing blood vessels were visualized in the RFP-expressing mouse mammary tumor on day 14 after orthotopic inoculation of MTT-RFP cells.

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4. Color-Coded Tumor-Host Models: Nestin-Driven–GFP Nude Mouse The stem-cell marker, nestin, is expressed in several different compartments within the mouse embryo as well as in the endothelial cells of developing blood vessels in the adult mouse. Immunocompetent mice expressing GFP under the control of the nestin promoter express GFP in all tissues in which the nestin promoter is active, including the nervous system, pancreas, hair follicles and developing blood vessels (Amoh et al., 2004, 2005b). GFP expression in nascent vasculature makes this animal model ideal for imaging developing tumor vasculature. The nestin-driven–GFP (ND-GFP) mouse was crossed with an athymic mouse strain on a C57/B6 background to generate the ND-GFP nude mouse (Amoh et al., 2005b). In this nude ND-GFP mouse, GFP is expressed in neural tissue, pancreas, stomach and esophagus, hair follicles, and developing blood vessels (Amoh et al., 2005b). This new mouse strain allows the imaging of human cancer cell interaction with mouse stromal tissues, especially with the developing tumor vasculature. Human cancer cell lines that have been engineered to express RFP have been orthotopically implanted into several different sites in this model, including lung, pancreas, colon, and skin (melanoma), and grew predictably and extensively (Amoh et al., 2005b). GFP was expressed within these growing tumors in the developing vasculature in all types of cancer evaluated. Mouse cell lines expressing RFP can likewise be imaged in this model, allowing visualization of the developing blood vessels in the murine tumor (Fig. 2.4). ND-GFP mice nude mice allow imaging of early, intermediate, and late-stage vascular development in transplanted tumors (Amoh et al., (C) RFP-expressing U87 human glioma growing in the ND-GFP transgenic nude mouse. ND-GFPêxpressing blood vessels were visualized in the RFP-expressing human glioma on day 14 after subcutaneous injection of U87-RFP cells. (D) Human HT1080 fibrosarcoma on day 14 after injection. Dual-color tumor cells expressing GFP in the nucleus and RFP in the cytoplasm are polarized towards ND-GFPêxpressing blood vessels (white arrows). (E) RFP-expressing Bx-PC-3 human pancreatic tumor vascularized with ND-GFP vessels on day 14 after orthotopic implantation. (F) RFPexpressing human HCT-116 colon tumor vascularized with ND-GFP vessels on day 14 after orthotopic implantation. (G) Extensive ND-GFPêxpressing blood vessels were visualized in the RFP-expressing human fibrosarcoma 8 days after injection of HT1080 cells. Only ND-GFP vessels are visualized. (H) Extensive inhibition of ND-GFP^ expressing blood vessel formation in the RFP-expressing, HT-1080 human fibrosarcoma by 5-mg/g doxorubicin (intraperitoneal) on days 0, 1, and 2. Bar ¼ 100 mm. (From Amoh,Y.,Yang, M., Li, L., Reynoso, J., Bouvet, M., Moossa, A. R., Katsuoka, K., and Hoffman, R. M. (2005b). Nestin-linked green fluorescent protein transgenic nude mouse for imaging human tumor angiogenesis. Cancer Res. 65, 5352^5357.)


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2005a). Within the first 5 days after subcutaneous implantation of RFPpositive cancer cells, GFP-expressing blood vessels could be imaged in the periphery of the tumor (day 1). Green tumor vessels were imaged first extending to the tumor margin (day 2) and later growing into the tumor (day 5). In the more intermediate stages of tumor growth, the blood vessels could be seen forming a network within the tumor (day 7) and later dilating as they began to support blood flow (day 14). At the late stages of tumor growth, the more mature blood vessels that were supporting blood flow could be distinguished by the presence of red blood cells within their lumen. The mature blood vessels had lost their GFP expression (day 28 to 35), although GFP expression was preserved in the smaller developing blood vessels at the tumor periphery. Immunohistochemical staining revealed CD31 and GFP expression colocalized in blood vessels within the growing tumor (Amoh et al., 2005a).

5. Imaging Cancer Cell Trafficking in Lymphatics: Experimental Metastasis Model Cancer cell and host interactions have been studied within lymph node tissue by various methods including intravital fluorescence microscopy. Recent work has used methods such as microlymphography to study peritumoral lymphatic interstitial fluid pressure and lymphatic flow (Leu et al., 2000; Jain and Fenton, 2002). More recently, mouse models that allow the study of cancer-cell movement through lymphatics themselves have become available. Cancer cell movement through lymphatic vessels can be followed in real time at the cellular and subcellular level using dual-color fluorescence imaging. In an experimental model of lymphatic metastasis, cancer cells engineered to express fluorescent proteins are injected into the inguinal lymph node of an anesthetized animal (Fig. 2.6). For simultaneous visualization of the lymphatic architecture and cancer cells trafficking within them, the use of dilute FITCdextran along with RFP-expressing cancer cells allows the specific imaging of those cells traveling through lymphatics ( Hayashi et al., 2007). Maximum image resolution is achieved using a reversible skin flap exposing the inguinal and axillary lymph nodes as well as the interconnecting lymphatics of the anterior abdominal wall ( Hayashi et al., 2007). Differential evaluation of cancer cell movement through lymphatics and survival in receiving lymphatic tissue in this experimental model can be achieved by simultaneous delivery of two or more cancer cell lines expressing different fluorescent proteins. The human osteosarcoma cell line 143BGFP and the murine melanoma cell line B16-RFP were compared in this manner, with no difference noted in cell trafficking through the lymphatics, although much greater survival within the receiving lymph node was found

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A

B

Cell trafficking

Axillary lymph node

Inject into inguinal lymph node

C

Surface of the axillary lymph node

Figure 2.6 Experimental model of lymph node metastasis: Cancer cells homing to the axillary lymph node after injection in the inguinal lymph node. (A) Dual-color HT-1080 human fibrosarcoma cells were injected into the inguinal lymph node. Nude mice were anesthetized and an arc-shaped incision was made in the abdominal skin from the axillary to the inguinal region.The subcutaneous connective tissue was separated to free the skin flap without injuring the lymphatic.The skin flap was spread and fixed on the flat stand. The axillary lymph node was exposed. (B) Cancer cells entering the axillary lymph node via the afferent lymph duct. Atotal10 ml of medium containing 5 104 HT-1080 dual-color cells was injected into the center of the inguinal lymph node. (C) Schematic of (B). (From Hayashi, K., Jiang, P.,Yamauchi, K.,Yamamoto, N.,Tsuchiya, H.,Tomita, K., Moossa, A. R., Bouvet, M., and Hoffman, R. M. (2007). Real-time imaging of tumor-cell shedding and trafficking in lymphatic channels. Cancer Res. 67,8223^8228.)

in the murine melanoma cell line (Hayashi et al., 2007). Again, image resolution was improved by elevating a reversible skin flap, which could be sutured closed and re-elevated several days later to facilitate repeated imaging in the same living animal. In addition to labeling lymphatics transiently with FITC-dextran, a more durable and high-resolution fluorescence signal within the lymphatic tissue, including lymph nodes and lymphatic vessels, can be achieved using fluorescein-conjugated anti-LYVE1 antibody. Injection of a few micrograms of labeled antibody into the inguinal lymph node is sufficient for labeling of the major anterior abdominal wall lymphatics, with signal lasting out to 3 days after a single administration. When RFP-expressing cancer cells are delivered to the inguinal lymph node as described above, these cells can be imaged traveling through the local lymphatic network and depositing into the subcapsular sinus of the receiving lymph node in real time (McElroy et al., 2008) (Fig. 2.7).

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A

B

C

D

E

F

G

Axillary lymph node

Tumor cells

Figure 2.7 Sequential images of cancer cells traveling through anterior abdominal wall lymphatics. Following administration of conjugated LYVE-1, the pancreatic cancer cell line XPA-1 RFP was injected into the inguinal lymph node. Red fluorescent XPA-1 cells, both individually (small arrows) and in clusters could be seen trafficking through the fluorescent LYVE-1 labeled lymphatics (A to F). RFP-labeled cancer cells could also be seen collecting in the axillary lymph node after labeling of the node and lymphatics with green fluorescent LYVE-1antibody (G). (From McElroy, M., Hayashi, K., GarmySusini, B., Kaushal, S., Varner, J. A., Moossa, A. R., Hoffman, R. M., and Bouvet, M. (2008). Fluorescent LYVE-1 antibody to image dynamically lymphatic trafficking of cancer cells in vivo. J. Surg. Res., Epub ahead of print.

6. Imaging Cancer Cell Trafficking in Lymphatics: Spontaneous Metastasis Model While experimental metastasis models can easily facilitate the study of differential tumor cell movement through lymphatic channels in real time, the early events that initiate tumor cell shedding into lymphatic channels are

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more accurately recapitulated in spontaneous models of metastasis. The mouse footpad is an ideal site for the observation of spontaneous tumor metastasis to the lymphatics. Exposure of the draining lymphatics of the leg as well as the first downstream lymph node, the popliteal, allows imaging of tumor cell trafficking and survival outside the primary tumor. The dualcolor human fibrosarcoma cell line HT-1080, which had been engineered to express GFP in the nucleus and RFP in the cytoplasm, was injected into the footpad of a nude mouse. Four weeks after cell injection, the animals were anesthetized and the draining lymphatics as well as the popliteal lymph node were exposed (Fig. 2.8). Cancer cells within the lymphatic were A

B Popliteal lymph node with spontaneous metastasis

Metastasis of HT-1080

Exposed popliteal lymph node Primary tumor in the foot pad 10 mm

C

D Vein Poplitical lymph node Dual-colored HT-1080 cells Footpad

Lymph duct 0.1 mm

Figure 2.8 Footpad spontaneous metastasis model. (A) Dual-color HT-1080 cells expressing RFP in the cytoplasm and H2B-GFP in the nucleus were injected into the footpad. Four weeks later, mice were placed in the prone position and the primary tumor and popliteal lymph node metastasis were observed by whole-body fluorescence imaging with the Olympus OV100. (B) Fluorescence image of exposed popliteal lymph node as seen noninvasively in (A). Dual-color HT-1080 cells metastasized to the lymph node as observed by fluorescence imaging. (C) Exposed popliteal lymph node. Area of lymphatic with trafficking cancer cells shown at high magnification in (D). (D) Fluorescence image of trafficking HT-1080 dual-color cells in the lymphatic denoted by dotted lines in (C). Bar ¼ 10 mm (A);1 mm (B); 2 mm (C); and 100 mm (D). (From Hayashi, K., Jiang, P., Yamauchi, K., Yamamoto, N., Tsuchiya, H., Tomita, K., Moossa, A. R., Bouvet, M., and Hoffman, R. M. (2007). Real-time imaging of tumorcell shedding and trafficking in lymphatic channels. Cancer Res. 67, 8223^8228.)


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imaged, and single cells as well as clumps of cells could easily be distinguished from cellular debris and fragments using dual-color fluorescence imaging (Hayashi et al., 2007).

7. Protocol: Production of FluorescentProtein–Expressing Cancer Cell Lines 7.1. GFP retrovirus production 1. GFP expression vector: Use the pLEIN or an equivalent retroviral vector expressing GFP and a neomycin-resistance gene on the same bicistronic message. 2. PT67 (an NIH3T3-derived packaging cell line) expressing the 10-A1 viral envelope is used for retroviral production. The PT67 cells are cultured in Dubecco’s Modified Essential Medium (DMEM) supplemented with 10% (vol/vol) heat-inactivated, fetal calf serum (FCS). 3. PT67 cells are grown to 70% confluence and harvested. Cells are plated at 60 to 80% confluence on a 60-mm culture dish 12 h prior to transfection. The Lipofectamine Plus transfection kit is used with 10 mg of pLEIN-GFP. Add 7 ml of precomplexed pLEIN-GFP DNA in 87 ml of serum-free medium and mix. Next, add 6 ml of PLUS reagent, mix, and incubate at room temperature (RT; 22 to 26 C) for 15 min. 4. Dilute 4 ml of Lipofectamine in 96 ml of serum-free medium in a separate tube. Incubate this mixture at RT for 15 min. 5. The pre-complexed DNA and diluted Lipofectamine are combined, mixed, and incubated at RT for 15 min. 6. During complex formation, cell medium is replaced with 800 ml of serum-free DMEM. Add the DNA-Lipofectamine reagent complex to the cells and gently mix the complexes with the cells. Incubate this mixture at 37 C, 5% CO2 for 4 h. 7. After 4 h, increase the medium volume to 5 ml, and continue to incubate at 37 C, 5% CO2 for 24 h. 8. After the 24 h incubation, the packaging cells are cloned by limiting dilution in 96-well plates with cells plated to a density of less than one cell per well. 9. Examine the clones by fluorescence microscopy at 48 h after transduction. 10. For selection, culture the cells in stepwise increasing doses of G418 starting at 300 mg/ml and increasing up to 500 to 2000 mg/ml to select for clones producing high levels of GFP retroviral vector (PT67-GFP). Culture the cells for 1 to 2 days in each concentration of G418. Highviral-production clones of GFP PT67 with a titer of more than 106

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plaque-forming units (pfu) are used for GFP vector production (Hoffman and Yang, 2006b, 2006c). Note: A stepwise increase in G418 concentration is critical for selecting packaging cells with high viral titers.

7.2. RFP retrovirus production 1. The pLNCX2-Ds-Red2 plasmid is produced by inserting the HindIII/ NotI fragment from pDsRed2, which contains the full-length red fluorescent protein cDNA, into the HindIII/NotI site in the pLNCX2 vector, which contains a neomycin-resistance gene. 2. Incubate PT67 cells at 70% confluence for 2 h at 37 C, 5% CO2. The Lipofectamine Plus transfection kit is used as described above to transfect the pLNCX2-DsRed2 vector into the PT67 packaging cells. 3. The PT67-DsRed2 cells are cultured in the presence of 200 to 1000 mg/ ml G418 in a stepwise manner as described above in order to select a clone expressing high titers of viral product (Hoffman and Yang, 2006a; Hoffman and Yang, 2006c). Note: A stepwise increase in G418 concentration is critical for selecting packaging cells with high viral titers.

7.3. Production of the histone H2B-GFP vector 1. The histone H2B-GFP fusion vector is inserted into the pLHCX retrovirus containing the hygromycin-resistance gene at its HindIII/ClaI site. 2. In order to establish a packaging cell clone producing high levels of the histone H2B-GFP retroviral vector, transfect the pLHCX histone H2BGFP plasmid in PT67 packaging cells as described for GFP and DsRed2 retrovirus production. 3. Culture cells in the presence of 200 to 400 mg/ml hygromycin to establish stable PT67 H2B-GFP packaging cells. The amount of hygromycin is increased stepwise as described for G418. Note: As with G418 selection, the stepwise increase in hygromycin concentration is critical for selecting packaging cells with high viral titers.

7.4. GFP or RFP transduction of tumor cell lines 1. Twenty percent confluent tumor cells are used for transduction. Twelve to 18 h prior to transduction, cells are plated at a density of approximately 1 to 2 105 per 60-mm plate. 2. Collect conditioned medium from the PT67 packaging cells (PT67/ pLEIN GFP or PT67/pLNCX2-DsRed2), and filter through a 0.45-mm


3. 4. 5. 6. 7.

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polysulfonic filter. Add this virus-containing filtered medium to the target cells. Add polybrene to a final concentration of 8 mg/ml. Incubate the cells for 24 h at 37 C, 5% CO2. Replace medium with DMEM containing 10% FCS (vol/vol) after 24-h incubation. Fluorescence microscopy is used to check for GFP- or RFP-expressing cells. Harvest tumor cells and subculture at a ratio of 1:15 in selective medium containing 50 mg/ml G418. For selection of brightly fluorescent cells, increase the G418 level in a stepwise manner to a final concentration of 800 mg/ml. Culture the cells for 1 to 2 days at each G418 concentration. Isolate clones expressing GFP or RFP with cloning cylinders and amplify them in DMEM with 10% FCS in the absence of selection medium (Hoffman and Yang, 2006b; Hoffman and Yang, 2006c).

Note: Amplification of clones in the absence of selection medium will ensure that the cells will stably GFP or RFP without the presence of G418, which will not be present during in vivo experimental use (Hoffman and Yang, 2006a,b,c).

7.5. Double RFP and histone H2B-GFP gene transduction of cancer cells 1. In order to establish dual-color tumor cells, start with tumor cells expressing RFP in the cytoplasm at 70% confluence. 2. RFP-expressing cells are incubated as described above with filtered conditioned medium from the PT67 H2B-GFP cells for 48 h at 37 C, 5% CO2. 3. In order to select for double-transformants, transfer cells with the PLHCX-histone H2B-GFP retroviral vector and then select cells in hygromycin-containing media as described above (Hoffman and Yang, 2006b,c).

7.6. Protocol: Conjugation of anti LYVE-1 antibody 1. Reconstitute antibody at 1 mg/ml in phosphate buffered saline (PBS). 2. Add 100 ml of the 1 mg/ml solution to the AlexaFluor 488 reactive dye mixture. Adjust the pH of the mixture to 8 using sodium bicarbonate as needed. 3. Incubate the mixture 1 h at RT followed by overnight at 4 C. 4. The conjugated antibody is then separated from the remaining unconjugated dye on a purification column by centrifugation.

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5. Antibody and dye concentrations in the final sample are determined spectrophotometrically based on absorbance of the final purified conjugate at 280 nm and 494 nm, respectively (McElroy et al., 2008). Note: For optimal in vivo imaging, the appropriate final ratio of moles dye:moles antibody is approximately 5 to 10:1. Improved conjugation ratios can be achieved by using double the dye mixture during the initial conjugation.

7.7. Protocol: Establishment of imageable tumor models—cell injection 1. Harvest GFP-expressing tumor cells by trypsinization and wash three times with serum-free medium. Keep harvested cells on ice. 2. Re-suspend cells in serum-free medium. The concentration of resuspended cells will vary depending on the planned site of injection. Cells should be injected within 30 min of harvest to preserve viability. 3. For subcutaneous tumors, cells are re-suspended at 1 106 in 50 to 100 ml of medium and injected subcutaneously where desired. For cell implantation into the pancreas, mammary fat pad, or footpad, cells are resuspended at 1 106 in 10 to 20 ml of medium and injected (Hoffman and Yang, 2006a,c). Note: Rapid use of cells after trypsinization is critical as cell viability decreases significantly with time after harvest.

7.8. Surgical orthotopic implantation 1. Animals are anesthetized with a ketamine-xylazine mixture (50% ketamine, 38% xylazine, and 12% agepromazine maleate injected intramuscularly at a dose of 2 ml/g). 2. Subcutaneously-growing fluorescent tumor is harvested and sectioned into 1-mm3 fragments under sterile conditions. 3. After sterile surgical exposure of the target organ, 1-mm3 fragments of fluorescent tumor are sutured into the organ using 8-0 nylon surgical sutures. 4. The surgical site is closed using 6-0 absorbable surgical sutures. Animals are maintained in a HEPA-filtered barrier facility. Note: Surgical implantation of tumor tissue reduces the risk of tumor cell spillage and contamination of adjacent tissues.

7.9. Experimental lymphatic metastasis model 1. Animals are anesthetized as described and placed supine. The anterior abdominal wall is sterilized and a skin flap is elevated exposing the anterior abdominal wall lymphatics with the inguinal and axillary lymph nodes.


2.

3. 4. 5. 6.

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Careful attention must be paid to preserving the lymphatics during the dissection. The internal organs remain covered by the intact peritoneum. The axillary lymph node can be further exposed by elevating the pectoral muscle and freeing the lymph node from either the skin or chest wall side. Again, careful attention must be paid to minimizing trauma to the afferent lymphatics leading to the node. The skin flap is stabilized away from the body of the mouse, and fluorescent protein-expressing tumor cells are injected into the inguinal lymph node at 5 104 in 10 ml of PBS with or without 2.5 ml of FITC-dextran. Alternatively, 10 ml of conjugated LYVE-1 antibody at 0.25 mg/ml can be injected into the inguinal lymph node for repeated lymphatic imaging, followed by cell injection in PBS. After cell injection, imaging is acquired using either the OV-100 or IV100 systems as described below. The previously elevated skin flap can be re-secured under sterile conditions using absorbable 6-0 surgical sutures. The flap can be re-elevated to allow repeated imaging of cell movement and tumor growth (Hayashi et al., 2007; McElroy et al., 2008).

7.10. Spontaneous lymphatic metastasis model 1. Anesthetized animals are given a single injection of 1 to 2 106 fluorescent protein-expressing tumor cells in 10-ml volume into the rear footpad. 2. Four weeks later, the animals are again anesthetized and placed prone. The tumor-bearing rear foot is stabilized. 3. Careful surgical dissection through a 2-cm incision in the ipsilateral popliteal region is used to expose the draining lymphatics of the hind leg and the popliteal lymph nodes. 4. The lymphatics and lymph nodes are imaged using either the OV-100 or IV-100 imaging systems. Lymphatic resolution can be improved by the injection of 5 ml of FITC-dextran into the peritumoral tissue of the footpad (Hayashi et al., 2007).

8. Protocol: Imaging of Cancer Cell Interactions with Blood Vessels and Lymphatics 8.1. Whole-body imaging of tumor vasculature 1. Animals are anesthetized as described, and, as needed, overlying hair is removed using either clippers or depilatory creams. The animals are

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placed within the chamber of the OV-100 Small Animal Imaging System (Olympus Corp., Tokyo, Japan) oriented such that the tumor is visible. Note: Adequate removal of all hair is critical to imaging. 2. Images through the skin are collected at varying exposure times and magnifications to visualize tumor vasculature. Functional blood vessels carrying blood are visible as nonfluorescent regions against the background of the green fluorescent tumor in nonfluorescent mice and by their green fluorescence against the RFP-expressing tumor in the case of the GFP or ND-GFP mouse (Hoffman and Yang, 2006a,c). Note: While whole-body imaging is rapid and noninvasive, and can be repeated multiple times in the same animal, disadvantages include signal scatter and attenuation through the overlying tissues making high-resolution imaging difficult.

8.2. Skin flap elevation for improved imaging of superficial tumor vasculature for lymphatic migration 1. After anesthetization of the animals and removal of hair, a local skin flap can be elevated by creating a three-sided trapdoor incision overlying the superficial tumor. A larger skin flap can be elevated for the exposure of lymphatic channels and nodes. 2. Imaging is completed as described in Section 9. 3. The skin flap can be reapproximated and sutured closed under sterile conditions using 6-0 absorbable surgical sutures. The same skin flap can be re-elevated days to weeks later to allow repeated high-resolution imaging in the same living animal (Hayashi et al., 2007; Yang et al., 2002). Note: While skin flap elevation offers improved resolution over wholebody imaging, scar tissue formation can limit the total number of times that skin-flap opening can be repeated.

8.3. Intravital imaging for deep tumors 1. Following anesthetization of the tumor-bearing animal, deep internal organs can be imaged with high resolution by intravital imaging. 2. Under sterile conditions, the organ of interest containing the tumor is surgically exposed and stabilized, and high-resolution images are acquired. 3. Following imaging, the tumor and organ are returned to their native position, and the surgical site is closed and sutured. Animals can be reimaged a limited number of times by this method (Yang et al., 2001).


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Note: While intravital offers improved resolution for deep tumors, the length of imaging time must be minimized and in general this imaging technique can be repeated fewer times than whole-body imaging or skin flap elevation in the living animal.

8.4. Tumor tissue sampling for highest-resolution imaging of tumor vasculature 1. Tumor tissue can be harvested from 3 days to 4 weeks following implantation into either GFP or ND-GFP mice. Tissue can be collected ex vivo, or as a biopsy from living anesthetized animals. Bleeding from the surgical site is controlled with gentle pressure. 2. Fresh tumor tissue is sectioned into 1-mm3 pieces and compressed between glass slides for imaging. 3. For maximal tumor vasculature measurement, the tissues can be digested briefly with trypsin-EDTA for 5 min at 37 C prior to compression and imaging. 4. Tumor vasculature can be quantified in by obtaining images of all fields and calculation of the total vasculature length in all fields. Blood vessel density is calculated by dividing the total length of GFP-expressing vessels by the tumor volume imaged (in millimeters per square millimeter) (Hoffman and Yang, 2006a). Note: This technique offers the highest level of imaging resolution, but in general can not be repeated in the same animal.

9. Protocol: Imaging Methods for Mice Expressing Fluorescent Proteins 9.1. Imaging with a handheld flashlight 1. A blue LED flashlight with an excitation filter (mid-point wavelength peak of 470 nm) and a DP470/40 emission filter can be used for wholebody imaging of animals with either GFP- or RFP-expressing tumors. 2. Images can be acquired with a camera system such as the Nikon CoolPIX or simple CCD camera with an appropriate emission filter and stored (Hoffman and Yang, 2006a,c).

9.2. Imaging with fluorescence microscopy 1. An Olympus BH 2-RFCA fluorescence microscope equipped with a mercury 100-watt lamp power supply or its equivalent can be used.

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2. For visualization of GFP and RFP, the excitation light can be produced through a D425/60 band-pass filter and a 470 DCXR dichroic mirror. 3. Emitted fluorescent light is collected through a GG475 long-pass filter. 4. High-resolution images are captured with a Hamamatsu C5810 threechip–cooled color CCD camera or its equivalent. 5. Images can be processed as needed after collection (Hoffman and Yang, 2006a,c).

9.3. Imaging with the Olympus OV-100 small-animal imaging system 1. The Olympus OV-100 with a 470-nm excitation light originating from an MT-20 light source can be used for imaging tumor blood vessels. Emitted fluorescence is collected through appropriate filters on a filter wheel with a DP70 CCD camera. 2. Adjustment of emission filters can allow specific imaging of various different fluorophores including GFP and RFP. 3. Variable magnification in this system is available with a series of four objective lenses. Images ranging from whole-body to single-cell resolution can be acquired through the variable microscopy settings. 4. Images are stored and can be processed further after acquisition for tumor volume or fluorescence measurements as well as for quantification of tumor vasculature (Hoffman and Yang, 2006a,c).

9.4. Imaging with the Olympus IV-100 scanning laser microscope system 1. The tissue to be imaged using this microscopy system may be imaged either ex vivo or in a deeply anesthetized animal while secured. It is critical for optimal image resolution that the tissue being imaged does not move with the respiratory and cardiac variation in the animal (Yang et al., 2007). 2. Ex vivo tissue can be simply placed on a dark surface under the IV-100 objective with frequent application of PBS to keep the tissue moist during imaging. 3. Imaging of lymphatics, blood vessels, and tumor tissue in a skin flap requires stabilization of the skin flap itself away from the body of the animal. 4. Likewise, imaging of tumor lymphatics or blood vessels in the leg can be achieved by stabilization of the extremity such that the animal’s respiratory variation does not cause movement artifact in the tissue being imaged. 5. Intravital imaging of deeper organs in living animals requires stabilization of the organ and tumor tissue in question. This can be achieved in some


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organs, such as the pancreatic tail, that can be moved and stabilized without sacrifice of the animal provided that the mouse remains deeply anesthetized throughout the duration of the imaging procedure. For tissues that cannot be stabilized in this manner, such as the heart and lung, ex vivo tissue analysis affords the best imaging. 6. Variable magnification down to the subcellular level can be imaged using the full range of objectives. Differential excitation of fluorophores can be achieved in this system by the use of three different lasers for excitation at 488 nm, 561 nm, and 633 nm. Note: While this imaging technique provides ultra-high-resolution imaging of cancer cells, tissue stabilization is absolutely critical to the acquisition of these high-resolution images.

9.5. Imaging using spectral separation 1. The standard fluorescence imaging system previously described is replaced with a cooled monochrome camera and liquid-crystal tunable filter (CRI, Inc., Woborn, MA) positioned in front of a conventional macro-lens. 2. A series of images is typically acquired every 10 nm from 500 to 650 nm and assembled into a spectral ‘‘stack.’’ 3. Using the predefined GFP and RFP emission spectra, the colleted spectral ‘‘stack’’ can be resolved into various images corresponding to specific wavelengths of interest that represent autofluorescence, GFP, and RFP signals. 4. This method allows for maximal signal-to-noise ratio acquisition by virtue of its ability to separate out the competing autofluorescence or other fluorescence signals. 5. It is critical for this image acquisition that there be no movement in the tissue imaged when overlay images of multiple fluorescence signals are to be created (Mansfield et al., 2005). Note: In general, spectral separation imaging systems can provide greater sensitivity for specific fluorophore emission, although not all systems are equipped for high-resolution imaging.

10. Summary and Conclusions A number of different methods are now available for the evaluation of cancer cell interaction with blood vessels and lymphatics. Newer techniques involve the use of fluorescence imaging technology and allow repeated imaging in the living animal. Combining these imaging strategies with the

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evaluation of orthotopic tumor models offers the opportunity to longitudinally follow the natural progression of the disease. In general, the less invasive imaging procedures such as simple whole-body fluorescence imaging are limited by fluorescence signal attenuation by overlying tissues, whereas the more invasive approaches such as intravital and skin flap elevation provide much higher resolution but can be repeated only a limited number of times. These models offer a range of different approaches to imaging tumor cell interactions with host lymphatics and blood vessels.

REFERENCES Al-Mehdi, A. B., Tozawa, K., Fisher, A. B., Shientag, L., Lee, A., and Muschel, R. J. (2000). Intravascular origin of metastasis from the proliferation of endothelium-attached tumor cells: A new model for metastasis. Nat. Med. 6, 100–102. Alessandri, G., Raju, K., and Gullino, P. M. (1983). Mobilization of capillary endothelium in vitro induced by effectors of angiogenesis in vivo. Cancer Res. 43, 1790–1797. Amoh, Y., Li, L., Yang, M., Jiang, P., Moossa, A. R., Katsuoka, K., and Hoffman, R. M. (2005a). Hair follicle-derived blood vessels vascularize tumors in skin and are inhibited by Doxorubicin. Cancer Res. 65, 2337–2343. Amoh, Y., Li, L., Yang, M., Moossa, A. R., Katsuoka, K., Penman, S., and Hoffman, R. M. (2004). Nascent blood vessels in the skin arise from nestin-expressing hair-follicle cells. Proc. Natl. Acad. Sci. USA 101, 13291–13295. Amoh, Y., Yang, M., Li, L., Reynoso, J., Bouvet, M., Moossa, A. R., Katsuoka, K., and Hoffman, R. M. (2005b). Nestin-linked green fluorescent protein transgenic nude mouse for imaging human tumor angiogenesis. Cancer Res. 65, 5352–5357. Auerbach, R., Kubai, L., Knighton, D., and Folkman, J. (1974). A simple procedure for the long-term cultivation of chicken embryos. Dev. Biol. 41, 391–394. Cowen, S. E., Bibby, M. C., and Double, J. A. (1995). Characterisation of the vasculature within a murine adenocarcinoma growing in different sites to evaluate the potential of vascular therapies. Acta Oncol. 34, 357–360. Crum, R., Szabo, S., and Folkman, J. (1985). A new class of steroids inhibits angiogenesis in the presence of heparin or a heparin fragment. Science 230, 1375–1378. Deutsch, T. A., and Hughes, W. F. (1979). Suppressive effects of indomethacin on thermally induced neovascularization of rabbit corneas. Am. J. Ophthalmol. 87, 536–540. Dewhirst, M., Gross, J. F., Sim, D., Arnold, P., and Boyer, D. (1984). The effect of rate of heating or cooling prior to heating on tumor and normal tissue microcirculatory blood flow. Biorheology 21, 539–558. Duda, D. G., Fukumura, D., Munn, L. L., Booth, M. F., Brown, E. B., Huang, P., Seed, B., and Jain, R. K. (2004). Differential transplantability of tumor-associated stromal cells. Cancer Res. 64, 5920–5924. Epstein, R. J., Hendricks, R. L., and Stulting, R. D. (1990). Interleukin-2 induces corneal neovascularization in A/J mice. Cornea 9, 318–323. Fournier, G. A., Lutty, G. A., Watt, S., Fenselau, A., and Patz, A. (1981). A corneal micropocket assay for angiogenesis in the rat eye. Invest. Ophthalmol. Vis. Sci. 21, 351–354. Fukumura, D., Xavier, R., Sugiura, T., Chen, Y., Park, E. C., Lu, N., Selig, M., Nielsen, G., Taksir, T., Jain, R. K., and Seed, B. (1998). Tumor induction of VEGF promoter activity in stromal cells. Cell 94, 715–725.


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Gimbrone, M. A., Jr., Cotran, R. S., Leapman, S. B., and Folkman, J. (1974). Tumor growth and neovascularization: An experimental model using the rabbit cornea. J. Natl. Cancer Inst. 52, 413–427. Hayashi, K., Jiang, P., Yamauchi, K., Yamamoto, N., Tsuchiya, H., Tomita, K., Moossa, A. R., Bouvet, M., and Hoffman, R. M. (2007). Real-time imaging of tumor-cell shedding and trafficking in lymphatic channels. Cancer Res. 67, 8223–8228. Hoffman, R. (2002). Green fluorescent protein imaging of tumour growth, metastasis, and angiogenesis in mouse models. Lancet Oncol. 3, 546–556. Hoffman, R. M. (1999). Orthotopic metastatic mouse models for anticancer drug discovery and evaluation: A bridge to the clinic. Invest. New Drugs 17, 343–359. Hoffman, R. M. (2005). The multiple uses of fluorescent proteins to visualize cancer in vivo. Nat. Rev. Cancer 5, 796–806. Hoffman, R. M., and Yang, M. (2006a). Color-coded fluorescence imaging of tumor-host interactions. Nat. Protoc. 1, 928–935. Hoffman, R. M., and Yang, M. (2006b). Subcellular imaging in the live mouse. Nat. Protoc. 1, 775–782. Hoffman, R. M., and Yang, M. (2006c). Whole-body imaging with fluorescent proteins. Nat. Protoc. 1, 1429–1438. Jain, R. K., and Fenton, B. T. (2002). Intratumoral lymphatic vessels: A case of mistaken identity or malfunction? J. Natl. Cancer Inst. 94, 417–421. Korey, M., Peyman, G. A., and Berkowitz, R. (1977). The effect of hypertonic ointments on corneal alkali burns. Ann. Ophthalmol. 9, 1383–1387. Leu, A. J., Berk, D. A., Lymboussaki, A., Alitalo, K., and Jain, R. K. (2000). Absence of functional lymphatics within a murine sarcoma: A molecular and functional evaluation. Cancer Res. 60, 4324–4327. Li, C. Y., Shan, S., Huang, Q., Braun, R. D., Lanzen, J., Hu, K., Lin, P., and Dewhirst, M. W. (2000). Initial stages of tumor cell-induced angiogenesis: Evaluation via skin window chambers in rodent models. J. Natl. Cancer Inst. 92, 143–147. Mahoney, J. M., and Waterbury, L. D. (1985). Drug effects on the neovascularization response to silver nitrate cauterization of the rat cornea. Curr. Eye Res. 4, 531–535. Mansfield, J. R., Gossage, K. W., Hoyt, C. C., and Levenson, R. M. (2005). Autofluorescence removal, multiplexing, and automated analysis methods for in vivo fluorescence imaging. J. Biomed. Opt. 10, 41207. McElroy, M., Hayashi, K., Garmy-Susini, B., Kaushal, S., Varner, J. A., Moossa, A. R., Hoffman, R. M., and Bouvet, M. (2008). Fluorescent LYVE-1 antibody to image dynamically lymphatic trafficking of cancer cells in vivo. J. Surg. Res., Epub ahead of print. Miller, J. W., Stinson, W. G., and Folkman, J. (1993). Regression of experimental iris neovascularization with systemic alpha-interferon. Ophthalmology 100, 9–14. Muthukkaruppan, V., and Auerbach, R. (1979). Angiogenesis in the mouse cornea. Science 205, 1416–1418. Okabe, M., Ikawa, M., Kominami, K., Nakanishi, T., and Nishimune, Y. (1997). ‘Green mice’ as a source of ubiquitous green cells. FEBS Lett. 407, 313–319. Papenfuss, H. D., Gross, J. F., Intaglietta, M., and Treese, F. A. (1979). A transparent access chamber for the rat dorsal skin fold. Microvasc. Res. 18, 311–318. Passaniti, A., Taylor, R. M., Pili, R., Guo, Y., Long, P. V., Haney, J. A., Pauly, R. R., Grant, D. S., and Martin, G. R. (1992). A simple, quantitative method for assessing angiogenesis and antiangiogenic agents using reconstituted basement membrane, heparin, and fibroblast growth factor. Lab. Invest. 67, 519–528. Yang, M., Baranov, E., Li, X. M., Wang, J. W., Jiang, P., Li, L., Moossa, A. R., Penman, S., and Hoffman, R. M. (2001). Whole-body and intravital optical imaging of angiogenesis in orthotopically implanted tumors. Proc. Natl. Acad. Sci. USA 98, 2616–2621.

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Yang, M., Baranov, E., Wang, J. W., Jiang, P., Wang, X., Sun, F. X., Bouvet, M., Moossa, A. R., Penman, S., and Hoffman, R. M. (2002). Direct external imaging of nascent cancer, tumor progression, angiogenesis, and metastasis on internal organs in the fluorescent orthotopic model. Proc. Natl. Acad. Sci. USA 99, 3824–3829. Yang, M., Jiang, P., and Hoffman, R. M. (2007). Whole-body subcellular multicolor imaging of tumor–host interaction and drug response in real time. Cancer Res. 67, 5195–5200. Yang, M., Li, L., Jiang, P., Moossa, A. R., Penman, S., and Hoffman, R. M. (2003). Dualcolor fluorescence imaging distinguishes tumor cells from induced host angiogenic vessels and stromal cells. Proc. Natl. Acad. Sci. USA 100, 14259–14262. Yang, M., Reynoso, J., Jiang, P., Li, L., Moossa, A. R., and Hoffman, R. M. (2004). Transgenic nude mouse with ubiquitous green fluorescent protein expression as a host for human tumors. Cancer Res. 64, 8651–8656.

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Bone Marrow–Derived Vascular Progenitors and Proangiogenic Monocytes in Tumors Kan Lu,* Chrystelle Lamagna,* and Gabriele Bergers*,† Contents 1. Introduction 2. Methods for the Visualization of BMDCs in Tumors 2.1. Transplantation of GFP-expressing bone marrow to visualize BMDC in tumors 2.2. Visualization of GFPþ BMDC subpopulations in tumors 2.3. Quantification of GFPþ BMDC subpopulations in tumors 3. Analysis of Pericyte Progenitors in Tumors 3.1. Detection of pericyte progenitors and pericytes in tumors 3.2. Isolation of PDGFRbþ pericytes from Rip1Tag2 tumor cell suspension 3.3. In vitro differentiation of pericyte progenitors References

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Abstract In tumors, new blood vessels develop not only from pre-existing vessels (angiogenesis), but can also be comprised of circulating vascular progenitor cells originating from the bone marrow (vasculogenesis). Besides endothelial progenitor cells (EPC) and pericyte progenitor cells (PPCs) that are incorporated into the growing vasculature, other subpopulations of bone marrow–derived cells (BMDC) contribute indirectly to tumor neovascularization by providing growth factors, cytokines, and other key proangiogenic molecules. Here, we describe specific methods that allow for the identification and functional characterization of these distinct BMDC populations in tumors as exemplified in mouse models of pancreatic neuroendocrine tumors and glioblastomas.

* {

Department of Neurological Surgery, Brain Tumor Research Center, University of California-San Francisco, San Francisco, California Comprehensive Cancer Center, University of California-San Francisco, San Francisco, California


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1. Introduction A growing body of evidence supports the notion that new blood vessels do not exclusively originate from the existing vasculature by activation of vascular cells (endothelial cells and pericytes) within injured tissues or tumors, but are also formed with the help of vascular progenitors and proangiogenic myelocytic cells that are recruited from the bone marrow to sites of active vascular remodeling in the adult. Vascular progenitor cells consist of endothelial cell progenitor cells (EPCs) and pericyte progenitor cells (PPCs) that structurally contribute to the formation of new blood vessels. While EPCs incorporate into the vessel wall and mature into endothelial cells, PPCs envelop blood vessels and differentiate into pericytes supporting stabilization and maturation of the newly formed vasculature (Aghi and Chiocca, 2005; Allt and Lawrenson, 2001; Asahara et al., 1997; De Palma et al., 2005; Dome et al., 2008; Lyden et al., 2001; Rafii et al., 2003; Rajantie et al., 2004; Song et al., 2005). The ontogeny of vascular progenitors in the adult has become more complex with the revelation that vascular progenitors can originate from mesenchymal as well as from hematopoietic cells. The predominant portion of EPCs that are found in growing tumor vessels are derived from the bone marrow as CD45– VEGFR2þ CD133þ c-kitþ cells (Asahara et al., 1997; Rafii et al., 2002), but hematopoietic sources such as myeloid progenitor cells have also been described as having the propensity to differentiate into endothelial-like cells (Bailey et al., 2006; Rohde et al., 2006; Yang et al., 2004). Interestingly, in tumors bone marrow–derived (BMD) pericyte progenitors thus far appear to be of hematopoietic origin. BMD pericytes were identified as CD11bþ NG2þ cells in a subcutaneous Bl6-F1 melanoma model (Rajantie et al., 2004), and were found to originate from CD45þ Sca-1þ PDGFRbþ hematopoietic cells in a transgenic mouse model of pancreatic islet tumorigenesis (Song et al., 2005). In the latter tumor model, bone-marrow transplant experiments combined with FACS and immunohistochemical analyses revealed that BMD-Sca-1þ PDGFRbþ cells were able to develop into mature pericytes expressing the markers NG2, a-SMA, and desmin in vivo. Interestingly, only 15 to 20% of the pericytes in these tumors expressed the mature pericyte markers, whereas mature pericytes were devoid of PDGFRb. Complementary co-culture experiments of BMD or tumor-derived Sca-1þ cells with endothelial cells in a three-dimensional (3D) Matrigel matrix confirmed the propensity of these cells to differentiate into pericytes in vitro (Song et al., 2005). In addition, mesenchymal Sca-1þ Tie2þ CD13þ pericyte progenitors were identified in some tumors; however, these cells did not originate from the bone marrow but rather appeared to be recruited from the tumor stroma or neighboring tissue (De Palma et al., 2005). These cells are more reflective of

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pericytes that develop during vasculogenesis in the embryo, as those, albeit not exclusively like endothelial cells, originate from mesenchymal stem cells (Betsholtz et al., 2005; Carmeliet, 2003; Lamagna and Bergers, 2006). The extremely diverse incorporation rates of EPCs in blood vessels of different tumors, and even of the same tumor type at distinct stages of progression, have raised intense debate about the functional significance of EPCs in tumor neovascularization. Certainly in most instances, the incorporation rate of EPCs, and also of PPCs, is rather low but one cannot exclude the possibility that these cells can act as transient but significant catalysts of tumor neovascularization and progression. Congruent with this hypothesis, vascular-disrupting agents that ablate tumor blood vessels within a few hours and thereby cause severe hypoxia and necrosis, have been found to trigger transient homing of EPCs to the tumor margin, an effect sufficient to facilitate neovascularization and tumor regrowth (Shaked et al., 2006). Notably, these tumors did not contain substantial numbers of BMD cells (BMDCs) prior to treatment, but recruited such cells when confronted with a drug that scrutinized the tumor vasculature and caused hypoxia. The largest and most heterogeneous group of BMDCs in tumors consists of CD45þ myelocytic cells that contribute indirectly to neovascularization by expressing a variety of proangiogenic cytokines, growth factors, and proteases (Grunewald et al., 2006). Such cells include tumor-associated macrophages (TAMs) (Condeelis and Pollard, 2006; Pollard, 2004), immature monocytic cells including Tie2þ monocytes (TEMs) (De Palma et al., 2005), CXCR4þ VEGFR1þ hemangiocytes (Hattori et al., 2002; Jin et al., 2006), and Gr1þ CD11bþ myeloid cells (Bunt et al., 2006; Yang et al., 2004). Inhibition of any of these monocytic subpopulations in tumor models reduced or restrained tumor neovascularization (De Palma et al., 2005; Du et al., 2008; Grunewald et al., 2006; Pollard, 2004; Yang et al., 2004). Recent data revealed that one of the proangiogenic factors commonly expressed among these monocytic subpopulations is the matrix metalloproteinase MMP-9 (Ahn and Brown, 2008; Bergers et al., 2000; Coussens et al., 2000; Du et al., 2008; Seandel et al., 2008; Yang et al., 2004). MMP-9 can promote neovascularization by different means. It degrades extracellular matrix components to allow endothelial cell invasion and cleaves c-kit ligand to facilitate mobilization of vascular progenitor cells from the bone marrow into the bloodstream (Egeblad and Werb, 2002; Heissig et al., 2002; Page-McCaw et al., 2007). In addition, MMP-9 has also been shown to be essential in initiating neovascularization in mouse models of pancreatic islet tumorigenesis and glioblastomas whereby MMP-9 expressed from tumor-recruited BMDCs can liberate sequestered VEGF from the extracellular matrix, thus increasing VEGFR-2 activation and angiogenesis (Bergers et al., 2000; Du et al., 2008). Importantly, CD45þ cells that expressed MMP-9 were shown to be sufficient to initiate the angiogenic switch in pancreatic and brain tumors.

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How do BMDCs specifically home to sites of vascular remodeling in tumors? Thus far only a few factors are known to enable the mobilization of BMDC from bone marrow into the bloodstream, as well as their subsequent recruitment and retention into the tumor. The most prominent factors identified to date include tumor-produced VEGF and PlGF, which act as mobilization factors of EPCs, and PDGF-B, which mobilizes PPCs (Gerhardt and Betsholtz, 2003; Gerhardt et al., 2003; Li et al., 2006; Luttun et al., 2002; Rabbany et al., 2003); and stromal-derived factor 1a(SDF1a), which acts to retain CXCR4þ BMDCs within tumors (Du et al., 2008; Grunewald et al., 2006; Hattori et al., 2003). Evidence that microenvironmental influences such as low oxygen tension can trigger BMDC recruitment stems from observations in experimentally induced ischemic tissues in which endothelial progenitors and other CXCR4þ BMDCs were recruited, in part through increases in hypoxia-inducible factor 1a (HIF-1a) and its downstream targets SDF1a and VEGF (Ceradini et al., 2004; De Falco et al., 2004; Petit et al., 2007). In another line of investigation, HIF-1a–induced recruitment of proangiogenic BMD CD45þ myeloid cells, which included proangiogenic Tie2þ, VEGFR1þ, CD11bþ, and F4/80þ subpopulations, as well as endothelial and pericyte progenitor cells, was found to promote neovascularization in glioblastoma, a tumor type characterized by extensive hypoxia and necrosis (Aghi et al., 2006; Du et al., 2008). These data suggest that all of these cell types can participate as functionally significant constituents in neovascularization, although MMP-9 expressing monocytic BMDCs appeared to be sufficient to initiate angiogenesis in these tumors (Du et al., 2008). The significant variation in recruitment and utilization of BMDCs in different tumor models underscores the need to further delineate the mechanisms by which they are activated and recruited, as well as their contributions to tumor neovascularization in these contexts. In particular, it will be critical to dissect in more detail the specific BMDC populations recruited into various tumors and better characterize their functional roles in promoting angiogenesis. In this article, we discuss general methods used to visualize and identify recruited BMDCs in two tumor models. We describe the use of bone marrow transplantation to facilitate the analysis of BMDCs, and also present methods that allow the investigator to begin characterizing and isolating the different subtypes of BMDCs found in tumors.

2. Methods for the Visualization of BMDCs in Tumors Adult bone marrow constitutes a reservoir of various stem and progenitor cells capable of contributing to the regeneration of a variety of tissues. Methods whereby genetically marked bone marrow is transplanted


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into tumor-bearing recipient mice have allowed for the positive identification and visualization of BMDCs within tumors. Not only do such approaches permit the tracking and quantification of BMDC recruitment in various tumor systems, they further allow for the isolation and functional characterization of the various BMDC populations in promoting tumor vessel growth. Bone marrow transplantation of mouse tumor models also offers the opportunity to study the angiogenic roles of the various BMDC subtypes in the context of genetic and pharmaceutical manipulations, as well as the mechanisms by which they are recruited to tumors. As more is understood about how BMDCs contribute to tumor neovascularization, it will be important to continue refining the identity of these cells and characterizing their functions.

2.1. Transplantation of GFP-expressing bone marrow to visualize BMDC in tumors Bone marrow transplantation techniques have been widely used in animal models to study a broad spectrum of biological phenomena. Genetic marking of bone marrow unique to that of the donor animal allows one to detect the specific presence of BMDCs within tissues of recipient animals. One example of this approach is to transplant bone marrow from male donor mice into female recipients (Ahn and Brown, 2008). The presence of cells originating from bone marrow can subsequently be detected by in situ hybridization with a Y-chromosome probe. In the following methods sections, we will use GFP-expressing bone marrow cells from b-actineGFP donor mice to fluorescently mark BMDCs transplanted into nonfluorescent recipient animals. This approach provides several advantages: (1) it allows for analysis of bone marrow reconstitution at any time by simply taking a peripheral blood sample and analyzing by flow cytometry; (2) no further staining of the BMDCs is necessary when performing histologic or immunofluorescent analyses; and (3) it permits the detection, fractionation, quantification, and marker characterization of tissue-incorporated BMDCs by flow cytometry. One of most important considerations when optimizing a bone marrow transplantation protocol is determination of the lethal irradiation dose. As different strains of mice tolerate varying levels of irradiation, it is a good idea to perform an initial kill curve to avoid overly lethal doses, or conversely, sublethal doses that result in chimeric bone marrow reconstitution. We routinely apply these methods to two mouse tumor models: C57BL6 b-actin EGFP donor mice with C57BL6 Rip1Tag2 recipient mice (pancreatic islet cell carcinoma) (Song et al., 2005), and FVBN b-actin-EGFP Rag1ko donor mice with FVBN Rag1ko recipient mice (brain tumor model in which glioblastoma cells are orthotopically injected (Du et al., 2008).

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2.1.1. Bone marrow transplantation Reagents C57BL6 b-actin EGFP donor mice C57BL6 Rip1Tag2 recipient mice FVBN b-actin-EGFP Rag1ko donor mice FVBN Rag1ko recipient mice Irradiator Antibiotic water: 1.1 g/l Neomycin sulfate (Sigma-Aldrich, St. Louis, MO, N-1876), 106 units/l polymyxin B sulfate (Sigma-Aldrich, P-1004) final concentration in water, filter sterilized Sterile phosphate-buffered saline lacking Ca2þ and Mg2þ (PBS) supplemented with 2% fetal bovine serum (Hyclone, Logan, UT) (PBS-FBS) 5-ml syringe (Becton Dickinson, Franklin Lakes, NJ) 25-gauge, 5/800 needle (Becton Dickinson) 70-mm nylon cell strainer (BD Biosciences, San Jose, CA, 352350) Ficoll density-gradient solution (density, 1.119) (Histopaque-1119, SigmaAldrich, 11191) Hemacytometer {1/2} cc, 28-gauge insulin syringes (Becton Dickinson) Methods

1. Prepare and give antibiotic water to recipient mice the day before irradiation. 2. Lethally irradiate recipient mice the day before bone marrow transfer. For initial studies, perform a killing curve to establish the lethal dose: Irradiate four mice per group at 800 R, 900 R, 1000 R, and 1200 R. Use the minimum dose at which all four animals in a group die 12 to 15 days after irradiation. The total lethal dose should be split into two equal half-doses 3 h apart. If animals die within 2 to 3 days after irradiation, the mice are likely suffering from gastrointestinal toxicity and the dose is too high. If the mice do not die after 15 days, then the dose is likely sublethal. 3. The next day, anesthetize and euthanize GFP donor mice. Using sterilized tools, dissect out femurs and tibias, removing the skin and as much muscle and soft tissue as possible. 4. Place bones in a dish containing sterile PBS on ice, and then move to a tissue culture hood for all remaining work. 5. Using another set of sterile tools, clean bones of any residual muscle or tissue, and then place cleaned bones into another dish containing sterile PBS. 6. Separate the femur from the tibia by cutting at the knee. Cut the ends of the bones to reveal the medullary compartment containing the marrow.


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7. Fill the 5-ml syringe with cold PBS-FBS and attach the 25-gauge needle. Flush out the bone marrow from each bone into a 50-ml conical tube on ice. Flush from both ends of each bone to ensure maximal recovery. 8. Repeat until all bones have been flushed. 9. Place a 70-mm nylon cell strainer onto a new 50-ml conical tube and pipette the flushed bone marrow through to remove debris. Adjust final volume to 20 ml with PBS-FBS if necessary. 10. Gently add to the bottom of the tube an equal volume (20 ml) of prewarmed, room-temperature Histopaque 1119. 11. Centrifuge for 30 min at 700 g, room temperature, with the brake set to off. 12. Pipette the interphase containing the bone marrow cells into a new 50-ml conical tube. Adjust the final volume to 50 ml with PBS-FBS. 13. Centrifuge for 8 min at 1200 rpm, 4 C. 14. Resuspend the pellet in 25 ml of PBS-FBS and count the cells on a hemacytometer. 15. Centrifuge again for 8 min at 1200 rpm, 4 C, and then resuspend the bone marrow cells at 1 to 2 106 cells/100 ml. 16. Intravenously transfer 1 to 2 106 bone marrow cells (100 ml) to each irradiated recipient mouse using {1/2} cc, 28-gauge insulin syringes.

2.2. Visualization of GFPþ BMDC subpopulations in tumors The investigator can visually verify the recruitment of BMDCs within tumors from GFP-bone marrow transplanted mice by the presence of GFP-positive cells in tumor sections. Quantification and spatial distribution of the GFP-positive BMDCs within tumors in relation to tumor vascular morphology can also be simultaneously analyzed. In addition, the identities of these BMDCs can begin to be determined by immunohistochemical or immunofluorescent staining of lineage specific markers. Here we discuss how tumor-bearing mice are perfused and the relevant tissues prepared for histologic analysis. We will focus on immunofluorescent characterization of the various CD45þ monocytic BMDC subtypes that have been found to considerably modulate tumor neovascularization in an orthotopic model of mouse GBMs (Du et al., 2008). 2.2.1. Cardiac perfusion, tissue fixation, and processing for immunohistochemical analysis Reagents Hemostat 2% 2,2,2-tribromoethanol (Avertin, Sigma-Aldrich) Phosphate-buffered saline lacking Ca2þ and Mg2þ (PBS)

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4% paraformaldehyde in PBS (PFA, Sigma-Aldrich) 25-gauge winged infusion set (Becton Dickinson) 10-ml syringe (Becton Dickinson) 20-ml syringe (Becton Dickinson) Fluorescein-conjugated (FITC) Lycopersicon esculentum lectin (Vector Laboratories, Burlingame, CA, FL-1171), diluted 1:1 with sterile PBS before use Rhodamine Ricinus communis Agglutinin I (Vector RL-1082) 30% sucrose in water, sterile filtered Optimal Cutting Temperature (OCT) Compound (Sakura Tissue-Tek, Torrance, CA, 4583) Dry ice Peel-A-Way tissue embedding molds (Polysciences, Warrington, PA, 18646A) Aluminum foil Cryostat Methods

1. Anesthetize tumor-bearing mouse by intraperitoneal injection of Avertin (250 to 400 mg/kg). 2. Preload 10 ml of PBS into 10-ml syringe and 20 ml of 4% PFA into 20-ml syringe. 3. Immerse tail in warm water to dilate veins in preparation for lectin infusion. 4. Intravenously inject 100 ml of FITC lectin (green fluorescence) or rhodamine agglutinin I (red fluorescence) via tail vein if desired. Wait 3 min for lectin to circulate before cardiac perfusion. 5. Tack mouse arms down with belly side up. Open peritoneal cavity and make an incision along each side of the sternum. Retract the sternum with the hemostat to allow for visualization of the heart. 6. Attach 25-gauge winged infusion set to 10-ml syringe containing PBS, and heart infuse PBS 1 minute. 7. Attach the same 25-gauge winged infusion set to the 20-ml syringe containing 4% PFA, and heart infuse PFA 3 min. Avoid removal of the needle from the heart during syringe transfer. 8. Collect the relevant tissue and place in 10 ml of 4% PFA in a 15-ml conical tube. Cover tube with aluminum foil if tissue contains GFP bone marrow or fluorescent lectin. Place on rocker at 4 C for 6 h. 9. Remove 4% PFA and wash three times with PBS. 10. Immerse tissue in 30% sucrose and place back on rocker at 4 C overnight (12 to 24 h).


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11. Wash tissue three times with PBS. 12. Squeeze a few drops of OCT into an empty tissue-embedding mold and carefully lay tissue in, avoiding air bubbles. Orient according to tumor location or other considerations. 13. Fill cryo-mold with OCT, completely covering the tissue. 14. Cover with aluminum foil and freeze each block on dry ice (10 to 15 min), and then store frozen blocks at –80 C. 15. Cut sections on a cryostat. 2.2.2. Immunohistochemical staining of bone marrow–derived monocytic modulatory cells in tumor sections Reagents PAP PEN hydrophobic barrier pen (The Binding Site, San Diego, AD100) Phosphate buffered saline, Ca2þ and Mg2þ free (PBS-CMF) Blocking buffer: 5% normal goat serum (NGS, Jackson Immunoresearch Lab, West Grove, PA) in PBS-CMF Staining buffer: 2% normal goat serum, 0.3% Triton-X-100 (Sigma-Aldrich) in PBS-CMF PBD wash buffer: 0.1% Tween-20 (Sigma-Aldrich) in PBS-CMF Humidified chamber Rat IgG2b anti-mouse CD45 antibody (BD Biosciences 550539, Clone 30-F11) Rat IgG2b anti-mouse VEGFR1 antibody (Imclone Systems, Clone MF1) Rat IgG1 anti-mouse Tie-2 antibody (eBioscience, San Diego, CA, 145987, Clone TEK4) Rat IgG2b anti-mouse CD11b antibody (BD Biosciences 550282, Clone M1/70) Rat IgG2b anti-mouse F4/80 antibody (Serotec, Oxford, UK, MCAP497, Clone C1:A3-1) Rat IgG2a anti-mouse CD31 antibody (BD Biosciences 553370, Clone MEC 13.3) Rat IgG isotype-matched control antibodies Alexa Fluor 594 goat anti-rat IgG (HþL) secondary detection antibody (Invitrogen-Molecular Probes, Eugene, OR, A-11007) or other desired fluorophore ProLong Gold antifade reagent with DAPI (Invitrogen-Molecular Probes, P-36931) Coverslips Clear nail polish 4% paraformaldehyde (PFA) in PBS Fluorescence microscope (Zeiss Axiophot, Carl Zeiss, Germany)

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Methods

1. Wash sections 3 times, 5 minutes per wash, in PBS-CMF. 2. Draw circles around tissue on the slide with the PAP PEN to create a hydrophobic barrier. Pipette subsequent solutions within this ring. 3. Block in blocking buffer for 30 min at room temperature. 4. Incubate with the desired primary antibody (rat anti-mouse CD45, VEGFR1, Tie-2, CD11b, or F4/80) in staining buffer at 4 C overnight in a humidified chamber. 5. Use rat IgG isotype control antibodies as negative controls on duplicate sections. 6. Wash with PBD 3 times, 5 minutes per wash. 7. Wash with PBS-CMF for 5 min. 8. Incubate with secondary Alexa Fluor 594-conjugated goat antirat antibody (1:200 dilution) in staining buffer for 1 h at room temperature in a humidified chamber. 9. Wash with PBD twice, 5 minutes per wash. 10. Wash with PBS-CMF for 5 min. 11. Mount sections with one or two drops of Prolong Gold antifade reagent with DAPI. Cover with a coverslip, to avoid trapping air bubbles. 12. Cure for 24 h at room temperature in the dark. 13. Seal edges with clear nail polish. 14. Analyze with a fluorescent microscope and store at 4 C short term or – 20 C or colder long term. 15. BMD monocytic modulatory cells can be identified by positive staining of the described markers along with GFP positivity if animals have been transplanted with GFP bone marrow. Fig. 3.1A shows an example of GFPþ BMDCs found within an orthotopic glioblastoma, and Fig. 3.1B reveals CD11b staining of GFPþ BMDCs. Immunohistochemical staining of vasculature 1. Postfix sections in 4% PFA for 20 min at 4 C, protected from light. 2. Wash sections 3 5 min in PBS-CMF. 3. Draw circles around tissue on the slide with the PAP PEN to create a hydrophobic barrier. Pipette subsequent solutions within this ring. 4. Block in blocking buffer for 30 min at room temperature. 5. Incubate with rat anti-mouse CD31 antibody (1:100 dilution) in staining buffer at room temperature overnight in a humidified chamber. 6. Use rat IgG isotype control antibodies as a negative control on duplicate sections. 7. Wash with PBD 3 5 min. 8. Wash with PBS-CMF for 5 min. 9. Incubate with secondary Alexa Fluor 594 (or other desire fluorophore)– conjugated goat antirat antibody (1:200 dilution) in staining buffer for 1 h at room temperature in a humidified chamber.

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A

B BM-GFP-cells

DAPI

BM-GFP-cells

CD11b

C BM-GFP-cells CD31 DAPI

Figure 3.1 Visualization and immunohistochemical staining of BMDCs in tumors. Orthotopic glioblastomas implanted into GFP-bone marrow transplanted mice were sectioned, stained, and analyzed by fluorescence microscopy. (A) GFPþ bone marrow cells (green) are readily detected within the tumor. Scale bar, 50 mm. (B) Staining of tumor sections identifies a fraction of GFPþ bone marrow cells (green) recruited to the tumor as CD11bþ myeloid cells (red and green merge). Scale bar,15 mm. (C) Staining of the tumor vasculature with CD31antibodies (red) reveals dilated tumor vessel morphology, while recruited GFPþ bone marrow cells (green) reside among tumor cells (yellow arrowheads) and are also incorporated into the vasculature (white arrowheads). Scale bar,15 mm.

10. Wash with PBD 2 5 min. 11. Wash with PBS-CMF for 5 min. 12. Mount sections with one or two drops of Prolong Gold antifade reagent with DAPI. Cover with a coverslip, and avoid trapping air bubbles. 13. Cure for 24 h at room temperature in the dark. 14. Seal edges with clear nail polish. 15. Analyze with fluorescent microscope and store at 4 C short-term or – 20 C or colder long term. 16. Vasculature can be visualized without immunohistochemical staining if tissues were perfused with fluorescent lectin prior to animal euthanization. CD31 staining enhances and allows for complete staining of the vasculature regardless of vessel functionality. Fig. 3.1C shows staining of tumor vasculature with CD31, revealing hyperdilated vessels alongside GFPþ BMDCs.

2.3. Quantification of GFPþ BMDC subpopulations in tumors While tumor-recruited BMDCs can be quantified by counting the number of GFP-positive cells (and relevant marker stained cells) in tissue sections, a more efficient and versatile way to quantify and characterize BMDCs is by flow cytometry. FACS analysis allows for GFP-positive BMDCs to be quickly identified and quantified, and further fractionation of BMDC

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subtypes can be easily performed by staining with the relevant markers. In the following we discuss the methods used to analyze BMDCs in tumor tissues by flow cytometry, specifically detecting for endothelial progenitor cells, pericyte progenitor cells, and various CD45þ monocytic modulatory cells (Du et al., 2008). 2.3.1. Flow cytometric analysis of bone marrow–derived vascular progenitor and modulatory cells in tumors Reagents 2% 2,2,2-tribromoethanol (Avertin, Sigma-Aldrich) Collagenase digestion buffer: 0.2 g bovine serum albumin (BSA, Sigma Aldrich, B4287) 12,500 units collagenase II (Worthington Biochemical, Lakewood, NJ, LS004176) 12,500 units collagenase IV (Worthington Biochemical, LS004188) 20 ml DNase I (RNase, Proteinase-Free, Worthington Biochemical, LS006333) 20 ml PBS 70-mm nylon cell strainers (BD Biosciences 352350) 5-ml syringes BD Pharm Lyse 10, red-blood-cell lysing buffer (BD Biosciences 555899) Phosphate-buffered saline lacking Ca2þ and Mg2þ supplemented with 5% fetal bovine serum (Hyclone SH30070) (5% FBS-PBS) Phosphate-buffered saline lacking Ca2þ and Mg2þ supplemented with 2% fetal bovine serum (Hyclone SH30070) (2% FBS-PBS) Fc Block: rat anti-mouse CD16/CD32 (BD Biosciences 553142) Phycoerythrin (PE)-labeled, isotype-matched control Allophycocyanin (APC)-labeled isotype-matched control PE-Cy7–labeled, isotype-matched control Alexa Fluor 647–labeled, isotype-matched control Antibodies for detection of monocytic modulatory cells: PE-Cy7-conjugated rat IgG2b anti-mouse CD45 antibody (eBioscience 25-0451, clone 30-F11) APC-conjugated rat IgG2b anti-mouse CXCR4 antibody (BD Biosciences 558644, clone 2B11) PE-conjugated rat IgG1 anti-mouse Tie-2 antibody (eBioscience 12-5987, clone TEK4) PE-conjugated rat IgG2a anti-mouse F4/80 antibody (eBioscience 12-4801, clone BM8) PE-conjugated rat IgG2b anti-mouse CD11b antibody (eBioscience 12-0112, clone M1/70)


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Biotinylated rat IgG2b anti-VEGFR1 antibody clone MF1 (Imclone Systems), biotinylation of antibody must be performed (FluoReporter Mini-biotin-XX Protein Labeling Kit, Invitrogen-Molecular Probes F6347) Streptavidin-PE (eBioscience 12-4317) Antibodies for detection of endothelial progenitor cells: PE-conjugated rat IgG2a anti-mouse VEGFR2 antibody (eBioscience 12-5821, clone Avas12a1) Alexa 647-conjugated rat IgG1 anti-mouse VE-cadherin antibody (eBioscience 51-1441, clone BV13) Antibodies for detection of pericyte progenitor cells: PE-conjugated rat IgG2a anti-mouse PDGFRb antibody (eBioscience 12-1402, clone APB5) APC-conjugated rat IgG2a anti-mouse Sca-1 antibody (eBioscience 17-5981, clone D7) BD Via-Probe cell viability solution (ready-to-use 7-AAD solution, BD Biosciences 555815) BD LSR II flow cytometer (BD Biosciences) 5-ml polystyrene round-bottom tubes (Becton Dickinson) Methods

1. Anesthetize bone marrow–transplanted, tumor-bearing mice by intraperitoneal injection of Avertin (250 to 400 mg/kg). 2. Carefully dissect out tumor and place in a dish containing PBS on ice. 3. Finely mince tumor with a razor blade on ice in a clean and dry dish. 4. Transfer minced tumor into 10 ml of collagenase digestion buffer in a 50-ml conical tube. 5. Incubate in a 37 C water bath for 13 min, with manual shaking/ stirring every 2 to 3 min. 6. Immediately place on ice and stop digestion by adding 5% FBS-PBS up to 40 ml. 7. Strain cells through a 70-mm nylon cell strainer into a new 50-ml conical tube. 8. Remove the plunger from a 5-ml syringe and vigorously rub and scrape any residual chunks of tumor tissue on top of the strainer against the mesh. Rinse with 10 ml of 5% FBS-PBS to maximize tumor cell recovery. 9. Centrifuge at 1200 rpm for 5 min at 4 C. 10. Aspirate supernatant and resuspend cell pellet in 1 ml of 1 Pharm Lyse (diluted in water). Place on ice for 3 min, and then stop reaction by adding 5% FBS-PBS to 50 ml.

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11. Centrifuge at 1200 rpm for 5 min at 4 C. 12. Resuspend cells in 2% FBS-PBS according to the cell yield and the number of controls and staining combinations required; 0.5 to 1 106 cells per 100 ml are recommended. Split the cells in 100-ml aliquots into 5-ml polystyrene, round-bottom tubes. 13. Set aside single color- and isotype-matched control samples. Add Fc block to remaining samples at 1:50 dilution. 14. Apply primary antibody multicolor combinations according to Table 3.1 to detect different populations of BMD CD45þ monocytes, endothelial progenitor cells, or pericyte progenitor cells. The FITC channel is reserved for identifying GFP-positive BMDCs in the tumor sample, and 7-AAD is reserved for staining and exclusion of dead cells. 15. Apply appropriate antibodies to single color controls and isotypematched controls. 16. Incubate primary antibody combinations on ice for 30 min, protected from light. 17. Wash cells with 3 ml of 2% FBS-PBS, and centrifuge at 1200 rpm for 5 min at 4 C. 18. Resuspend cells in 200 ml of 2% FBS-PBS, except for staining of VEGFR1þ hemangiocytes. 19. For detection of VEGFR1þ hemangiocytes, resuspend cells in 400 ml of 2% FBS-PBS and add streptavidin-PE at a 1:400 dilution. Incubate on ice for 30 min, protected from light. Following secondary staining, wash and centrifuge as described in Step 19. Resuspend in 200 ml of 2% FBS-PBS. 20. Prior to analysis of each sample, add 20 ml of BD Via-Probe cell viability solution (7-AAD) to each tube and incubate for 10 min. Analyze samples on the flow cytometer. 21. Exclude dead cells and gate for GFPþ BMDCs. Use single-color and isotype-matched controls to set gates for marker analysis. Fig. 3.2 demonstrates the identification and analysis of GFPþ CD45þ CD11bþ BMDCs in an orthotopic glioblastoma tumor by FACS analysis.

3. Analysis of Pericyte Progenitors in Tumors RIP1Tag2 mice express the viral SV40Tag oncoproteins under the control of the rat insulin gene II promoter in the pancreatic b cells of the 400 islets of Langerhans, setting in motion a multistep pathway to beta islet cell carcinoma as characterized by the temporal and synchronistic appearance of distinctive lesional stages in all transgenic mice (Bergers et al., 1998; Hanahan, 1985). Aberrant hyperproliferation of the b cells starts at about 3 to 4 weeks of age, producing hyperplastic and dysplastic

Table 3.1 Antibody and fluorochrome combinations for detection and analysis of BMDCs by flow cytometry.

Color

TEMs

FITC PE PE-Cy7 APC 7-AAD

GFP-BM Tie-2 CD45 CXCR4 Dead cells

Pericyte progenitor cells

Monocytic accessory modulatory cells

Endothelial progenitor cells

Tumor- associated macrophages

Myeloid cells

Hemangiocytes

Color

GFP-BM F4/80 CD45 CXCR4 Dead cells

GFP-BM CD11b CD45 CXCR4 Dead cells

GFP-BM VEGFR1 B* CD45 CXCR4 Dead cells

FITC PE

GFP-BM VEGFR2

GFP-BM PDGFR b

APC/Alexa 647 7-AAD

VE-cadherin Dead cells

Sca-1 Dead cells

B* denotes biotinylated antibody requiring secondary detection.

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Exclude 7-AAD + dead cells 250 K

Forward scatter

Side scatter

77.1% 200 K 150 K 100 K 50 K

200 K 150 K 100 K 50 K

0

0 0

Gate for GFP + BMDCs 250 K

85.3%

Forward scatter

Gate initial cell population 250 K

50 K 100 K 150 K 200 K 250 K

150 K 100 K 50 K 0

5

1000 10,000 1 ⫻ 0

0 100

Forward scatter

7.24%

200 K

0 100

7-AAD

1000 10,000 1 ⫻ 105

GFP

Set gating parameters for marker expression CD45 positive control

CD45 isotype control 1 ⫻ 105

1 ⫻ 105

0.96%

10,000

10,000

1000

1000

100

100

86%

% CD45 + CD11b + GFP BMDCs in tumor 1 ⫻ 105

0 1000 10,000 1 ⫻ 05

0 100

CD11b isotype control

CD11b positive control

1 ⫻ 105

1 ⫻ 105

10,000

10,000

1000

1000

100 0 0 100

1000 10,000 1 ⫻ 0

0 0

32.3%

0 100

100

1000

10,000 1 ⫻ 105

CD11b

0 5

1000

100

100

1.91%

10,000

5 1000 10,000 1 ⫻ 0

CD45

CD45

26.9%

0 0 100

5

1000 10,000 1 ⫻ 0

CD11b

Figure 3.2 Flow cytometric analysis and characterization of BMDCs in tumors. Orthotopic glioblastomas implanted into GFP-bone marrow transplanted mice were resected and processed into a single cell suspension for flow cytometric analysis. In this example, the tumor cell suspension was stained with PE-Cy7 conjugated CD45 antibodies and PE conjugated CD11b antibodies to detect and quantify BMD myeloid cells, and then counterstained with 7-AAD. During anaylsis, cellular debris and 7-AADþ dead cells were sequentially gated out, followed by selective gating of GFPþ cells within the tumor suspension for interrogation of CD45 and CD11b expression. Gating parameters for positive CD45 and CD11b expression were set by staining bone marrow cells with isotype-matched control antibodies or CD45 and CD11b antibodies alone (lower left corner). Note that the majority of GFPþ BMDCs recruited in this tumor model are CD45þ monocytes, of which about one-fourth of them are CD11bþ myeloid cells.

islets, initially with a quiescent vasculature. Then, at 6 to 7 weeks, angiogenesis is switched on in a subset of the dysplastic islets. Subsequently, solid tumors form at about 10 to 11 weeks as encapsulated tumors and invasive carcinomas. The mice die between 13.5 to 15 weeks of age with substantial tumor burden and consequent hyperinsulinemia. Tumor pericytes originate not only from a pre-existing pool of pericytes, but also by maturation of undifferentiated progenitors recruited to the


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newly formed tumor blood vessels. In pancreatic islet tumors of Rip1Tag2 mice, PDGFRb is expressed in perivascular cells closely associated with the tumor vasculature, while the PDGF ligands B and D for PDGFRb are expressed in tumor endothelial cells, reflective of paracrine communication pathways similar to the vascular processes during development (Bergers et al., 2003; Hellstrom et al., 1999; Song et al., 2005). One advantage of studying pericyte progenitors in pancreatic islet tumors is based on the fact that PDGFRb is exclusively expressed in pericytes in these tumors and that these tumors are rather devoid of stromal fibroblasts that can also express PDGFRb. We have identified PDGFRbþ pericyte progenitors (PPPs) closely associated with angiogenic endothelial cells in Rip1Tag2 tumors. Furthermore, a subset of these cells is recruited from the bone marrow, which indicates that recruitment of BMD cells to sites of a growing vasculature is not limited to endothelial cells, but can also include pericytes (Song et al., 2005). Importantly, three populations of tumor pericytes can be detected in tumors, each reflecting a distinct differentiation stage: (1) PPPs that are positive for PDGFRb but negative for the mature pericyte markers NG2, desmin, and a-SMA; (2) intermediate pericytes that express PDGFRb and the mature pericyte markers NG2, desmin, and a-SMA; and (3) mature pericytes that are positive for the mature pericyte markers NG2, desmin, and a-SMA, but have lost the expression of PDGFRb. The three populations of tumor pericytes can be detected by FACS and immunohistochemistry of tumor sections. Pericyte progenitors can also be isolated from tumor cell suspensions using magnetic beads based on PDGFRb expression. They can later be cultured as monoculture in vitro to allow their differentiation into mature pericytes. However, we have demonstrated that in order to upregulate desmin expression, it is necessary to co-culture pericyte progenitors with endothelial cells. We use the 3D Matrigel gel as an in vitro assay, which permits the investigator to observe the interactions between endothelial cells and pericyte progenitors during vessel formation.

3.1. Detection of pericyte progenitors and pericytes in tumors Prior to sacrifice, mice are injected intravenously with FITC-conjugated tomato lectin to detect the blood vessels, and tumor tissues are fixed and processed as described in Section 2.2.1. Tumor pericytes are detected by immunostaining of 16-mm frozen sections with antibodies against PDGFRb, NG2, desmin, and a-SMA. Commercially available desmin and a-SMA antibodies are of the mouse isotype. Hence, to avoid unspecific background, antibodies are conjugated to the AlexaFluor647 fluorochrome just prior to immunostaining using the Zenon AlexaFluor647 Mouse IgG-specific Labeling Kit (Invitrogen), according to the manufacturer’s instructions.

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Reagents

PAP PEN hydrophobic barrier pen (The Binding Site) Phosphate buffered saline, Ca2þ and Mg2þ free (PBS) Fixation buffer: 4% paraformaldehyde (PFA) in PBS Blocking/permeabilization buffer: PBS supplemented with 5% normal goat serum (NGS; Jackson Immunoresearch Lab) and 0.3% Triton X-100 (Sigma-Aldrich) Incubation buffer: PBS supplemented with 2% NGS and 0.3% Triton X-100 Washing buffer: PBS supplemented with 0.1% Tween-20 (Sigma-Aldrich) Zenon AlexaFluor647 Mouse IgG1 Labeling Kit (Invitrogen) Zenon AlexaFluor647 Mouse IgG2a Labeling Kit (Invitrogen) Rabbit polyclonal anti-NG2 chondroitin sulfate proteoglycan antibody (Chemicon, Temecula, CA), used at 2 mg/ml Mouse IgG1 monoclonal anti-desmin antibody (Clone D33, DAKO, Carpinteria, CA), used at 1:200 Mouse IgG2a monoclonal anti-alpha smooth muscle actin (a-SMA) antibody (Clone 1A4, DAKO), used at 1:500 Rat IgG2a monoclonal, anti-PDGFRb antibody (Clone APB5, eBiosciences), used at 10 mg/ml Rabbit IgG isotype-matched control ( Jackson Immunoresearch Lab) Mouse IgG1 and IgG2a isotype-matched controls (BD Biosciences) Rat IgG2a isotype-matched control (eBiosciences) AlexaFluor647-conjugated goat anti-rabbit IgG (HþL) antibody (Invitrogen-Molecular Probes), used at 1:400 AlexaFluor546-conjugated goat anti-rat IgG (HþL) antibody (InvitrogenMolecular Probes), use at 1:400 ProLong Gold antifade reagent with DAPI nucleic acid stain (InvitrogenMolecular Probes) Fluorescence confocal microscope (Zeiss LSM510, Carl Zeiss, Germany) Methods

1. Draw circles around tissue on the slide with the PAP PEN to create a hydrophobic barrier. Pipette subsequent solutions within this ring. 2. Post-fix the frozen sections in 4% PFA/PBS for 10 min at room temperature. 3. Wash the tissue sections three times in PBS for 5 min at room temperature. 4. Block the tissues with 5%NGS/0.3% Triton X-100/PBS for 30 min at room temperature. 5. Wash the tissue sections once in PBS for 5 min at room temperature. 6. Incubate the sections with the primary antibodies diluted in the incubation buffer overnight at 4 C. Use isotype-matched control antibodies as a negative control on duplicate sections.

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7. Wash the tissue sections twice in PBS/0.1% Tween-20 for 5 min at room temperature. 8. Wash the tissue sections in PBS for 5 min at room temperature. 9. Incubate the sections with the secondary antibodies diluted in the incubation buffer for 1 h at room temperature. 10. Wash the tissue sections twice in PBS/0.1% Tween-20 for 5 min at room temperature. 11. Wash the tissue sections in PBS for 5 min at room temperature. 12. Mount the slides with ProLong Gold antifade reagent with DAPI nucleic acid stain included. 13. Evaluate immunostaining with a fluorescence confocal microscope. Fig. 3.3 demonstrates pericyte staining of a Rip1Tag2 tumor with desmin and PEGFRb. 14. Isolation of pericyte progenitors and pericytes in tumors. 15. PDGFRbþ pericyte progenitors are isolated from single-cell suspensions of Rip1Tag2 tumors. Given that this progenitor population represents a small percentage of the total tumor cell population (2%), it is necessary to start with a high number of cells, that is, five to seven mice per isolation. 3.1.1. Preparation of single-cell suspension from Rip1Tag2 tumors Reagents Sterile conical 50-ml tubes Sterile Petri dishes One disposable 70-mm cell strainer (Becton Dickinson) One sterile razor blade Lectin

Desmin

Lectin PDGFRβ

Merge

Figure 3.3 Visualization of tumor pericytes on tissue sections.13.5-week-old Rip1Tag2 mice were injected intravenously with FITC-conjugated tomato lectin that binds to blood vessels and allows their visualization in pancreatic tumors (green).Tumor sections are then stained with antibodies against PDGFRb (red) and desmin (blue). Pictures were acquired using a fluorescent confocal microscope. Scale bar, 50 mm.

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Water bath (37 C) 2% 2,2,2-tribromoethanol (Avertin, Sigma-Aldrich) Sterile phosphate-buffered saline without Ca2þ and Mg2þ (PBS) Sterile PBS supplemented with 63.7 mg/ml penicillin, 100 mg/ml streptomycin, and 2.5 mg/ml fungizone Collagenase digestion buffer: 0.2 g bovine serum albumin (BSA, Sigma Aldrich) 12,500 units collagenase II (Worthington Biochemical) 12,500 units collagenase IV (Worthington Biochemical) 20 ml DNase I (RNase, proteinase-free, Worthington Biochemical) 20 ml PBS Phosphate-buffered saline lacking Ca2þ and Mg2þ supplemented with 5% fetal bovine serum (Hyclone) (5% FBS-PBS) Phosphate-buffered saline lacking Ca2þ and Mg2þ supplemented with 2% fetal bovine serum (Hyclone) and 1 mM EDTA (Sigma-Aldrich) (2% FBS-PBS-EDTA) Ficoll density gradient solution (density 1.119) (Histopaque-1119, SigmaAldrich) 0.4% trypan blue solution (Sigma-Aldrich) Methods

1. Euthanize 13-week-old Rip1Tag2 mice bearing substantial tumor burden by intraperitoneal injection of Avertin (250 to 400 mg/kg). 2. Harvest the pancreases and wash them in PBS containing antibiotics and fungizone and place them in fresh solution on ice. 3. Carefully dissect the tumors out of the pancreas on ice and place them in fresh PBS containing antibiotics and fungizone. 4. Place the tumors in a clean, sterile Petri dish, and finely mince them with a sterile razor blade. Add the minced tumors to 20 ml of collagenase digestion buffer. 5. Incubate in a 37 C water bath for 13 min, with manual shaking/ stirring every 2 to 3 min. 6. Add 20 ml of 5% FBS-PBS and mechanically dissociate the digested tissue by pipetting up and down. 7. Place a 70-mm disposable cell strainer on top of a 50-ml conical tube and filter the digested tumor solution. 8. Remove the plunger from a 5-ml syringe and vigorously rub and scrape any residual chunks of tumor tissue on top of the strainer against the mesh. Rinse with 10 ml of 5% FBS-PBS to maximize tumor cell recovery. 9. Centrifuge at 1200 rpm for 8 min at 4 C. 10. Resuspend the cell pellet into 10 ml of 2% FBS-PBS-EDTA at room temperature. 11. Transfer 10 ml of Histopaque-1119 into a 50-ml conical tube.


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12. Carefully layer 10 ml of tumor cell suspension onto the Histopaque1119 layer 13. Centrifuge at 700 g for 30 min at room temperature with the brake set to off. 14. Red blood cells pellet at the bottom of the tube while cells from the tumors band at the interface between the Histopaque-1119 and the upper layer. Carefully aspirate the cells with a Pasteur pipette without disturbing the interface and transfer into a 50-ml conical tube. 15. Bring the cell suspension to a total volume of 50 ml by adding 2% FBSPBS-EDTA. Transfer 10 ml into an Eppendorf tube for cell counting. Centrifuge the rest at 1200 rpm for 8 min at 4 C. 16. Cell counting: Mix 10 ml of tumor cell suspension and 10 ml of trypan blue (final dilution 1:2). Determine the total number of cells using a hemacytometer chamber.

3.2. Isolation of PDGFRbþ pericytes from Rip1Tag2 tumor cell suspension Reagents

One sterile, polystyrene 5-ml, round-bottom tube (Becton Dickinson) Phosphate-buffered saline lacking Ca2þ and Mg2þ supplemented with 2% fetal bovine serum (Hyclone) and 1 mM EDTA (Sigma-Aldrich) (2% FBS-PBS-EDTA) EasySep magnet (cat. 18000, Stem Cell Technologies, Canada) EasySep Biotin Selection Kit (cat. 18556, Stem Cell Technologies). Contains the mouse-specific Fc Receptor blocking antibody, the EasySep Biotin Selection Cocktail and EasySep Magnetic Nanoparticles Biotin-conjugated Rat IgG2a anti-PDGFRb antibody (Clone APB5, eBiosciences), used at 7.5 mg/ml Methods

1. PDGFRbþ pericyte progenitors are isolated from Rip1Tag2 mouse tumors using the EasySep Biotin Selection Kit according to the exact manufacturer’s instructions, as described below. 2. Resuspend the cell pellet at 2 108 cells/ml in 2% FBS-PBS-EDTA and place the cell suspension into a 5-ml, round-bottom polystyrene tube. 3. Add mouse-specific Fc receptor blocking antibody at 10 ml/ml. Mix well. 4. Add biotin-conjugated rat anti-PDGFRb antibody. Mix well and incubate at room temperature for 15 min. 5. Add EasySep Biotin Selection Cocktail at 100 ml/ml. Mix well and incubate at room temperature for 15 min. 6. Add EasySep Magnetic Nanoparticles at 50 ml/ml. Mix well and incubate at room temperature for 10 min.

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7. Bring the cell suspension to a total volume of 2.5 ml by adding 2% FBSPBS-EDTA. Mix the cells in the tube by gently pipetting up and down. Place the tube (without cap) into the magnet. Set aside for 5 min. 8. Pick up the magnet, and in one continuous motion invert the magnet and tube, pouring off the supernatant fraction. The magnetically labeled cells remain inside the tube. Leave the magnet and tube in inverted position for 2 to 3 s, and then return to upright position. 9. Remove the tube from the magnet and add 2.5 ml 2% FBS-PBSEDTA. Mix the cells by gently pipetting up and down. Place the tube back in the magnet and set aside for 5 min. 10. Repeat Steps 7 and 8 twice, and then Step 7 once more, for a total of four separations in the magnet. Remove the tube from the magnet and resuspend cells in differentiation medium.

3.3. In vitro differentiation of pericyte progenitors PDGFRbþ pericyte progenitors isolated from pancreatic Rip1Tag2 tumors have the capacity to differentiate into mature pericytes in vitro (Song et al., 2005). Here we describe procedures to differentiate pericyte progenitors both under monoculture conditions, as well as co-cultures with endothelial cells on a 3D Matrigel matrix. 3.3.1. In vitro differentiation of pericyte progenitors as monoculture Reagents Eight-well Labtek Permanox chamber slides (Nalge Nunc Int., Rochester, NY) Coating solution: PBS supplemented with 0.1% gelatin from porcine skin, type A (Sigma-Aldrich) Differentiation medium: mesenchymal stem cell basal medium (MSCBM, Lonza Group, Switzerland) supplemented with MSCBM SingleQuot Kit (Lonza Group) and 5 ng/ml of recombinant human TGFb1 (R&D Systems, Minneapolis, MN). Methods

1. Incubate an eight-well Labtek Permanox slide chamber with 0.1% gelatin/PBS for 1 h at 37 C. 2. Remove the uncoated 0.1% gelatin/PBS. Wash once with PBS. 3. Transfer freshly isolated PDGFRbþ pericyte progenitors to gelatincoated slide chamber at a density of 10,000 cells per well in differentiation medium.


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4. Culture the cells for 7 days at 37 C in 5% CO2. Replace half the medium every second day. 3.3.2. Differentiation culture of PDGFRbþ pericyte progenitors with endothelial co-cultures in 3D Matrigel Pericytes and endothelial cells are known to closely interact in vivo through both direct contact as well as paracrine signaling mechanisms. Therefore, co-cultures of pericyte progenitors with endothelial cells in vitro may more closely mimic the maturation and behavior of pericytes in vivo. Indeed, we have observed that desmin is upregulated in mature pericytes only under these co-culture conditions. Reagents

Four-well Labtek Permanox chamber slides (Nalge Nunc Int.) CDC/EU.HMEC-1 (HMEC-1) immortalized human microvascular endothelial cells (Ades et al., 1992) PDGFRbþ pericyte progenitor cells (as isolated in Section 3.2) CellTracker fluorescent cell-labeling probes in color of choice (Molecular Probes-Invitrogen) Endothelial growth medium: MCDB131 (Gibco-Invitrogen) supplemented with 10% FBS, 1 L-glutamine Serum-free MCDB131 medium (no additives) Endothelial Cell Medium-2 (EGM-2, Lonza Group) supplemented with EGM-2 SingleQuot Kit, 63.7 mg/ml penicillin, 100 mg/ml streptomycin, 1 antibiotic antimycotic stabilized solution (AASS, Sigma-Aldrich) Pericyte differentiation medium: mesenchymal stem cell basal medium (MSCBM, Lonza Group) supplemented with MSCBM SingleQuot Kit and 5 ng/ml of recombinant human TGFb1 (R&D Systems) Cold sterile PBS PBS supplemented with 0.5 mM EDTA (Sigma-Aldrich) (PBS-EDTA) Growth-factor-reduced (GFR)-Matrigel (Becton Dickinson) Hemacytometer Chilled sterile pipette tips 4% paraformaldehyde in PBS 0.1 M glycine Methods

1. Culture HMEC-1 immortalized human microvascular endothelial cells in complete MCDB131 endothelial culture medium. 2. Thaw GFR-Matrigel at 4 C on ice overnight.

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3. Under sterile conditions in the tissue culture hood, place GFR-Matrigel and chamber slides on ice. Using chilled pipette tips, mix Matrigel to homogeneity. 4. Pipette enough Matrigel to cover the surface of each chamber on the slide and make a thick gel (150 to 200 ml per square centimeter of growth surface area). 5. Place slides and Matrigel in the tissue culture incubator at 37 C for 30 min. After incubation, check to see that Matrigel has set. 6. Wash HMEC-1 cells with cold PBS and detach from the plates by adding 2 ml of PBS-EDTA. Incubate at 37 C for 1-2 min. After cells have lifted off plate, add 10 ml complete growth medium containing 10% FBS. 7. Spin down cells at 1200 rpm for 5 min, and resuspend the pellet in 1 ml of complete growth medium. 8. Label cells with CellTracker fluorescent probes at a 10-mM working concentration and incubate for 15 min at 37 C. 9. Spin endothelial cells down and resuspend in 10 ml serum-free medium. Incubate for 30 min at 37 C. 10. Spin labeled endothelial cells down and resuspend in 5 to 10 ml of complete EGM-2 medium. 11. Count cells with a hemacytometer and seed 50,000 labeled endothelial cells per square centimeter of growth surface on top of the solidified Matrigel. Add enough complete medium if necessary to ensure complete coverage of the well (final volume 0.5 ml for a four-chamber slide). 12. Place chamber slide containing Matrigel matrices and labeled endothelial cells overnight in the tissue culture incubator. Endothelial tube-like structures should form within a few hours. 13. Isolate PDGFRbþ pericyte progenitor cells as described in Section 3.2. (Optional: Fluorescently label PDGFRbþ pericyte progenitor cells with CellTracker in a color different from that used to label endothelial cells.) Resuspend pericyte progenitor cells in complete MSCBM medium. 14. Aspirate medium from chamber slides, being careful not to disturb the Matrigel bed and endothelial tubes. 15. Seed pericyte progenitor cells at one-third the number of endothelial cells seeded (16,666 pericyte progenitor cells per square centimeter of growth surface) on top of the Matrigel bed containing endothelial tubes. 16. Allow co-cultures to incubate for 3 to 7 days in tissue culture incubator. 17. Rinse cultures with PBS and fix by covering the Matrigel/endothelial cell/pericyte co-cultures with 4% paraformaldehyde for 15 min at room temperature. 18. Rinse with 0.1 M glycine 2 5 min.


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19. Cover chambers with OCT and place on dry ice to freeze. 20. Pry frozen Matrigel OCT blocks from the chambers and store wrapped in foil at –70 C until use. 21. Cut sections on a cryostat, then proceed to stain for mature pericyte markers as described below. 3.3.3. Detection of mature pericytes after differentiation After 7 days of culture or endothelial cell co-culture in differentiation medium, PDGFRbþ pericyte progenitors have acquired a mature pericyte phenotype identified by (1) a changed morphology from round to elongated, and (2) expression of the mature pericyte markers a-SMA, desmin, and NG2. Reagents

Fixation buffer: 4% paraformaldehyde (PFA) in PBS Blocking/permeabilization buffer: PBS supplemented with 5% normal goat serum (NGS, Jackson Immunoresearch Lab) and 0.3% Triton X-100 (Sigma-Aldrich) Incubation buffer: PBS supplemented with 2% NGS and 0.3% Triton X-100 Vector M.O.M. (mouse-on-mouse) blocking reagent (Vectors Lab) Rabbit IgG polyclonal anti-NG2 chondroitin sulfate proteoglycan antibody (Chemicon), used at 2 mg/ml Mouse IgG1 monoclonal anti-desmin antibody (Clone D33, DAKO), used at 1:200 Mouse IgG2a monoclonal anti-a-SMA antibody (Clone 1A4, DAKO), used at 1:500 Rat IgG2a monoclonal anti-PDGFRb antibody (Clone APB5, eBiosciences), used at 10 mg/ml Rabbit IgG isotype-matched control (Jackson Immunoresearch Lab) Mouse IgG1 and IgG2a isotype-matched controls (BD Biosciences) Rat IgG2a isotype-matched control (eBiosciences) AlexaFluor594-conjugated goat anti-mouse IgG (HþL) antibody (Invitrogen-Molecular Probes), used at 1:200 AlexaFluor594-conjugated goat anti-rabbit IgG (HþL) antibody (Invitrogen-Molecular Probes), used at 1:200 AlexaFluor488-conjugated goat anti-rat IgG (HþL) antibody (InvitrogenMolecular Probes), used at 1:200 ProLong Gold antifade reagent with DAPI nucleic acid stain (InvitrogenMolecular Probes) Fluorescence microscope (Zeiss Axiophot, Carl Zeiss, Germany)

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DAPI PDGFRb NG2

DAPI PDGFRb a-SMA

DAPI PDGFRb

Desmin

Figure 3.4 PDGFRbþ pericyte progenitors differentiate into mature pericytes in vitro. PDGFRbþ cells isolated from Rip1Tag2 tumors were cultured in vitro in complete MSCBM medium. After 7 days, cells were fixed and immunostained with antibodies against PDGFRb (green) and NG2, desmin, or a-SMA (red, as indicated).

Figure 3.5 Differentiation of PDGFRbþ pericyte progenitors under co-culture conditions with endothelial cells. Endothelial cells were labeled with CellTracker Green dye and cultured on a 3D Matrigel matrix to form endothelial tube-like structures. Pericyte progenitors were co-cultured with the endothelial cells in differentiation medium for 7 days. Staining for the mature pericyte marker NG2 (red) reveals mature pericytes wrapped around the endothelial tubes. Nuclei are counterstained with DAPI (blue).


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1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Wash cultured PDGFRbþ pericyte progenitors twice in PBS. Fix the cells in 4%PFA/PBS for 15 min at room temperature. Wash the cells three times in PBS, 5 min each. If immunostaining with antibodies against a-SMA or desmin, block the cells with Vector M.O.M. diluted in PBS according to the manufacturer’s instructions. Incubate for 30 min at room temperature. Briefly wash the cells with PBS. Block the cells with 5%NGS/0.3% Triton X-100/PBS for 30 min at room temperature. Incubate the cells with the primary antibodies for 1 h at room temperature (or overnight at 4 C). Wash the cells three times in PBS for 5 min at room temperature. Incubate the cells with the secondary antibodies for 1 h at room temperature. Wash the cells three times in PBS for 5 min at room temperature. Mount the slides with ProLong Gold antifade reagent with DAPI nucleic acid stain included. Evaluate the positive cells using a fluorescence microscope. Figs. 3.4 and 3.5 illustrate staining of differentiated mature pericytes under monoculture and endothelial co-culture conditions, respectively.

REFERENCES Ades, E. W., Candal, F. J., Swerlick, R. A., George, V. G., Summers, S., Bosse, D. C., and Lawley, T. J. (1992). HMEC-1: Establishment of an immortalized human microvascular endothelial cell line. J. Invest. Dermatol. 99, 683–690. Aghi, M., and Chiocca, E. A. (2005). Contribution of bone marrow-derived cells to blood vessels in ischemic tissues and tumors. Mol. Ther. 12, 994–1005. Aghi, M., Cohen, K. S., Klein, R. J., Scadden, D. T., and Chiocca, E. A. (2006). Tumor stromal–derived factor-1 recruits vascular progenitors to mitotic neovasculature, where microenvironment influences their differentiated phenotypes. Cancer Res. 66, 9054–9064. Ahn, G. O., and Brown, J. M. (2008). Matrix metalloproteinase-9 is required for tumor vasculogenesis but not for angiogenesis: Role of bone marrow–derived myelomonocytic cells. Cancer Cell 13, 193–205. Allt, G., and Lawrenson, J. G. (2001). Pericytes: Cell biology and pathology. Cells Tissues Organs 169, 1–11. Asahara, T., Murohara, T., Sullivan, A., Silver, M., van der Zee, R., Li, T., Witzenbichler, B., Schatteman, G., and Isner, J. M. (1997). Isolation of putative progenitor endothelial cells for angiogenesis. Science 275, 964–967. Bailey, A. S., Willenbring, H., Jiang, S., Anderson, D. A., Schroeder, D. A., Wong, M. H., Grompe, M., and Fleming, W. H. (2006). Myeloid lineage progenitors give rise to vascular endothelium. Proc. Natl. Acad. Sci. USA 103, 13156–13161.

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Bergers, G., Brekken, R., McMahon, J., Vu, T., Itoh, T., Tamakki, K., Tanzawa, K., Thorpe, P., Itohara, S., Werb, Z., and Hanahan, D. (2000). Matrix metalloproteinase-9 triggers the angiogenic switch during carcinogenesis. Nat. Cell Biol. 2, 737. Bergers, G., Hanahan, D., and Coussens, L. M. (1998). Angiogenesis and apoptosis are cellular parameters of neoplastic progression in transgenic mouse models of tumorigenesis. Dev. Biol. Int. J. Dev. Biol. 42, 995–1002. Bergers, G., Song, S., Meyer-Morse, N., Bergsland, E., and Hanahan, D. (2003). Benefits of targeting both pericytes and endothelial cells in the tumor vasculature with kinase inhibitors. J. Clin. Invest. 111, 1287–1295. Betsholtz, C., Lindblom, P., and Gerhardt, H. (2005). Role of pericytes in vascular morphogenesis. EXS 94, 115–125. Bunt, S. K., Sinha, P., Clements, V. K., Leips, J., and Ostrand-Rosenberg, S. (2006). Inflammation induces myeloid-derived suppressor cells that facilitate tumor progression. J. Immunol. 176, 284–290. Carmeliet, P. (2003). Angiogenesis in health and disease. Nat. Med. 9, 653–660. Ceradini, D. J., Kulkarni, A. R., Callaghan, M. J., Tepper, O. M., Bastidas, N., Kleinman, M. E., Capla, J. M., Galiano, R. D., Levine, J. P., and Gurtner, G. C. (2004). Progenitor cell trafficking is regulated by hypoxic gradients through HIF-1 induction of SDF-1. Nat. Med. 10, 858–864. Condeelis, J., and Pollard, J. W. (2006). Macrophages: Obligate partners for tumor cell migration, invasion, and metastasis. Cell 124, 263–266. Coussens, L. M., Tinkle, C. L., Hanahan, D., and Werb, Z. (2000). MMP-9 supplied by bone marrow-derived cells contributes to skin carcinogenesis. Cell 103, 481–490. De Falco, E., Porcelli, D., Torella, A. R., Straino, S., Iachininoto, M. G., Orlandi, A., Truffa, S., Biglioli, P., Napolitano, M., Capogrossi, M. C., and Pesce, M. (2004). SDF-1 involvement in endothelial phenotype and ischemia-induced recruitment of bone marrow progenitor cells. Blood 104, 3472–3482. De Palma, M., Venneri, M. A., Galli, R., Sergi, L. S., Politi, L. S., Sampaolesi, M., and Naldini, L. (2005). Tie2 identifies a hematopoietic lineage of proangiogenic monocytes required for tumor vessel formation and a mesenchymal population of pericyte progenitors. Cancer Cell 8, 211–226. Dome, B., Dobos, J., Tovari, J., Paku, S., Kovacs, G., Ostoros, G., and Timar, J. (2008). Circulating bone marrow–derived endothelial progenitor cells: Characterization, mobilization, and therapeutic considerations in malignant disease. Cytometry A 73, 186–193. Du, R., Lu, K. V., Petritsch, C., Liu, P., Ganss, R., Passegue, E., Song, H., Vandenberg, S., Johnson, R. S., Werb, Z., and Bergers, G. (2008). HIF1alpha induces the recruitment of bone marrow–derived vascular modulatory cells to regulate tumor angiogenesis and invasion. Cancer Cell 13, 206–220. Egeblad, M., and Werb, Z. (2002). New functions for the matrix metalloproteinases in cancer progression. Nat. Rev. Cancer 2, 161–174. Gerhardt, H., and Betsholtz, C. (2003). Endothelial–pericyte interactions in angiogenesis. Cell Tissue Res. 314, 15–23. Gerhardt, H., Golding, M., Fruttiger, M., Ruhrberg, C., Lundkvist, A., Abramsson, A., Jeltsch, M., Mitchell, C., Alitalo, K., Shima, D., and Betsholtz, C. (2003). VEGF guides angiogenic sprouting utilizing endothelial tip cell filopodia. J. Cell Biol. 161, 1163–1177. Grunewald, M., Avraham, I., Dor, Y., Bachar-Lustig, E., Itin, A., Jung, S., Chimenti, S., Landsman, L., Abramovitch, R., and Keshet, E. (2006). VEGF-induced adult neovascularization: recruitment, retention, and role of accessory cells. Cell 124, 175–189. Hanahan, D. (1985). Heritable formation of pancreatic ß-cell tumors in transgenic mice harboring recombinant insulin/simian virus 40 oncogenes. Nature 315, 115. Hattori, K., Heissig, B., and Rafii, S. (2003). The regulation of hematopoietic stem cell and progenitor mobilization by chemokine SDF-1. Leuk. Lymphoma 44, 575–582.


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Hattori, K., Heissig, B., Wu, Y., Dias, S., Tejada, R., Ferris, B., Hicklin, D. J., Zhu, Z., Bohlen, P., Witte, L., Hendrikx, J., Hackett, N. R., et al. (2002). Placental growth factor reconstitutes hematopoiesis by recruiting VEGFR1(þ) stem cells from bone-marrow microenvironment. Nat. Med. 8, 841–849. Heissig, B., Hattori, K., Dias, S., Friedrich, M., Ferris, B., Hackett, N. R., Crystal, R. G., Besmer, P., Lyden, D., Moore, M. A., Werb, Z., and Rafii, S. (2002). Recruitment of stem and progenitor cells from the bone marrow niche requires MMP-9 mediated release of kit-ligand. Cell 109, 625–637. Hellstrom, M., Kalen, M., Lindahl, P., Abramsson, A., and Betsholtz, C. (1999). Role of PDGF-B and PDGFR-beta in recruitment of vascular smooth muscle cells and pericytes during embryonic blood vessel formation in the mouse. Development 126, 3047–3055. Jin, D. K., Shido, K., Kopp, H. G., Petit, I., Shmelkov, S. V., Young, L. M., Hooper, A. T., Amano, H., Avecilla, S. T., Heissig, B., Hattori, K., Zhang, F., et al. (2006). Cytokinemediated deployment of SDF-1 induces revascularization through recruitment of CXCR4þ hemangiocytes. Nat. Med. 12, 557–567. Lamagna, C., and Bergers, G. (2006). The bone marrow constitutes a reservoir of pericyte progenitors. J. Leukoc. Biol. 80, 677–681. Li, B., Sharpe, E. E., Maupin, A. B., Teleron, A. A., Pyle, A. L., Carmeliet, P., and Young, P. P. (2006). VEGF and PlGF promote adult vasculogenesis by enhancing EPC recruitment and vessel formation at the site of tumor neovascularization. FASEB J. 20, 1495–1497. Luttun, A., Carmeliet, G., and Carmeliet, P. (2002). Vascular progenitors: from biology to treatment. Trends Cardiovasc. Med. 12, 88–96. Lyden, D., Hattori, K., Dias, S., Costa, C., Blaikie, P., Butros, L., Chadburn, A., Heissig, B., Marks, W., Witte, L., Wu, Y., Hicklin, D., et al. (2001). Impaired recruitment of bone-marrow–derived endothelial and hematopoietic precursor cells blocks tumor angiogenesis and growth. Nat. Med. 7, 1194–1201. Page-McCaw, A., Ewald, A. J., and Werb, Z. (2007). Matrix metalloproteinases and the regulation of tissue remodelling. Nat. Rev. Mol. Cell Biol. 8, 221–233. Petit, I., Jin, D., and Rafii, S. (2007). The SDF-1-CXCR4 signaling pathway: A molecular hub modulating neo-angiogenesis. Trends Immunol. 28, 299–307. Pollard, J. W. (2004). Tumour-educated macrophages promote tumour progression and metastasis. Nat. Rev. Cancer 4, 71–78. Rabbany, S. Y., Heissig, B., Hattori, K., and Rafii, S. (2003). Molecular pathways regulating mobilization of marrow-derived stem cells for tissue revascularization. Trends Mol. Med. 9, 109–117. Rafii, S., Avecilla, S., Shmelkov, S., Shido, K., Tejada, R., Moore, M. A., Heissig, B., and Hattori, K. (2003). Angiogenic factors reconstitute hematopoiesis by recruiting stem cells from bone marrow microenvironment. Ann. N. Y. Acad. Sci. 996, 49–60. Rafii, S., Meeus, S., Dias, S., Hattori, K., Heissig, B., Shmelkov, S., Rafii, D., and Lyden, D. (2002). Contribution of marrow-derived progenitors to vascular and cardiac regeneration. Dev. Biol.Semin. Cell Dev. Biol. 13, 61–67. Rajantie, I., Ilmonen, M., Alminaite, A., Ozerdem, U., Alitalo, K., and Salven, P. (2004). Adult bone marrow–derived cells recruited during angiogenesis comprise precursors for periendothelial vascular mural cells. Blood 104, 2084–2086. Rohde, E., Malischnik, C., Thaler, D., Maierhofer, T., Linkesch, W., Lanzer, G., Guelly, C., and Strunk, D. (2006). Blood monocytes mimic endothelial progenitor cells. Stem Cells 24, 357–367. Seandel, M., Butler, J., Lyden, D., and Rafii, S. (2008). A catalytic role for proangiogenic marrow-derived cells in tumor neovascularization. Cancer Cell 13, 181–183. Shaked, Y., Ciarrocchi, A., Franco, M., Lee, C. R., Man, S., Cheung, A. M., Hicklin, D. J., Chaplin, D., Foster, F. S., Benezra, R., and Kerbel, R. S. (2006). Therapy-induced acute recruitment of circulating endothelial progenitor cells to tumors. Science 313, 1785–1787.

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Song, S., Ewald, A. J., Stallcup, W., Werb, Z., and Bergers, G. (2005). PDGFRbetaþ perivascular progenitor cells in tumours regulate pericyte differentiation and vascular survival. Nat. Cell Biol. 7, 870–879. Yang, L., DeBusk, L. M., Fukuda, K., Fingleton, B., Green-Jarvis, B., Shyr, Y., Matrisian, L. M., Carbone, D. P., and Lin, P. C. (2004). Expansion of myeloid immune suppressor GrþCD11bþ cells in tumor-bearing host directly promotes tumor angiogenesis. Cancer Cell 6, 409–421.

C H A P T E R

F O U R

Screening Phage-Display Peptide Libraries for Vascular Targeted Peptides Martin Trepel,* Renata Pasqualini,† and Wadih Arap† Contents 1. Introduction 1.1. Vascular heterogeneity and its characterization by phage display 1.2. Targeting vascular addresses in humans 1.3. Vascular specialization within single organs 1.4. Applications of phage-derived peptides for targeted therapies 1.5. Applications of phage-derived peptides for diagnostics 1.6. Ligand-directed vectors for vascular targeted gene delivery and targeted molecular imaging 2. Methods 2.1. General protocols and requirements 2.2. Cell-free screening of phage libraries on isolated receptors 2.3. Screening the molecular diversity of cell surfaces in vitro: The BRASIL method 2.4. Screening the vascular diversity in living animals: In vivo phage display technology 3. Concluding Remarks and Perspective Acknowledgments References

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Abstract Depending on their physiologic location and functional state, vascular endothelial cells express surface receptors differentially. Recognition of this molecular diversity is essential for the development of targeted therapies. Random phage display peptide libraries can be selected in vitro on recombinant proteins or on intact cells. After systemic injection, selection can be performed in animals and * {

Department of Oncology and Hematology, University Medical Center Hamburg-Eppendorf, Hamburg, Germany The University of Texas M. D. Anderson Cancer Center, Department of Genitourinary Medical Oncology, Houston, Texas


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humans in vivo for the isolation of ligands for tissue-specific receptors. For the screening of libraries on intact cells or tissues, no a priori knowledge of the targeted receptor is needed, as the recovered peptide ligands can identify their corresponding receptors. Furthermore, the isolated peptides can be used to target therapeutic chemicals, biologicals, gene therapy vectors, or diagnostic compounds to specific tissues in vivo. Protocols for the screening of phage libraries in these three settings—on proteins, on cells in vitro and in the living animal—are described in this chapter.

1. Introduction 1.1. Vascular heterogeneity and its characterization by phage display The vasculature of each organ expresses a unique pattern of cell surface molecules that characterize the endothelial heterogeneity specific to that particular tissue. Similarly, tumor vasculature expresses unique receptors that distinguish it from normal, quiescent vasculature (Hajitou et al., 2006a; Sergeeva et al., 2006; Trepel et al., 2002). Such tissue-specific endothelial cell surface proteins could serve as receptors for tissue-specific homing of soluble circulating ligands, or for homing of cells such as leukocytes or possibly cancer stem cells for the formation of metastases. These specific endothelial receptors have been defined as functional vascular addresses or ‘‘vascular zip codes’’ (Folkman, 1999; Narasimhan, 2002; Rafii et al., 2003; Ruoslahti, 2004; Thorpe and Ran, 2002) and can be targeted systemically throughout the circulation. Ligand-directed targeting involves profiling of vascular addresses in organs under both normal and pathological conditions that include cancer, inflammation, or atherosclerosis. High-throughput methods for the identification of novel tissue-specific vascular markers have been developed by several groups, as described in this and other chapters within this volume. One of these techniques is the screening of random phage display peptide libraries in vitro and in vivo. Such libraries represent large collections of phage particles displaying peptides with up to 109 unique sequences (Smith and Scott, 1993). To identify and functionally validate tissue specific targets such as outlined above, it is critical to look beyond receptor expression levels and take ligand accessibility as a key determinant in pharmacodelivery systems into account (Ozawa et al., 2008). Selection of phage libraries in vivo as described in detail in Section 2.4 of this chapter does exactly that. It allows the recovery of displayed peptides homing preferentially to the target tissue due to their binding to tissue-specific vascular receptors. The selection is performed without a priori knowledge of the target receptor and is based not only on the level of expression, but also on the accessibility of the receptor to

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circulating ligands. The selection process is therefore unbiased and functionally controlled. The procedure involves intravenous injection of a random phage-displayed peptide library for a predetermined circulation time in which suitable peptides home to vascular receptors, followed by removal and washing of the target tissue, subsequent elution, and amplification of recovered peptide displaying phage which are then subject to further rounds of selection. The amino acid sequences conferring binding to vascular addresses in various tissues can be identified by gene sequencing. A potential natural ligand containing a similar sequence can often be identified by homology analysis in protein sequence databases, and its receptor can be deduced accordingly (Arap et al., 2002b; Giordano et al., 2001; Pasqualini et al., 2000; Rajotte and Ruoslahti, 1999). The method of in vivo phage display library screening is described in Section 2.4 of this chapter. In vivo phage display is used both for the identification of novel peptide ligands homing to specific vascular addresses as well as the characterization of receptor expression heterogeneity in both normal and pathologic blood vessels. A variety of peptides that bind to tissue-specific receptors and selectively home to tissues in mice have been described, including those recognizing brain, kidney, lung, skin, pancreas, intestine, uterus, adrenal gland, retina, lymph nodes, muscle, prostate, breast, placenta, and fat tissue (Arap et al., 2002a; Essler and Ruoslahti, 2002; Kolonin et al., 2002; Kolonin et al., 2004; Kolonin et al., 2006b; Pasqualini and Ruoslahti, 1996; Rajotte et al., 1998; Trepel et al., 2001). In addition, peptides and their receptors homing to angiogenic vasculature in tumors or wounds have been identified (Arap et al., 1998; Christian et al., 2003; Ja¨rvinen and Ruoslahti, 2007; Pasqualini et al., 2000; Porkka et al., 2002). The progression of tumors from a benign to metastatic state was profiled by phagederived peptides in a model of murine pancreatic cancer (Hoffman et al., 2003; Joyce et al., 2003). Vascular endothelial diversity for lymphatic vessels has also recently been described by the use of in vivo phage display (Laakkonen et al., 2002; Zhang et al., 2006). In many of these studies, isolation of the tissue-homing peptides led to the identification of the corresponding tissue-specific receptors via protein database analysis and/or biochemical strategies such as affinity chromatography, immunoprecipitation, and/or mass spectrometry. Tissue-specific receptors that have been identified include membrane dipeptidase expressed in the lung (Rajotte and Ruoslahti, 1999), interleukin 11 receptor in the prostate (Arap et al., 2002b), Fc-receptor in the placenta (Kolonin et al., 2002), prohibitin in white fat (Kolonin et al., 2004), EphA4 receptor in pancreas (Yao et al., 2005), aminopeptidase N (Pasqualini et al., 2000), aminopeptidase P (Essler and Ruoslahti, 2002), and nucleolin (Christian et al., 2003) in tumor angiogenic vasculature. As opposed to the in vivo phage display technology, peptides targeting vascular receptors can also be selected on known tissue-specific receptors,

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presented as recombinant proteins or receptors overexpressed on cultured cells, that can subsequently be validated in vivo as tissue homing ligands (Arap et al., 2004; Burg et al., 1999; Koivunen et al., 1999; Marchio et al., 2004; Pasqualini et al., 1997). These approaches are described in Sections 2.2 and 2.3 of this chapter.

1.2. Targeting vascular addresses in humans Tissue-homing ligands and their receptors initially identified in animal models have been useful for the validation of corresponding targets in human disease (Arap et al., 2002a; Carson-Walter et al., 2001; Pasqualini et al., 2000). However, many examples demonstrate the diversity of tissuespecific protein expression patterns and ligand-receptor accessibility in humans and mice (Bacich et al., 2001; Chang et al., 1999). Therefore, the systematic mapping of receptors in human blood vessels is mandatory for the development of targeted therapies in the clinic and for successful translational and clinical trials (St Croix et al., 2000). Consequently, in vivo selection of random phage display peptide libraries in human patients is a major advance in the identification of clinically relevant vascular targeting ligands and their receptors. Using a novel experimental approach, we selected a phage peptide library in a human subject and performed a high-throughput analysis of peptide motif distribution in bone marrow, prostate, fat, muscle, and skin (Arap et al., 2002b). We showed that the tissue distribution of circulating peptides in vivo in humans is nonrandom. For each of the isolated tissues, we found a tripeptide sequence motif mediating homing to the target after intravenous injection. Eleven tripeptides were enriched in a unique tissue, and an additional 14 were enriched in multiple organs. This result implies that some of the isolated peptides bind to tissue-specific endothelial cell surface receptors, whereas others bind to ubiquitous vascular markers (Arap et al., 2002b). A protein database search for sequence similarities between the recovered motifs and natural proteins revealed that the peptide GRRAGGS, recovered from the prostate, mimics interleukin 11 (IL-11); we subsequently showed that this IL-11-mimicking peptide indeed bound to the IL-11 receptor (IL-11Ra) (Arap et al., 2002b). Later, we validated IL-11Ra as a morphologic and functional marker during the progression of human prostate cancer in a large panel of patient samples (Zurita et al., 2004). Recently, tumor vasculature–targeting peptides have also been isolated from colon cancer specimens by an ex vivo approach (Maruta et al., 2007). We have recently refined the in vivo targeting technology to allow for synchronous combinatorial selection of ligands from multiple organs (as opposed to only one at a time) in mice (Kolonin et al., 2006b). This approach was subsequently adopted for use in human subjects (unpublished data). Systematic implementation of such screenings will broaden our


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knowledge of human vascular heterogeneity and will enable us to draft a functional human vascular receptor map for the development of liganddirected receptor–targeted therapies and diagnostics.

1.3. Vascular specialization within single organs Vascular heterogeneity has recently been refined at the level of functionally distinct regions within single organs. We have combined in vivo phage display with laser pressure catapult microdissection, used to isolate specific cells or compartments from sections of heterogeneous tissues (Eltoum et al., 2002). As a proof of principle, we have identified peptide ligands targeting vascular receptors to the islets within the mouse pancreas. A peptide homologous to ephrin A-type ligands was initially identified by Yao et al. (2005), and the corresponding receptor EphA4 was subsequently detected in pancreatic islets and shown to be enhanced in pancreatic islet tumors of the RIP-Tag2 transgenic mouse (Yao et al., 2005). It is highly likely that such specialization of the endothelium within functionally distinct tissue compartments also occurs in other organs such as the kidney, central nervous system, or the digestive tract.

1.4. Applications of phage-derived peptides for targeted therapies Peptides binding to cell type–specific receptors can be used in a variety of therapeutic and diagnostic applications, including the targeting of therapeutic agents and diagnostic compounds to certain tissues, and the targeting of gene therapy vectors to specific receptors. Targeting of therapeutic agents to certain tissues leads to increased therapeutic efficacy and fewer unintended side effects. In a pilot approach, we used the cytotoxic drug doxorubicin conjugated to tumor-homing peptides containing the peptide motif NGR or RGD-4C, respectively (Arap et al., 1998). This and subsequent confirmatory studies showed that the peptide-targeted cytostatic drug exhibited substantially stronger antitumor effects than the untargeted drug (Arap et al., 1998; Pastorino et al., 2006). Similar approaches were used in subsequent work targeting proapoptotic peptides, cytokines, or antisense oligonucleotides to tumors (Arap et al., 2004; Curnis et al., 2000, 2005; Ellerby et al., 1999; Henke et al., 2008; Zurita et al., 2004), the prostate (Arap et al., 2002a), or fat tissue (Kolonin et al., 2004). This targeting was followed by the selective destruction of the vasculature within the tumor and subsequent regression of the target tissue. Tissue-targeted peptides binding to receptor proteases can also be used as enzyme inhibitors in the absence of an additional conjugation of a drug moiety. Matrix metalloprotease-9 or aminopeptidase A are both increased in tumor vasculature and systemic administration of tumor-homing peptides

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that recognize these proteases inhibited tumor growth in vivo (Koivunen et al., 1999; Marchio et al., 2004). Phage-derived peptides have also been used for targeting noncytotoxic drugs such as antigens to lymph nodes (Trepel et al., 2001) or for transdermal protein delivery to the bloodstream (Chen et al., 2006).

1.5. Applications of phage-derived peptides for diagnostics Tissue-homing peptides have been used widely for targeting diagnostic compounds to certain tissues. Radiolabeled tumor-homing peptides selected from phage libraries were used for scintigraphic tumor imaging in vivo (Kuhnast et al., 2004; Sun et al., 2007). Vascular cell adhesionmolecule-1(VCAM-1)–targeted imaging agents, detectable by MRI and fluorescence imaging of tumors and atherosclerotic lesions in vivo, were produced from phage display-derived peptides and multimodal nanoparticles (Kelly et al., 2005). For ex vivo microscopic diagnosis, fluorescencelabeled tumor cell binding peptides identified by phage display have been utilized for the detection of bladder cancer cells in urine samples of patients with this disease. (Lee et al., 2007). For imaging of neoplastic tissue in vivo, we targeted streptavidinconjugated luminescent quantum dots to tumors in vivo by the use of a chimeric phage display system displaying tumor-homing peptides and a streptavidin-binding moiety (Chen et al., 2004). In another approach, we created spontaneously fabricating, biologically active molecular networks consisting of peptide-targeted phage with capsid-incorporated gold nanoparticles, for cell type–directed imaging (Souza et al., 2006). These targeted multifunctional features within a single entity may provide high potential for nanotechnology-based biomedical applications.

1.6. Ligand-directed vectors for vascular targeted gene delivery and targeted molecular imaging Unintended transduction of certain tissues, adverse immune reactions, and lack of efficient transduction of the cells of interest are the most important limitations of current gene therapy vectors (Trepel et al., 2000a). Therefore, addressing the biological safety and the poor efficacy of gene transfer pose significant challenges for gene therapy (Somia and Verma, 2000; Thomas et al., 2003). Targeting vectors to specific cellular receptors could solve some or all of these problems. Strategies to use peptides for viral vector targeting typically include (1) bispecific molecular conjugates consisting of antivector antibodies and peptide ligands directed toward the target receptor (Trepel et al., 2000b), or (2) insertion of specific peptide ligands into certain sites of the vector capsid (Girod et al., 1999; Grifman et al., 2001; Loiler et al., 2003; Mu¨ller et al., 2003; Nicklin et al., 2001; Reynolds et al., 1999; Shi and


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Bartlett, 2003; White et al., 2001, 2004; Work et al., 2004). Since the second strategy is not based on conjugates, it has numerous advantages, such as ease of handling, maintenance of the small size of the vector particle, better stability in vitro and in vivo, and no additional conjugate-related immunogenicity. Peptide ligands isolated by phage display libraries can be incorporated into eukaryotic vectors (Grifman et al., 2001; Loiler et al., 2003; Reynolds et al., 1999; Shi and Bartlett, 2003; White et al., 2004), but the success rate of this approach is variable. Our own experience has been that only a minority of selected peptide ligands (less than 30%) function equally well in targeted phage particles and in modified vector capsids, such as adenovirus or adenoassociated virus (AAV). One of the limitations accounting for this observation may be that the phage-derived peptides were selected only for cell or receptor binding but not for the subsequent post-targeting cell entry required for gene transfer. Moreover, the structural context is probably crucial. The binding property of a ligand peptide may change unpredictably when it is incorporated into a virus capsid protein; for example, it might be subjected to structural constraints not present in the phage capsid that was initially used for selection of the ligand from the random library. Taking these limitations into account, we and others have developed random peptide-display libraries based on the gene therapy vector capsid itself for AAVs (Mu¨ller et al., 2003; Perabo et al., 2003) and later, for retroviruses (Bupp and Roth, 2003; Hartl et al., 2005; Khare et al., 2003a,b). Peptide ligands binding specifically to a cell type within the context of the specific viral capsid protein can therefore be selected. With this technology, vectors were isolated that specifically and efficiently transduced the cell types on which they were selected (Michelfelder et al., 2007; Mu¨ller et al., 2003; Perabo et al., 2003; Waterkamp et al., 2006). By using the targeted phage particle itself for gene delivery, we have combined the advantages of phage-derived peptides for targeted gene delivery with that of peptides in the structural protein context on which they were selected. Phage have been used for transduction of eukaryotic cells (Larocca et al., 1999; Piersanti et al., 2004; Poul and Marks, 1999), but inefficient transduction and immunogenicity prevented broader application of phage vectors in vivo. We have therefore introduced a new hybrid vector containing genetic cis-elements from adeno-associated virus (AAV) and from a single-stranded M13 bacteriophage. This vector system was termed AAV phage (AAVP) (Hajitou et al., 2006b). An AAVP prototype was established displaying the RGD-4C peptide targeted to alpha V integrins, which are enhanced in tumor vessels. The vector mediated tumor-specific transduction after systemic administration in vivo. AAVP-mediated gene transfer was used both for suicide gene therapy and molecular-genetic imaging (Hajitou et al., 2006b) and can even be used as a ‘‘theranostic’’ (Ozawa et al., 2008) in a clinic-ready setting with positron emission tomography (PET) to predict tumor response to a systemically administered cytotoxic

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agent (Hajitou et al., 2008). This new class of targeted prokaryotic/eukaryotic viral hybrid vectors has a multitude of potential applications in biomedical research. A detailed methodology for the use of phage display technology for targeted gene delivery has been published recently (Hajitou et al., 2006b, 2007) and is therefore not described here.

2. Methods 2.1. General protocols and requirements 2.1.1. Materials F-pilus—positive Escherichia coli Theoretically, any F-pilus—positive bacteria could be used for phage amplification. We use E. coli K91kan bacteria (originally a gift from G. Smith, University of Missouri, Columbia, MO). Libraries Cloning and production of random phage display peptide libraries have been described in detail elsewhere (Smith and Scott, 1993). Several random phage display peptide libraries in different types of vectors are also commercially available. Libraries should comprise a diversity of 108 or more unique phage peptide sequences. We prefer peptide lengths of nine or fewer random residues, because their selection yields good affinity ligands; moreover, the potential diversity of longer library inserts is so high that libraries representing this diversity cannot be achieved under regular experimental conditions. Amplified libraries may not always be optimal for some of the selection protocols described here. We therefore recommend the use of primary, unamplified libraries only. Medium Supplement for Terrific Broth Mix 11.55 g KH2PO4, 105 g K2HPO4, and 500ml H20. Autoclave and use at a 1:10 dilution in terrific broth (TB). PEG/NaCl Mix 100 g PEG (particle size 8000), 110 g NaCl, and 450 ml H2O. Shake well and autoclave. Shake repeatedly while the solution is cooling down. General recommendation For working with phage, we recommend autoclaved or at least sterile-filtered solutions and aerosol-blocking filter pipette tips.


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2.1.2. Phage titering The bacteria used for recovery of the phage (we use K91kan) must be grown on the day of the panning. A streak from a K91kan agar plate is inoculated in 5 ml TB (supplemented with kanamycin at 200 mg/ml and 10% of medium supplement; see above) and grown at 37 C in a shaker. Approximately 2 to 4 h after initiation of the culture, the bacteria are typically in early stationary phase (OD600 of a 1:10 dilution in TB is 0.16 to 0.20). At this point, the shaker speed is slowed (to regenerate sheared pili), and bacteria must be used within 30 min. In a microtiter plate, five wells are prepared containing phage in serial 1:10 dilutions at amounts of 10–5 ml to 10–9 ml of the initial stock. To each well, 180 ml of the k91kan bacteria culture are added. After a 30-min incubation of phage with bacteria at room temperature, 100 ml of each of the solutions are plated on LB-tet plates labeled with the amount of phage being plated with the bacteria (10–5ml primary phage solution, 10–6 ml phage solution, and so on, up to 10–9 ml primary phage solution). Plates are grown overnight at 37 C in the incubator, and colonies are counted to determine the number of transducing units per microliter of phage solution (e.g., 20 colonies on the 10–6-plate correspond to 20 106 transducing phage units (TU) per microliter of primary phage solution, that is, 2 107 TU per microliter). Only plates with 20 to 600 colonies yield reliable results. 2.1.3. Phage single-clone sequencing A single colony of phage-transduced E. coli K91kan (or equivalent) is grown in LB containing 20 mg/ml tetracycline, and DNA plasmid minipreparation is performed following standard protocols. For sequencing of the random phage insert, we use the primer 50 -GCAAGCTGATAAACCGATA CAATT-30 . The recognition pattern for the insert in the phage genome is 50 -GCCGACGGGGCT–INSERT–GGGGCCGCTGGG-30 .

2.2. Cell-free screening of phage libraries on isolated receptors 2.2.1. Principle Screening of random phage display peptide libraries has been widely used to select peptides binding to purified and immobilized molecules in vitro (Sergeeva et al., 2006; Smith and Petrenko, 1997). The aim of such screening can be the characterization of protein–protein interactions, the mapping of their interaction domains, or the isolation of novel ligands for targeted drug delivery, such as targeting to vascular endothelial markers. The target protein is coated on a plastic surface in a well of a microtiter plate or equivalent matrix. After adding the library to the coated well, phage

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clones displaying a peptide specifically binding to the target protein remain bound in the well, while nonbinding phage are removed by washing. Bacteria are added to recover the phage from the target, because the phage transduce E. coli effectively via the bacterial F-pilus. Subsequently, phage are amplified within the bacteria, bacteria are eliminated by centrifugation, and the phage are purified. This procedure is repeated three to five times to enhance selectivity. Finally, DNA from the remaining phage is isolated and sequenced; hence, the amino acid sequences of the peptides binding to the target protein can be deduced. The phage-displayed inserts binding to the target protein ideally share common peptide motifs, typically consisting of three to five residues (Binder et al., 2006, 2007). 2.2.2. Preparation A well of a noncharged microtiter plate is coated overnight at 4 C with 1 to 20 mg of the target protein diluted in the working solution (usually 50 to 100 ml PBS). The amount of protein used depends on the coating efficiency. Two additional wells are coated with 3% BSA in PBS for preselection purposes. Coating can also be performed for 4 h at room temperature or for 2 h at 37 C. The K91kan bacteria used for recovery of the phage particles should be grown approximately 2 to 4 h prior to phage harvest (Section 2.1.2). 2.2.3. Preselection For many applications, a negative preselection of the phage library on irrelevant proteins is advantageous. The procedure may be omitted or reduced for the first selection round to increase phage yield (because yield is very important in the first round) and increased in stringency over the subsequent two rounds. For this purpose, one of the preselection wells is rinsed once with sterile PBS, after which 1010 TU of the library in 25 to 50 ml sterile PBS is added, and the well is incubated for 1 h at room temperature on a rocking platform. Subsequently, the second preselection well (if applicable) is washed with PBS, and the library is transferred from the first to this second preselection well. At the same time, the well coated with the target protein is washed with sterile PBS and filled with 200 ml sterile-filtered BSA (3% in PBS, to block nonspecific binding) for 1 h at room temperature. 2.2.4. Phage selection, recovery, and amplification The well coated with the target protein is washed once with sterile PBS, and the preselected library is transferred to the well containing the target protein. The phage library is incubated on the target for 2 h at room temperature on a rocking platform. The well is washed slowly four times with washing buffer (1% BSA in PBS plus 0.01% Tween 20) and subsequently once with PBS alone.


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The washing volume should slightly exceed the coating volume. To the empty well, 200 ml of the K91kan culture (see Section 2.1.2; after reaching early stationary growth phase, the bacteria should be used within 1 h; otherwise, they will lose infectability) are added and the well is incubated for 1 h at room temperature. After 1 h, the bacteria are transferred into a tube containing 10 ml of LB supplemented with tetracycline (0.4 mg/ml) and are incubated for 20 min at room temperature. Aliquots from this 10-ml mixture are plated on LB agar plates containing 40 mg/ml tetracycline. Usually 1, 10, and 100 ml are plated in duplicates. (For 1 ml and 10 ml platings the bacterial solution is diluted in medium to 100 ml to allow equal distribution on the plate.) The tetracycline concentration is adjusted to 20 mg/ml, and the bacterial solution is amplified overnight on a 37 C shaker. After overnight growth, the bacterial culture is centrifuged at 8000 rpm for 15 min, and the supernate is transferred to a clean tube. PEG/NaCl (see Section 2.1.1) is added at 1.5 ml per 10 ml of supernate. The tube must now be firmly closed and vigorously shaken by hand until frothing occurs. The sample is incubated for 1 h on ice and subsequently centrifuged at 8000 rpm for 20 min at 4 C. The resulting small white pellet contains the phage particles, and the supernate is discarded. The tube is spun again for 5 min at 8000 rpm to remove all PEG and to concentrate the pellet; remaining supernate is removed by vacuum or pipette. Three hundred ml TBS are added to the phage pellet. For re-suspension, we prefer placing the tube in the shaker for 10 min rather than resuspending the pellet with a pipette. The solution is transferred to a 1.5-ml tube and centrifuged at 14,000 rpm for 10 min to remove any bacterial proteins or debris. The supernate that contains the phage solution is transferred to a new tube. The titer of the recovered phage solution from Round 1 is determined following the protocol described in Section 2.1.2. 2.2.5. Additional rounds of selection The selection as described above is repeated with some modifications; for example, 1010 TU of the recovered phage is used for panning Round 2. Modifications in Round 2 versus Round 1 include the following: Coating: The amount of coated protein is decreased by 50%. In addition, two extra wells for preselection and one control well with a suitable control protein (e.g., IgG or BSA, used at the same concentration as the target protein, is used). Selection: After the second preselection step, the phage solution is distributed 1:1 into the target protein well and the control protein well, and is adjusted with 3% BSA to a total volume of 50 ml. Phage binding is restricted to 1 h at room temperature.

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Washing: After phage binding, the well is washed eight times with 1% BSA/ 0.01% Tween 20 in PBS. Plating: Platings have to be done both for the target well and for the control well. After the counting of colonies, one of the plates must be saved for sequencing, if enrichment of recovered phage from the target over phage recovered from the control is observed. The selection is repeated one more time (Round 3) with the following slight modifications: Coating: Only 20 to 30% of the initial amount (Round 1) of the target protein is coated. Selection: Phage binding is restricted to 30 min at room temperature Washing: After phage binding, the well is washed 10 to 12 times with 1% BSA/0.01% Tween 20 in PBS. The number of TU recovered from the target protein is compared to the number of TU recovered from the control. If enrichment in favor of the target protein is observed (usually in the range of 20- to 1000-fold), 30 single clones are grown for sequencing (following Section 2.1.3) and singleclone binding tests. Clones sharing sequence homology (i.e., a common sequence motif ) are evaluated in single-clone binding assays. 2.2.6. Single-clone binding assays on immobilized proteins in vitro The single-phage clones are titered according to Section 2.1.2 of this chapter. Evaluation of binding of single-phage clones to the target and control proteins is performed as that of phage binding during selection Round 3, but without preselection. Phage detection and quantification are accomplished by enzyme-linked immunosorbent assay (ELISA) following standard protocols with a primary polyclonal antiphage antibody. If specific binding is observed, the positively tested clones are sequenced. If binding to the target protein is not sufficiently enhanced over binding to the control protein, one to three more rounds of selection using the phage pool recovered from Round 3 are recommended, with increased selection stringency.

2.3. Screening the molecular diversity of cell surfaces in vitro: The BRASIL method 2.3.1. Principle A method that allows quick and effective selection of ligands from phage libraries on intact cells in suspension has been termed ‘‘biopanning and rapid analysis of selective interactive ligands’’ (BRASIL) (Giordano et al., 2001) and is described here. Since its introduction, BRASIL has been widely used for the selection of specific ligands binding to vascular, epithelial, neuronal,


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or tumor cells, respectively (Ardelt et al., 2003; Giordano et al., 2001; Hardy et al., 2007; Ja¨ger et al., 2007; Jia et al., 2007; Kolonin et al., 2006a; Schmidt et al., 2007). This strategy can be applied to systematic preclinical cell surface proteome mapping of human cancer cells and vascular endothelial cells. For selection on primary cells or cell lines in vitro, the library is incubated with a cell suspension rather than a recombinant protein, followed by removal of unbound phage particles. Instead of repetitive washing of the target cells, the separation of cell-bound versus unbound phage can be achieved by a single-step differential centrifugation of the cells with bound phage through an organic phase (Giordano et al., 2001). 2.3.2. Preparation and preselection For most applications, a ligand must be selected that binds via a type-specific receptor to a single type (or class) of cell. Therefore, the library is negatively selected on cells distinct from the cell type of interest (‘‘control cells’’). These control cells are grown under regular conditions, released with trypsin and washed in 10 ml RPMI with 1% BSA. A volume of 107 cells are subsequently pelleted by centrifugation at 800g, after which the supernate is discarded and the pellet is resuspended in 300 ml RPMI/BSA. Between 1 1010 and 1 1011 TU of random phage display peptide library (depending on the titer and diversity of the library) are added. This solution is transferred to a 1.5-ml tube, mixed, and incubated for 1 h on a rotating platform at room temperature to adsorb phage that bind to control cells. After 1 h, the suspension is centrifuged at 4000 rpm for 5 min. The supernate is saved, and the pellet is discarded. 2.3.3. Phage selection The target cell population should be viable (fresh primary tissue or tissue culture cells) and the cells must be in suspension. A volume equivalent to slightly more than 107 cells is centrifuged at 800g for 5 min. After the negative selection of the library, the supernate in the tube containing the pelleted target cells is removed, and the pellet is gently resuspended in the supernate derived from the preselection mix (see above). After transfer of the sample to a new 1.5-ml tube, 200 ml of RPMI/1% BSA is added, and the sample tube is put on ice for 5 min. Subsequently, the library is incubated with the target cells at 4 C on a rocking or turning platform for 4 to 10 h. 2.3.4. Differential centrifugation and removal of unbound phage During the incubation of the phage on the target cells, the organic phase for the differential centrifugation step is prepared by mixing dibutylphthalate and cyclohexane at a 9:1 v:v ratio (BRASIL oil). Usually, a stock solution of several ml is made that lasts for a few days. Larger stocks are not recommended, as

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repetitive opening of the stock tubes leads to evaporation of cyclohexane with consequent increase in the density of the oil). Two hundred microliters of fresh BRASIL oil are added to each of two 400-ml tubes. After 4 to 10 h of incubation (see above), the cell-phage solution is carefully added on top of the oil in the tube (150 ml into each tube). The organic phase and the cell/phage-containing aqueous phase must not be mixed. The tubes are centrifuged for 10 min at 10,000g and 4 C. The resulting cell pellet at the bottom of the tube (underneath the organic phase) contains the cell-bound phage, while the unbound phage remains in the aqueous supernate on top of the organic phase. Following centrifugation, the tubes are frozen for 10 min in dry ice or for 20 min at –80 C. 2.3.5. Phage recovery and amplification The tips of the tubes containing the pellet are cut off with single-use disposable scalpels. To avoid contamination, scalpels and gloves are discarded after the processing of target cell pellets and prior to that of control cell pellets (Rounds 2 and 3 only; see below). Excess oil from the tip is removed, and each tip is transferred into a single empty 2-ml tube. The pellets are resuspended in 100 ml PBS with the plastic tip in the tube. To each of the tubes, 1 ml of K91kan culture in early stationary phase is added (see Section 2.1.2). The solution is mixed gently and incubated at 37 C. After 30 min, bacteria from all tubes containing material from one cell population (target cells only in the first round, target or control cells, respectively, in subsequent rounds) are transferred to a 50-ml tube, and 20 ml prewarmed NZY medium containing tetracycline (0.2 mg/ml) are added. After 30 min at 37 C, 10 ml and 100 ml of this solution are plated on LB agar plates containing 40 mg/ml tetracycline. The tetracycline concentration in the rest of the liquid culture is adjusted to 20 mg/ml, and the culture is grown in two 15-ml tubes overnight (16 h maximum) at 37 C in the shaker. After overnight growth, the bacterial culture is centrifuged and the phage pellet is recovered as described in Section 2.2.4. The titer of the recovered phage solution from Round 1 is determined according to the protocol described in Section 2.1.2. 2.3.6. Additional rounds of selection The selection as described above is repeated with some modifications, with 2 1010 TU of the recovered phage for panning Round 2. Cells: The cell quantity can be decreased to 106. Negative selection: The negative preselection is repeated once on another vial of control cells (i.e., two preselection steps).


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Selection: After the second preselection step, the phage solution is distributed 1:1 to the target cells and the control cells, and solutions are adjusted with 3% BSA in RPMI to a total volume of 300 ml per tube. Phage binding is restricted to 5 min at room temperature. (If the BRASIL oil that is needed for further processing of the sample is not ready at this time, the sample is put on ice after 5 min to avoid internalization of the bound phage into the cells.) Plating: Platings are now complete for the target cells and the control cells. After the counting of colonies, one of the target cell–derived plates must be saved for sequencing, if enrichment of phage recovered from the target over phage recovered from the control is observed. The selection is repeated one more time (Round 3) with 2 109 TU of phage recovered from Round 2, again with slight modifications: Cells: 5 105 target and control cells are used. Negative selection: The negative preselection on control cells can be increased to four preselection steps, especially if there is no enrichment of phage on target cells after Round 2. In both Rounds 2 and 3, the number of TU recovered from the target cells is compared to the number of TU recovered from the control. If enrichment in favor of the target cells is observed (usually in the range of 5- to 100-fold), 30 single clones are grown for sequencing (see Section 2.1.3) and single-clone binding tests. Clones with a common sequence motif are evaluated in single-clone binding assays. 2.3.7. Single-clone binding assays on cells in vitro Single-phage clones are titered as described in Section 2.1.2. Binding of single-phage clones to target and control cells is performed as described for phage binding during selection in Round 3, but without preselection. Phage quantification from each cell population is accomplished by bacterial recovery and colony counting, as described for the selection procedure. If binding to the target cells is not sufficiently enhanced over binding to the control protein, one to three more rounds of selection using the phage pool recovered from Round 3 may be considered (performed as for Round 3).

2.4. Screening the vascular diversity in living animals: In vivo phage display technology 2.4.1. Principle Vascular heterogeneity can be characterized by ligands homing to tissuespecific endothelial cell receptors or ‘‘vascular addresses’’ (see Section 1.1). Our group has developed the in vivo phage-display technology, a strategy that has been widely used to identify tissue-specific ligand-receptor

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interactions in living animals and humans. Different peptide motifs binding tissue-specific receptors have been recovered from several tissues (Arap et al., 1998, 2002a,b; Christian et al., 2003; Essler and Ruoslahti, 2002; Ja¨rvinen and Ruoslahti, 2007; Kolonin et al., 2002, 2004, 2006b; Laakkonen et al., 2002; Pasqualini et al., 2000; Pasqualini and Ruoslahti, 1996; Porkka et al., 2002; Rajotte et al., 1998; Trepel et al., 2001). In principle, random phage-display peptide libraries are injected intravenously, and the target tissue is surgically collected after a short circulation time. Phage clones displaying ligands that mediated their homing to the collected tissue can be rescued and amplified by bacterial infection. The preselected, amplified library is re-injected for further enrichment of clones displaying peptides with optimal homing capacity. After three or four rounds of selection, recovered phage clones are sequenced to identify the DNA corresponding to the inserts displayed, as described for phage applications in vitro (see above). We have also established a strategy for simultaneous screening of multiple organs with phage display libraries in the mouse (Kolonin et al., 2006b), according to similar technical principles as outlined in this protocol. Therefore, we confine the description of the method to the screening of one organ at a time. 2.4.2. Animals The following protocol is described for a murine model. However, it can be adapted to other species with modifications based on the amount of phage injected and the strategy for phage recovery. We use primarily 2-month-old Balb/c mice. Nude mice are advantageous for avoidance of fur-related bacterial cross-contamination. The influence of different mouse strains on phage selection has not been investigated systematically, but we do not consider this to be a major concern since most isolated ligand receptors seem to be found valid across various strains. 2.4.3. Selection in vivo The phage library is diluted in DMEM tissue culture medium or PBS (1010 TU to a total volume of no more than 300 ml) and injected intravenously into the tail vein. The animal is kept alive for 5 min while the library circulates. 2.4.4. Tissue isolation, phage recovery, and phage amplification Transcardial perfusion of the animal following standard protocols prior to the surgical collection of the target tissue can decrease nonspecific phage recovery. For certain organs (such as the kidney), however, perfusion increases phage trapping and therefore recovery of nonspecific phage. Furthermore, perfusion is not always advantageous for the first round of selection because excessive stringency may worsen the outcome by


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eliminating some binding phage that are present in small quantities. If perfusion is not performed, exsanguination of the animal is an option to decrease background phage recovery from the blood. If perfusion is done, we commonly use 5 ml of DMEM at room temperature injected into the left ventricle. The organs of interest (target organ plus at least one control organ such as the lung or the brain) are surgically removed and put on ice immediately. The organs are weighed, homogenized with a glass tissue grinder or equivalent, and washed once in ice-cold DMEM containing a protease inhibitor cocktail (DMEM-PI). One milliliter of DMEM-PI is added to the tissue homogenate in a 2-ml tube, centrifuged at 4 C at 3000 rpm for 4 min, and decanted. After the washing step, the sample is kept on ice until the addition of bacteria (see below). The K91kan bacteria used for recovery of the phage are grown as described in Section 2.1.2. Next, 1500 ml of a K91kan culture in early stationary phase at 37 C are added to the phage-cell pellet, which at this point is resuspended gently but thoroughly. The mixture is incubated for 30 min at 37 C by swirling or inverting the tube at 10-min intervals. The entire contents of the tube are transferred to a 500-ml Erlenmeyer flask, and 100 ml prewarmed NZY medium containing tetracycline at 0.2 mg/ml are added. After a 30-min incubation at 37 C, 10 ml and 100 ml of the culture are plated on LB agar plates containing 40 mg/ml tetracycline to quantify the phage recovered from each tissue. The tetracycline concentration in the rest of the culture is adjusted to 20 mg/ml, followed by overnight growth in an Erlenmeyer flask (approximately 12 h, but no more than 16 h) at 37 C on the shaker. On the next day, the bacterial culture is centrifuged, and the phage pellet is recovered as described in Section 2.2.4. The titer of the recovered phage solution from Round 1 is determined according to the protocol described in Section 2.1.2. 2.4.5. Additional rounds of selection The selection as described above is repeated with some modifications, with 5 109 TU of the recovered phage for panning Round 2. Washing after selection: The homogenized tissue is washed 3 times (instead of once as in Round 1) in ice-cold DMEM-PI. One ml DMEM-PI is added to the tissue homogenate in a 2-ml tube and the contents are centrifuged at 4 C and 3000 rpm for 4 min. The supernate is removed (a small amount is retained on top of the cells). The tube is vortexed, after which 1 ml DMEM-PI is added and the contents centrifuged. The process is repeated for a total of three washes. Plating: If not already done in the first round, platings for the target tissue and for the control tissue(s) are now performed to evaluate numeric phage enrichment in the tissue of interest. After counting of the colonies,

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one of the target tissue–derived plates must be saved for sequencing, if enrichment of phage recovered from the target over phage recovered from the control is observed. The selection is repeated one more time (Round 3) with 1 109 TU of phage recovered from Round 2, and an increased washing stringency of four to five steps. In both Rounds 2 and 3, the number of TU recovered from the target tissue is compared to the number of TU recovered from the control tissues. If enrichment in favor of the target cells is observed (usually in the range of 3- to 50-fold), at least 30 single clones are grown for sequencing (Section 2.1.3) and single-clone binding tests. Clones sharing sequence homology (i.e., a common sequence motif ) are evaluated in single-clone binding assays. 2.4.6. Single-clone binding assays in vivo Single clones are titered as described in Section 2.1.2. Homing of singlephage clones to target and control tissues is evaluated as described for phage binding during selection Round 3. Quantification of phage from each cell population is performed by bacterial recovery and colony counting, as described for the selection procedure. If specific homing is not observed to a satisfactory degree, one to three more rounds of selection using the phage pool recovered from Round 3 may be considered (performed as described for Round 3). 2.4.7. Synchronous serial in vivo selection To circumvent the need for multiple-round screenings for peptides homing to each tissue of interest, we have recently introduced a simultaneous screening procedure for synchronous identification of organ-homing peptides in multiple tissues (Kolonin et al., 2006b). This method is performed basically as described here except that multiple organs are removed as target tissues in each round, and the phage recovered from these organs are pooled and reinjected for successive rounds of selection. The statistical algorithms for analysis of peptide sequences isolated in serial synchronous selections use Bayesian mixture models for complex high-dimensional count data (Kolonin et al., 2006b).

3. Concluding Remarks and Perspective Recognition of the diversity of cell surface–receptor expression as a function of specific tissues and states of cell activation is essential for the development of targeted therapies and diagnostics. Random phage-display peptide library biopanning is an invaluable tool for the production of a detailed receptor map of the vasculature. The systematic screening of


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libraries at three levels—recombinant target proteins, intact cells in certain functional states in vitro, and certain organs and diseased tissues in vivo, will lead to a comprehensive map of vascular ligand–receptor interactions and will direct the development of targeted drugs.

ACKNOWLEDGMENTS This work was supported by grants from the Deutsche Forschungsgemeinschaft Tr 448/5-3, the Wilhelm Sander-Stiftung 2005.114.1, and The Deutsche Jose Carreras Leuka¨mieStiftung DJCLS R 07/29f (MT), as well as the National Institutes of Health, Department of Defense, the Prostate Cancer Foundation, and the Gillson-Longenbaugh Foundation (RP and WA).

REFERENCES Arap, M. A., Lahdenranta, J., Mintz, P. J., Hajitou, A., Sarkis, A. S., Arap, W., and Pasqualini, R. (2004). Cell surface expression of the stress response chaperone GRP78 enables tumor targeting by circulating ligands. Cancer Cell 6, 275–284. Arap, W., Haedicke, W., Bernasconi, M., Kain, R., Rajotte, D., Krajewski, S., Ellerby, H. M., Bredesen, D. E., Pasqualini, R., and Ruoslahti, E. (2002a). Targeting the prostate for destruction through a vascular address. Proc. Natl. Acad. Sci. USA 99, 1527–1531. Arap, W., Kolonin, M. G., Trepel, M., Lahdenranta, J., Cardo-Vila, M., Giordano, R. J., Mintz, P. J., Ardelt, P. U., Yao, V. J., Vidal, C. I., Chen, L., Flamm, A., et al. (2002b). Steps toward mapping the human vasculature by phage display. Nat. Med. 8, 121–127. Arap, W., Pasqualini, R., and Ruoslahti, E. (1998). Cancer treatment by targeted drug delivery to tumor vasculature in a mouse model. Science 279, 377–380. Ardelt, P. U., Wood, C. G., Chen, L., Mintz, P. J., Moya, C., Arap, M. A., Wright, K. C., Pasqualini, R., and Arap, W. (2003). Targeting urothelium: Ex vivo assay standardization and selection of internalizing ligands. J. Urol. 169, 1535–1540. Bacich, D. J., Pinto, J. T., Tong, W. P., and Heston, W. D. W. (2001). Cloning, expression, genomic localization, and enzymatic activities of the mouse homolog of prostate-specific membrane antigen/NAALADase/folate hydrolase. Mamm. Genome 12, 117–123. Binder, M., Otto, F., Mertelsmann, R., Veelken, H., and Trepel, M. (2006). The epitope recognized by rituximab. Blood 108, 1975–1978. Binder, M., Vo¨gtle, F. N., Michelfelder, S., Mu¨ller, F., Illerhaus, G., Sundararajan, S., Mertelsmann, R., and Trepel, M. (2007). Identification of their epitope reveals the structural basis for the mechanism of action of the immunosuppressive antibodies basiliximab and daclizumab. Cancer Res. 67, 3518–3523. Bupp, K., and Roth, M. J. (2003). Targeting a retroviral vector in the absence of a known cell-targeting ligand. Hum. Gene Ther. 14, 1557–1564. Burg, M. A., Pasqualini, R., Arap, W., Ruoslahti, E., and Stallcup, W. B. (1999). NG2 proteoglycan-binding peptides target tumor neovasculature. Cancer Res. 59, 2869–2874. Carson-Walter, E. B., Watkins, D. N., Nanda, A., Vogelstein, B., Kinzler, K. W., and St Croix, B. (2001). Cell surface tumor endothelial markers are conserved in mice and humans. Cancer Res. 61, 6649–6655.

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Chang, S. S., Reuter, V. E., Heston, W. D., Bander, N. H., Grauer, L. S., and Gaudin, P. B. (1999). Five different anti-prostate-specific membrane antigen (PSMA) antibodies confirm PSMA expression in tumor-associated neovasculature. Cancer Res. 59, 3192–3198. Chen, L., Zurita, A. J., Ardelt, P. U., Giordano, R. J., Arap, W., and Pasqualini, R. (2004). Design and validation of a bifunctional ligand display system for receptor targeting. Chem. Biol. 11, 1081–1091. Chen, Y., Shen, Y., Guo, X., Zhang, C., Yang, W., Ma, M., Liu, S., Zhang, M., and Wen, L. P. (2006). Transdermal protein delivery by a coadministered peptide identified via phage display. Nat. Biotechnol. 24, 455–460. Christian, S., Pilch, J., Akerman, M. E., Porkka, K., Laakkonen, P., and Ruoslahti, E. (2003). Nucleolin expressed at the cell surface is a marker of endothelial cells in angiogenic blood vessels. J. Cell Biol. 163, 871–878. Curnis, F., Gasparri, A., Sacchi, A., Cattaneo, A., Magni, F., and Corti, A. (2005). Targeted delivery of IFNgamma to tumor vessels uncouples antitumor from counterregulatory mechanisms. Cancer Res. 65, 2906–2913. Curnis, F., Sacchi, A., Borgna, L., Magni, F., Gasparri, A., and Corti, A. (2000). Enhancement of tumor necrosis factor alpha antitumor immunotherapeutic properties by targeted delivery to aminopeptidase N (CD13). Nat. Biotechnol. 18, 1185–1190. Ellerby, H. M., Arap, W., Ellerby, L. M., Kain, R., Andrusiak, R., Rio, G. D., Krajewski, S., Lombardo, C. R., Rao, R., Ruoslahti, E., Bredesen, D. E., and Pasqualini, R. (1999). Anti-cancer activity of targeted pro-apoptotic peptides. Nat. Med. 5, 1032–1038. Eltoum, I. A., Siegal, G. P., and Frost, A. R. (2002). Microdissection of histologic sections: Past, present, and future. Adv. Anat. Pathol. 9, 316–322. Essler, M., and Ruoslahti, E. (2002). Molecular specialization of breast vasculature: A breasthoming phage-displayed peptide binds to aminopeptidase P in breast vasculature. Proc. Natl. Acad. Sci. USA 99, 2252–2257. Folkman, J. (1999). Angiogenic zip code. Nat. Biotechnol. 17, 749. Giordano, R. J., Cardo-Vila, M., Lahdenranta, J., Pasqualini, R., and Arap, W. (2001). Biopanning and rapid analysis of selective interactive ligands. Nat. Med. 7, 1249–1253. Girod, A., Ried, M., Wobus, C., Lahm, H., Leike, K., Kleinschmidt, J., Deleage, G., and Hallek, M. (1999). Genetic capsid modifications allow efficient re-targeting of adenoassociated virus type 2. Nat. Med. 5, 1052–1056. Grifman, M., Trepel, M., Speece, P., Gilbert, L. B., Arap, W., Pasqualini, R., and Weitzman, M. D. (2001). Incorporation of tumor-targeting peptides into recombinant adeno-associated virus capsids. Mol. Ther. 3, 964–975. Hajitou, A., Lev, D. C., Hannay, J. A., Korchin, B., Staquicini, F. I., Soghomonyan, S., Alauddin, M. M., Benjamin, R. S., Pollock, R. E., Gelovani, J. G., Pasqualini, R., and Arap, W. (2008). A preclinical model for predicting drug response in soft-tissue sarcoma with targeted AAVP molecular imaging. Proc. Natl. Acad. Sci. USA 105, 4471–4476. Hajitou, A., Pasqualini, R., and Arap, W. (2006a). Vascular targeting: Recent advances and therapeutic perspectives. Trends Cardiovasc. Med. 16, 80–88. Hajitou, A., Rangel, R., Trepel, M., Soghomonyan, S., Gelovani, J. G., Alauddin, M. M., Pasqualini, R., and Arap, W. (2007). Design and construction of targeted AAVP vectors for mammalian cell transduction. Nat. Protoc. 2, 523–531. Hajitou, A., Trepel, M., Lilley, C. E., Soghomonyan, S., Alauddin, M. M., Marini, F. C., 3rd, Restel, B. H., Ozawa, M. G., Moya, C. A., Rangel, R., Sun, Y., Zaoui, K., et al. (2006b). A hybrid vector for ligand-directed tumor targeting and molecular imaging. Cell 125, 385–398. Hardy, B., Raiter, A., Weiss, C., Kaplan, B., Tenenbaum, A., and Battler, A. (2007). Angiogenesis induced by novel peptides selected from a phage display library by screening human vascular endothelial cells under different physiological conditions. Peptides 28, 691–701.


103

Hartl, I., Schneider, R. M., Sun, Y., Medvedovska, J., Chadwick, M. P., Russell, S. J., Cichutek, K., and Buchholz, C. J. (2005). Library-based selection of retroviruses selectively spreading through matrix metalloprotease-positive cells. Gene Ther. 12, 918–926. Henke, E., Perk, J., Vider, J., de Candia, P., Chin, Y., Solit, D. B., Ponomarev, V., Cartegni, L., Manova, K., Rosen, N., and Benezra, R. (2008). Peptide-conjugated antisense oligonucleotides for targeted inhibition of a transcriptional regulator in vivo. Nat. Biotechnol. 26, 91–100. Hoffman, J. A., Giraudo, E., Singh, M., Zhang, L., Inoue, M., Porkka, K., Hanahan, D., and Ruoslahti, E. (2003). Progressive vascular changes in a transgenic mouse model of squamous cell carcinoma. Cancer Cell 4, 383–391. Ja¨ger, S., Jahnke, A., Wilmes, T., Adebahr, S., Vogtle, F. N., Delima-Hahn, E., Pfeifer, D., Berg, T., Lubbert, M., and Trepel, M. (2007). Leukemia-targeting ligands isolated from phage-display peptide libraries. Leukemia 21, 411–420. Ja¨rvinen, T. A., and Ruoslahti, E. (2007). Molecular changes in the vasculature of injured tissues. Am. J. Pathol.Am. J. Pathol. 171, 702–711. Jia, W. D., Sun, H. C., Zhang, J. B., Xu, Y., Qian, Y. B., Pang, J. Z., Wang, L., Qin, L. X., Liu, Y. K., and Tang, Z. Y. (2007). A novel peptide that selectively binds highly metastatic hepatocellular carcinoma cell surface is related to invasion and metastasis. Cancer Lett. 247, 234–242. Joyce, J. A., Laakkonen, P., Bernasconi, M., Bergers, G., Ruoslahti, E., and Hanahan, D. (2003). Stage-specific vascular markers revealed by phage display in a mouse model of pancreatic islet tumorigenesis. Cancer Cell 4, 393–403. Kelly, K. A., Allport, J. R., Tsourkas, A., Shinde-Patil, V. R., Josephson, L., and Weissleder, R. (2005). Detection of vascular adhesion molecule-1 expression using a novel multimodal nanoparticle. Circ. Res.Circ. Res. 96, 327–336. Khare, P. D., Rosales, A. G., Bailey, K. R., Russell, S. J., and Federspiel, M. J. (2003a). Epitope selection from an uncensored peptide library displayed on avian leukosis virus. Virology 315, 313–321. Khare, P. D., Russell, S. J., and Federspiel, M. J. (2003b). Avian leukosis virus is a versatile eukaryotic platform for polypeptide display. Virology 315, 303–312. Koivunen, E., Arap, W., Valtanen, H., Rainisalo, A., Medina, O. P., Heikkila, P., Kantor, C., Gahmberg, C. G., Salo, T., Konttinen, Y. T., Sorsa, T., Ruoslahti, E., et al. (1999). Tumor targeting with a selective gelatinase inhibitor. Nat. Biotechnol. 17, 768–774. Kolonin, M. G., Bover, L., Sun, J., Zurita, A. J., Do, K. A., Lahdenranta, J., Cardo-Vila, M., Giordano, R. J., Jaalouk, D. E., Ozawa, M. G., Moya, C. A., Souza, G. R., et al. (2006a). Ligand-directed surface profiling of human cancer cells with combinatorial peptide libraries. Cancer Res. 66, 34–40. Kolonin, M. G., Pasqualini, R., and Arap, W. (2002). Teratogenicity induced by targeting a placental immunoglobulin transporter. Proc. Natl. Acad. Sci. USA 99, 13055–13060. Kolonin, M. G., Saha, P. K., Chan, L., Pasqualini, R., and Arap, W. (2004). Reversal of obesity by targeted ablation of adipose tissue. Nat. Med. 10, 625–632. Kolonin, M. G., Sun, J., Do, K. A., Vidal, C. I., Ji, Y., Baggerly, K. A., Pasqualini, R., and Arap, W. (2006b). Synchronous selection of homing peptides for multiple tissues by in vivo phage display. FASEB J. 20, 979–981. Kuhnast, B., Bodenstein, C., Haubner, R., Wester, H. J., Senekowitsch-Schmidtke, R., Schwaiger, M., and Weber, W. A. (2004). Targeting of gelatinase activity with a radiolabeled cyclic HWGF peptide. Nucl. Med. Biol. 31, 337–344. Laakkonen, P., Porkka, K., Hoffman, J. A., and Ruoslahti, E. (2002). A tumor-homing peptide with a targeting specificity related to lymphatic vessels. Nat. Med. 8, 751–755. Larocca, D., Kassner, P. D., Witte, A., Ladner, R. C., Pierce, G. F., and Baird, A. (1999). Gene transfer to mammalian cells using genetically targeted filamentous bacteriophage. FASEB J. 13, 727–734.

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Lee, S. M., Lee, E. J., Hong, H. Y., Kwon, M. K., Kwon, T. H., Choi, J. Y., Park, R. W., Kwon, T. G., Yoo, E. S., Yoon, G. S., Kim, I. S., Ruoslahti, E., et al. (2007). Targeting bladder tumor cells in vivo and in the urine with a peptide identified by phage display. Mol. Cancer Res. 5, 11–19. Loiler, S. A., Conlon, T. J., Song, S., Tang, Q., Warrington, K. H., Agarwal, A., Kapturczak, M., Li, C., Ricordi, C., Atkinson, M. A., Muzyczka, N., and Flotte, T. R. (2003). Targeting recombinant adeno-associated virus vectors to enhance gene transfer to pancreatic islets and liver. Gene Ther. 10, 1551–1558. Marchio, S., Lahdenranta, J., Schlingemann, R. O., Valdembri, D., Wesseling, P., Arap, M. A., Hajitou, A., Ozawa, M. G., Trepel, M., Giordano, R. J., Nanus, D. M., Dijkman, H., et al. (2004). Aminopeptidase A is a functional target in angiogenic blood vessels. Cancer Cell 5, 151–162. Maruta, F., Akita, N., Nakayama, J., Miyagawa, S., Ismail, T., Rowlands, D. C., Kerr, D. J., Fisher, K. D., Seymour, L. W., and Parker, A. L. (2007). Bacteriophage biopanning in human tumour biopsies to identify cancer-specific targeting ligands. J. Drug Target. 15, 311–319. Michelfelder, S., Lee, M. K., deLima-Hahn, E., Wilmes, T., Kaul, F., Muller, O., Kleinschmidt, J. A., and Trepel, M. (2007). Vectors selected from adeno-associated viral display peptide libraries for leukemia cell-targeted cytotoxic gene therapy. Exp. Hematol. 35, 1766–1776. Mu¨ller, O. J., Kaul, F., Weitzman, M. D., Pasqualini, R., Arap, W., Kleinschmidt, J. A., and Trepel, M. (2003). Random peptide libraries displayed on adeno-associated virus to select for targeted gene therapy vectors. Nat. Biotechnol. 21, 1040–1046. Narasimhan, K. (2002). Zip codes: deciphering vascular addresses. Nat. Med. 8, 116. Nicklin, S. A., Von Seggern, D. J., Work, L. M., Pek, D. C., Dominiczak, A. F., Nemerow, G. R., and Baker, A. H. (2001). Ablating adenovirus type 5 fiber–CAR binding and HI loop insertion of the SIGYPLP peptide generate an endothelial cell– selective adenovirus. Mol. Ther. 4, 534–542. Ozawa, M. G., Zurita, A. J., Dias-Neto, E., Nunes, D. N., Sidman, R. L., Gelovani, J. G., Arap, W., and Pasqualini, R. (2008). Beyond receptor expression levels: The relevance of target accessibility in ligand-directed pharmacodelivery systems. Trends Cardiovasc. Med. 18, 126–132. Pasqualini, R., Koivunen, E., Kain, R., Lahdenranta, J., Sakamoto, M., Stryhn, A., Ashmun, R. A., Shapiro, L. H., Arap, W., and Ruoslahti, E. (2000). Aminopeptidase N is a receptor for tumor-homing peptides and a target for inhibiting angiogenesis. Cancer Res. 60, 722–727. Pasqualini, R., Koivunen, E., and Ruoslahti, E. (1997). Alpha v integrins as receptors for tumor targeting by circulating ligands. Nat. Biotechnol. 15, 542–546. Pasqualini, R., and Ruoslahti, E. (1996). Organ targeting in vivo using phage display peptide libraries. Nature 380, 364–366. Pastorino, F., Brignole, C., Di Paolo, D., Nico, B., Pezzolo, A., Marimpietri, D., Pagnan, G., Piccardi, F., Cilli, M., Longhi, R., Ribatti, D., Corti, A., et al. (2006). Targeting liposomal chemotherapy via both tumor cell–specific and tumor vasculaturespecific ligands potentiates therapeutic efficacy. Cancer Res. 66, 10073–10082. Perabo, L., Buning, H., Kofler, D. M., Ried, M. U., Girod, A., Wendtner, C. M., Enssle, R., and Hallek, M. (2003). In vitro selection of viral vectors with modified tropism: The adeno-associated virus display. Mol. Ther. 8, 151–157. Piersanti, S., Cherubini, G., Martina, Y., Salone, B., Avitabile, D., Grosso, F., Cundari, E., Di Zenzo, G., and Saggio, I. (2004). Mammalian cell transduction and internalization properties of lambda phages displaying the full-length adenoviral penton base or its central domain. J. Mol. Med. 82, 467–476.


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Porkka, K., Laakkonen, P., Hoffman, J. A., Bernasconi, M., and Ruoslahti, E. (2002). A fragment of the HMGN2 protein homes to the nuclei of tumor cells and tumor endothelial cells in vivo. Proc. Natl. Acad. Sci. USA 99, 7444–7449. Poul, M. A., and Marks, J. D. (1999). Targeted gene delivery to mammalian cells by filamentous bacteriophage. J. Mol. Biol. 288, 203–211. Rafii, S., Avecilla, S. T., and Jin, D. K. (2003). Tumor vasculature address book: Identification of stage-specific tumor vessel zip codes by phage display. Cancer Cell 4, 331–333. Rajotte, D., Arap, W., Hagedorn, M., Koivunen, E., Pasqualini, R., and Ruoslahti, E. (1998). Molecular heterogeneity of the vascular endothelium revealed by in vivo phage display. J. Clin. Invest. 102, 430–437. Rajotte, D., and Ruoslahti, E. (1999). Membrane dipeptidase is the receptor for a lungtargeting peptide identified by in vivo phage display. J. Biol. Chem. 274, 11593–11598. Reynolds, P., Dmitriev, I., and Curiel, D. (1999). Insertion of an RGD motif into the HI loop of adenovirus fiber protein alters the distribution of transgene expression of the systemically administered vector. Gene Ther. 6, 1336–1339. Ruoslahti, E. (2004). Vascular zip codes in angiogenesis and metastasis. Biochem. Soc. Trans. 32, 397–402. Schmidt, A., Haas, S. J. P., Hildebrandt, S., Scheibe, J., Eckhoff, B., Racek, T., Kempermann, G., Wree, A., and Putzer, B. M. (2007). Selective targeting of adenoviral vectors to neural precursor cells in the hippocampus of adult mice: New prospects for in situ gene therapy. Stem Cells 25, 2910–2918. Sergeeva, A., Kolonin, M. G., Molldrem, J. J., Pasqualini, R., and Arap, W. (2006). Display technologies: Application for the discovery of drug and gene delivery agents. Adv. Drug Deliv. Rev. 58, 1622–1654. Shi, W. F., and Bartlett, J. S. (2003). RGD inclusion in VP3 provides adeno-associated virus type 2 (AAV2)-based vectors with a heparan sulfate-independent cell entry mechanism. Mol. Ther. 7, 515–525. Smith, G. P., and Petrenko, V. A. (1997). Phage display. Chem. Rev. 97, 391–410. Smith, G. P., and Scott, J. K. (1993). Libraries of peptides and proteins displayed on filamentous phage. Methods Enzymol. 217, 228–257. Somia, N., and Verma, I. M. (2000). Gene therapy: Trials and tribulations. Nat. Rev. Genet. 1, 91–99. Souza, G. R., Christianson, D. R., Staquicini, F. I., Ozawa, M. G., Snyder, E. Y., Sidman, R. L., Miller, J. H., Arap, W., and Pasqualini, R. (2006). Networks of gold nanoparticles and bacteriophage as biological sensors and cell-targeting agents. Proc. Natl. Acad. Sci. USA 103, 1215–1220. St Croix, B., Rago, C., Velculescu, V., Traverso, G., Romans, K. E., Montgomery, E., Lal, A., Riggins, G. J., Lengauer, C., Vogelstein, B., and Kinzler, K. W. (2000). Genes expressed in human tumor endothelium. Science 289, 1197–1202. Sun, L., Chu, T., Wang, Y., and Wang, X. (2007). Radiolabeling and biodistribution of a nasopharyngeal carcinoma-targeting peptide identified by in vivo phage display. Acta Biochim. Biophys. Sin. (Shanghai) 39, 624–632. Thomas, C. E., Ehrhardt, A., and Kay, M. A. (2003). Progress and problems with the use of viral vectors for gene therapy. Nat. Rev. Genet. 4, 346–358. Thorpe, P. E., and Ran, S. (2002). Mapping zip codes in human vasculature. Pharmacogenomics J. 2, 205–206. Trepel, M., Arap, W., and Pasqualini, R. (2000a). Exploring vascular heterogeneity for gene therapy targeting. Gene Ther. 7, 2059–2060. Trepel, M., Arap, W., and Pasqualini, R. (2001). Modulation of the immune response by systemic targeting of antigens to lymph nodes. Cancer Res. 61, 8110–8112. Trepel, M., Arap, W., and Pasqualini, R. (2002). In vivo phage display and vascular heterogeneity: implications for targeted medicine. Curr. Opin. Chem. Biol. 6, 399–404.

106


Trepel, M., Grifman, M., Weitzman, M. D., and Pasqualini, R. (2000b). Molecular adaptors for vascular-targeted adenoviral gene delivery. Hum. Gene Ther. 11, 1971–1981. Waterkamp, D. A., Mu¨ller, O. J., Ying, Y., Trepel, M., and Kleinschmidt, J. A. (2006). Isolation of targeted AAV2 vectors from novel virus display libraries. J. Gene Med. 8, 1307–1319. White, S. J., Nicklin, S. A., Buning, H., Brosnan, M. J., Leike, K., Papadakis, E. D., Hallek, M., and Baker, A. H. (2004). Targeted gene delivery to vascular tissue in vivo by tropism-modified adeno-associated virus vectors. Circulation 109, 513–519. White, S. J., Nicklin, S. A., Sawamura, T., and Baker, A. H. (2001). Identification of peptides that target the endothelial cell–specific LOX-1 receptor. Hypertension 37, 449–455. Work, L. M., Nicklin, S. A., Brain, N. J. R., Dishart, K. L., Von Seggern, D. J., Hallek, M., Buning, H., and Baker, A. H. (2004). Development of efficient viral vectors selective for vascular smooth muscle cells. Mol. Ther. 9, 198–208. Yao, V. J., Ozawa, M. G., Trepel, M., Arap, W., McDonald, D. M., and Pasqualini, R. (2005). Targeting pancreatic islets with phage display assisted by laser pressure catapult microdissection. Am. J. Pathol. 166, 625–636. Zhang, L., Giraudo, E., Hoffman, J. A., Hanahan, D., and Ruoslahti, E. (2006). Lymphatic zip codes in premalignant lesions and tumors. Cancer Res. 66, 5696–5706. Zurita, A. J., Troncoso, P., Cardo-Vila, M., Logothetis, C. J., Pasqualini, R., and Arap, W. (2004). Combinatorial screenings in patients: The interleukin-11 receptor alpha as a candidate target in the progression of human prostate cancer. Cancer Res. 64, 435–439.

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Avian Embryos: A Model for the Study of Primary Vascular Assembly in Warm-Blooded Animals Paul A. Rupp, Mike B. Filla, Cheng Cui, and Charles D. Little Contents 1. 2. 3. 4. 5. 6.

Introduction Culture Insert Preparation Culture Chamber Preparation Construction of Microscope Incubator Construction of Electroporation Chamber Embryonic Culture Insert/Dish Preparation 6.1. Solutions 6.2. Embryo staging and preparation 7. Cell Labeling 7.1. Using antibodies and microinjection for tagging endothelial cells or ECM fibers 7.2. Embryo whole-mount electroporation of DNA plasmids expressing fluorescent proteins 7.3. In ovo electroporation of fluorescent protein expressing constructs 8. Post-Incubation Fixation and Processing 9. Whole-Mount Immunolabeling 10. Plastic Embedding and Sectioning References

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Abstract The formation of a primary vascular bed is a dynamic process, aspects of which are readily amenable to time-lapse imaging in avian embryos. At early developmental stages, the body plan of avian embryos is very similar to mammals and has many properties that make it ideal for imaging. We devised labeling, culturing, and imaging techniques that capture high-resolution images of intact avian embryos in four dimensions over large length scales (1 to 5000 mm). Here, Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03005-X

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we describe multiple techniques for labeling and culturing avian embryos to study the cellular, tissue, and extracellular matrix dynamics of vascular morphogenesis.

1. Introduction We have devised novel methods for studying avian endothelial cell migration in vivo (Czirok et al., 2002; Rupp et al., 2003a,b), as well as the dynamics of extracellular matrix (ECM) assembly and tissue movements (Czirok et al., 2004, 2006; Filla et al., 2004; Zamir et al., 2005, 2006). Our studies have centered on the use of time-lapse microscopy to observe embryos from day 1 until early circulation stages. Avian embryos are excellent specimens for the study of an emergent vascular pattern in warm-blooded animals. Not only do avian embryos have body plans nearly identical to those of mammals at early stages, they are readily accessible, easy to stage, exhibit excellent optical properties, and are inexpensive. Avians possess high-performance cardiovascular systems with four-chambered hearts and complex vessel wall structure virtually identical to those in mammals. Tagging endothelial cells in quail embryos for fluorescence microscopy can be conveniently accomplished using two methods—one entails microinjection of fluorochrome-conjugated antibodies and the other takes advantage of introducing DNA plasmids encoding fluorescent proteins. Each method has advantages and disadvantages. When labeling with antibodies, only the antigen present at the time of microinjection is labeled. If the embryo is used for time-lapse imaging, it is therefore not possible to observe the entire vascular structure unless additional antibody is introduced at intervals throughout the image acquisition experiment. This can be an advantage, since not all cells within the vascular structure will be labeled; making it easier to track specific cells. An additional advantage of injecting fluorescently-conjugated primary antibodies that bears mentioning is that the surrounding ECM and/or extracellular growth factors (e.g., VEGF) can be tagged in the same specimen. This can be accomplished in conjunction with endothelial cell surface labeling for double or even triple labeling protocols. If motion analysis (time-lapse microscopy) is planned, labeling a relevant ECM component will permit analysis of cellular motion with respect to the ECM scaffold—that is, autonomous versus passive (tissue) motility (see Zamir et al. [2006] for similar analysis during gastrulation). Transfection of cells, via microinjection and electroporation, with fluorescent protein-expressing plasmids has its own benefits and pitfalls. Cells can be labeled at very early stages (as early as HH stage 1); however,

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this labeling is nonspecific. To label a distinct population of cells, such as the endothelial cells, the DNA plasmids must contain cell type–specific promoters. As with the microinjection of tagged antibodies, not all endothelial cells will be labeled with this technique. However, all progeny of cells that are transfected will likewise be labeled. In order to study cellular and tissue dynamics in vivo, several technical issues must be addressed, including a clear way to label the cells or molecules of interest, sustaining the specimen’s health, maintaining a clear optical path, and acquiring/processing images over wide length scales. The first three of these issues are discussed below, the latter is not. The imaging software has evolved (Czirok et al., 2001), with the latest software available as opensource code. Here we describe experimental methods to label quail embryos using the endothelial cell–specific QH1 antibody in conjunction with another extracellular epitope (singly or simultaneously). We also describe labeling chicken or quail endothelial cells, and other mesodermal derivatives, using electroporation. Perhaps most useful, we describe approaches proven to maintain the health of the embryos on a microscope stage.

2. Culture Insert Preparation A bed of parallel filaments is used to support an embryo during timelapse imaging and provide optimal differential interference contrast (DIC) microscopy. To create a culture insert, the permeable membrane of a Millicell cell culture insert (PICMORG-50, Millipore, Bedford, MA) is first removed. Parallel grooves approximately 3 mm apart were cut into the cell culture insert using a Dremel tool. Trilene XL Smooth Casting fishing line (6-lb test, 0.23-mm diameter, Berkley, Spirit Lake, IA) was woven through the notches (Fig. 5.1A). This structure provides support for the embryo on its vitelline membrane and allows for unobstructed imaging of early avian embryos (Fig. 5.1B). Two newer systems exists that use stainless steel suture bed inserts, Bioptechs DTculture dishes (Bioptechs, Butler, PA), and culture chambers of our design. (In Fig. 5.2A to D, version 1 is shown, from Rupp et al. [2003b].)

3. Culture Chamber Preparation Depending on the type of time-lapse to be performed, three different culture chambers are used. The first is created from a six-well culture dish, employed when the plastic culture insert (described above) is used for highresolution DIC optics. The chamber is abbreviated as 6-WDCC (6-well DIC

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Figure 5.1 The components of a six-well DIC culture chamber and system (6-WDCC). (A) An image of a plastic culture insert created from a tissue culture insert (Millicel) and fishing line.The filaments are 2 mm apart. (B) An avian embryo, ventral side up, resting on a culture insert within one well of the 6-WDCC. Note that the embryo can be situated such that the filaments do not interfere with the optical path. (C) The culture chamber with four wells available for imaging, and two wells for holding water (humidity control).The entire chamber is 21 mm thick in the vertical direction with the filament bed raising the embryo 3 mm from the bottom. (D) An image of a microscope incubator that encompasses the stage and associated optics of a Leica DMRXA2 TM upright microscope (Leica Microsystems, Wetzlar, Germany). Heated air is piped into the incubator through standard metal ducts and distributed via two manifolds within the incubator (see Czirok et al., 2002) to maintain the embryos at 37 C. (Modified from Rupp, P. A., Rongish, B. J., Czirok, A., and Little, C. D. (2003a). Culturing of avian embryos for time-lapse imaging. Biotechniques 34, 274^278.)

culture chamber), and was originally described by Rupp and colleagues (2003a). Using this arrangement, four embryos are typically imaged simultaneously. In order to image in DIC, the plastic within the optical path is replaced with glass (Fig. 5.1C). A 20-mm cork borer is heated over an open flame and a hole is bored through each of the wells within the base. The rough edges are sanded smooth. A no. 2 coverslip is then glued to the underside of the plate using MarineGoop (Eclectic Products, Pineville, LA) thinned with xylene. A watertight seal must be achieved. The plastic in the culture chamber lid is also replaced using 1-mm thick glass (see Fig. 5.1C). The 6-WDCC is approximately 21 mm thick (vertical height).

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Figure 5.2 The components of a four-well DIC electronic culture chamber and system (4-WDECC). (A) An image of the bottom half of a heated self-contained four-well chamber used for embryo culture and image acquisition.The system consists of a chamber of our design (Rupp et al., 2003b) used in conjunction with Bioptechs culture dishes (left corner) (Bioptechs Inc., Butler, PA), and stainless steel filament bed inserts (upper right corner). (B) An HH stage-10 quail embryo is shown within one chamber on a filament bed. The distance between the filaments is 3 mm. (C) A fully assembled electronic culture chamber for imaging up to four embryos in high-resolution DIC and epifluorescence microscopy. (D) The sealed culture chamber mounted on a Leica DMRXA2 upright microscope (Leica Microsystems,Wetzlar, Germany) and connected to the controller box. (Modified from Rupp, P. A., Czirok, A., and Little, C. D. (2003b). Novel approaches for the study of vascular assembly and morphogenesis in avian embryos.Trends Cardiol. Med. 13, 283^288.)

The second culture chamber, used with embryonic culture inserts (described below), is created from two six-well culture dishes (Fig. 5.3A-B). To begin with, the bottom of one of the culture dishes is sanded down from the top until level with the ridged base. A belt sander used in an area with adequate ventilation works well. The sanded dish is then flipped over and a Dremel tool is used to detach the inner wells from the ridged base. A 20-mm hole is created in the bottom of each detached well set using a heated cork borer, and the rough edges are sanded smooth. Holes are likewise bored through one of the lids and sanded smooth. The detached wells with holes should be used as a template to mark where the holes should be bored in the lid. Cover glasses (no. 2, 25 mm, Fisher Scientific) are glued to the topside of the modified lid using MarineGoop thinned with xylene. The glass-containing lid is then flipped over and the

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Figure 5.3 The components of 6-well narrow and 12-well narrow culture chambers (6-WNCC and 12-WNCC, respectively). (A) The lower half of a 6-WNCC used for imaging embryos on embryonic culture inserts after transfection by electroporation. Displayed to the left of the chamber is a Millicell tissue culture insert used to make embryonic culture inserts (albumen/agar plated) as well as the plastic culture inserts used for DIC optics (shown in Figure 5.1). An insert after the addition of albumen/agar is shown in the upper left well of the chamber. (B) An assembled chamber. Note how the lid is inverted from its usual orientation.This allows for increased objective lens working distance. (C) A 12-WNCC assembled from two 12-well dishes is shown without albumen/agar. (D) A single embryo laid dorsal side down on the solidified culture medium within a 12-WNCC. The entire assembly is 12 mm thick (vertically on the microscope stage).

set of detached wells are aligned within the lid and glued into place. The plastic within the second lid is removed and a sheet of glass glued in its place. The second lid is then flipped over and aligned with the new ‘‘base’’ creating a vertically thin 6-well narrow culture chamber (6-WNCC) with glass in all optical paths (Fig. 5.3B). As with the first chamber, all plastic in the optical path has been replaced. This system was designed to limit the amount of embryonic manipulation required after electroporation, and thus increase the success rate for time-lapse imaging. After electroporation, the embryo need only be flipped over (dorsal side down) upon the embryonic culture insert and then placed in the chamber for imaging. In addition, higher magnification objectives may be used due to the additional working distance created by this thinner chamber (12 mm). A third chamber contains 12 wells, and is created in an identical manner to the 6-WNCC except that 12-well culture dishes are used. The

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12-WNCC does not use the embryonic culture inserts; rather a thin layer of albumen/agar is coated directly onto the glass of each well. Embryos that have been labeled either by injection of antibody or electroporation can then be placed dorsal side down directly into the well (Fig. 5.3D). An advantage with this arrangement is that many more embryos can be imaged; provided there is sufficient image acquisition time for each specimen. Like the 6-WNCC, the 12-well version is vertically thinner (12 mm), providing a greater working distance.

4. Construction of Microscope Incubator To culture and dynamically image embryos, an incubator for use with the 6-WDCC, 6-WNCC, and 12-WNCC was fabricated from cardboard (4 mm thickness) to enclose the optics and stage of a Leica DMRXA2 upright microscope (Fig. 5.1D) (Leica Microsystems, Wetzlar, Germany). A portable heater and standard ductwork are used to direct heated air into the incubator and across the culture chamber. The temperature within the incubator is maintained at 38.5 C using a Love 1600 controller unit (Dwyer Instruments, Michigan City, IN) with thermocouple sensors (PT6, Physitemp Instruments, Clifton, NJ). The culture media within the 6-WDCC is kept at 37.5 C, as calibrated with liquid crystal thermometer foils (Edmund Industrial Optics, Barrington, NJ). The design is similar to that of Kulesa and colleagues (1999), and modified from Czirok and colleagues (2001) and Rupp and colleagues (2003a).

5. Construction of Electroporation Chamber Cui and colleagues (2006, 2007) provide a thorough description of the construction of an electroporation chamber (Fig. 5.4A) for whole-mount embryo electroporation. Briefly, a 100-mm Petri dish with a 20-mm hole is aligned underneath a 60-mm dish with a similar hole and glued together with xylene-thinned MarineGoop. A no. 2 cover glass is glued to the bottom of the 100-mm Petri dish. The glass bottom allows for precise positioning of the embryo above the anode. The anode (þ) is prepared by running a 76- 0.25-mm platinum wire (A-M Systems, Carlsborg, WA) along the bottom of the 60-mm Petri dish. The anode is bent such that a 3-mm section over the middle of the bored hole is raised 1 mm above the floor of the Petri dish (Fig. 5.4A to D and G). One end of the anode extends through holes at the base of both dishes and then extends up the outer wall of the Petri dish to be connected to the power supply. Clear nail polish is used to seal the holes, to fix the anode in place, and to insulate the anode

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Figure 5.4 Images and diagram of the electroporation chamber. (A) The electroporation chamber (60-mm dish) is contained within a100-mm dish.This allows for the placement of a weighted ring in the outer chamber to provide stability to the overall device. The platinum wire electrodes are insulated by clear nail polish (tinted cyan in these images). (B) The chamber during electroporation. (C) The cathode (^) positioned by a micromanipulator in the flat orientation for electroporation of a ‘‘large’’area. (D) The cathode (^) positioned by a micromanipulator in the point orientation to focally label a select group of cells. (E) An embryo 3.5 h post-electroporation of a nuclear-GFP expressing (H2B-GFP, Rusty Lansford, California Institute of Technology) DNA plasmid. The entire interstitial space between the vitelline membrane and the epiblast was flooded with plasmid.The cathode was used in the flat orientation with each side of the embryo being electroporated at different voltages (3 Von the right and 4 Von the left). Note how there are fewer GFP-labeled cells on the right side (3 V) than on the left side (4 V) demonstrating that differences in voltage will affect transfection efficiency. Scale bar, 250 mm. (F) A small group of clustered cells labeled when a small amount of plasmid DNA was introduced into a localized region. The cathode was in the flat orientation.

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except for the raised 3-mm ‘‘working’’ section above the windowed hole. A cathode (–) is prepared by bending a 3- 0.25-mm platinum wire (A-M Systems, Carlsborg, WA) at a 45-degree angle approximately 2 mm from one end. The shorter of the two ends is coated with clear nail polish to act as an insulator when mounted on a micromanipulator. The cathode is coupled to a thin stainless steel rod and mounted on a micromanipulator. Alternatively, the platinum wire can be fed through a glass microcapillary, which is glued within the base of an 18-gauge needle (steel needle has been removed; see Fig. 5.4B).

6. Embryonic Culture Insert/Dish Preparation 6.1. Solutions Agar-salt solution: Make and sterilize 0.6% agar (Becton Dickson, 214010) in 123 mM NaCl (Fisher Scientific, S271-10). Aliquots are stored at room temperature. Egg albumen preparation: Egg albumen isolated from fresh chicken eggs that have been incubated 12 to 24 h at 37.5 to 38.5 C is homogenized using a Wheaton 40-ml glass homogenizer. The homogenized egg albumen is heated to 56 C for 40 min prior to aliquoting and storing at – 20 C (good for 1 month). Egg albumen/glucose solution: Just prior to pouring plates, 1.5 ml 10% glucose is added to 48.5 ml of homogenized and heat-treated chicken albumen. The preparation of culture dishes for ex ovo incubation of embryos has been modified from the ‘‘EC culture’’ described by Chapman and colleagues (2001). Briefly, both the agar-salt and egg albumen/glucose solutions are heated to 56 C for at least 10 min. The equilibrated solutions are homogeneously mixed in a 1:1 ratio and immediately distributed as follows: 2 ml per 35-mm plate; 1 ml per Millicell tissue culture insert (PICMORG-50, Millipore, Bedford, MA); or 1 ml per well of the 12-WNCC. The thin (nonviscous) albumen is recommended for use when the embryo is to be imaged using time-lapse microscopy because it is more transparent. Store the solidified plates in a humidified chamber at 4 C for up to 1 week. Standard

Alternatively, the cathode could be positioned in the point orientation to achieve similar results. Scale bar, 250 mm. (G) A cross-section of the electroporation chamber used to electroporate pre-gastrulation stage avian embryos. (Modified from Cui, C., Lansford, R., Filla, M. B., Little, C. D., Cheuvront,T. J., and Rongish, B. J. (2006). Electroporation and EGFP labeling of gastrulating quail embryos. Dev. Dyn. 235, 2802^2810; and Cui, C., Rongish, B. J., Little, C.D., and Lansford, R. (2007). Ex ovo electroporation of DNA vectors into pre-gastrulation avian embryos. CSH Protocols doi:10.1101/pdb.prot4894.)

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35-mm embryonic culture dishes are used for microinjection of antibody followed by non-imaging incubations. The embryonic culture inserts (albumen/agar on tissue culture inserts) are used for electroporation of plasmid DNA and subsequent nonimaging incubations or for imaging within the 6-WNCC. The 12-WNCC with embryonic culture medium is used for time-lapse imaging when higher magnification is needed and to increase the number of samples.

6.2. Embryo staging and preparation Incubate fertile quail eggs (Coturnix coturnix japonica) or chicken eggs in a humidified incubator at 38 C until the appropriate stages as determined by Hamburger and Hamilton (HH) (1951, reprinted in 1992). The embryos are manipulated ex ovo by mounting them to paper rings as described by Chapman et al. (2001) and Rupp et al. (2003a). Briefly, the incubated eggs are opened carefully to ensure that the vitelline membranes on the dorsal aspect of the embryos remain intact. The albumen and yolk (with embryo) are gently poured into a sterile Petri dish. A transfer pipette is used to remove the viscous albumen exposing the vitelline membrane. Kimwipe Tissues (Kimtech Science, Kimberly-Clark Global Sales, Roswell, GA) may be used to remove any additional albumen from the surface of the yolk. An embryo is centered within a Whatman 52 filter paper ring (Whatman International, Maidstone, England) having an inner diameter slightly larger than the size of the embryo. The ring is allowed to adhere to the membrane for 1 min. The ring is cut along the perimeter of the ring beginning at the caudal end of the embryo using an angled iris scissors (Fine Science Tools, Foster City, CA). The embryo is gently pulled free by grasping the paper ring at the caudal end and pulling at a low degree of angle. The embryo is carefully submerged ventral side up into ePBS to remove yolk. A transfer pipette is used to help rinse any adhering yolk by gently pulsing ePBS across the surface of the embryo. Once free of yolk, the embryos are placed onto the embryonic culture dishes (Chapman et al., 2001) for labeling by antibody injection or onto embryonic culture inserts for microinjection and electroporation. Embryos are placed ventral side up for labeling with antibodies and ventral side down during electroporation.

7. Cell Labeling 7.1. Using antibodies and microinjection for tagging endothelial cells or ECM fibers The conjugation of primary antibodies with fluorochromes can be used to label your cell or molecule of interest. The technique is used for labeling embryos using the in vivo whole-mount cultures. We employ Alexa-488,

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Alexa-555, or Alexa-647 (Invitrogen, CA) fluorochrome-conjugated primary QH1 antibodies (Developmental Studies Hybridoma Bank, University of Iowa, Ames, IA) to study the behavior of vascular endothelial cells in the developing quail. QH1 is not effective in labeling chicken endothelial cells. For labeling ECM fibers in either quail or chicken, we use an Alexa488, Alexa-555, or Alexa-647 conjugated nonperturbing primary fibrillin2 antibody ( JB3) or a similarly conjugated fibronectin antibody (B3D6) (both from the Developmental Studies Hybridoma Bank). Depending on the stage of the embryo to be injected, the antibodies are delivered by microinjection of 5- to 25-nl volumes 1 to 16 times per embryo, at 1 ng/nl concentrations. Delivery of the antibodies into the interstitial space is accomplished using a micropipette (18-mm bore) and a pneumatically driven Pico-Injector (Harvard Apparatus, Holliston, MA) mounted to a hydraulic micromanipulator assembly (Narishige Scientific Instrument Laboratory, Tokyo). The needle enters the interstitial space at an acute angle (less than 45 degrees) and the antibody is introduced in 5- to 10-ms pulses with a pressure of 5.0 psi or less.

7.2. Embryo whole-mount electroporation of DNA plasmids expressing fluorescent proteins 7.2.1. Solutions Hanks balanced salt solution (HBSS): Fisher Scientific-CellGro, 21020-CV Embryonic phosphate buffered saline (ePBS, pH 7.4): 137 mM NaCl, 2.69 mM KCl, 8.1 mM Na2HPO4, 1.47 mM KH2PO4, 0.68 mM CaCl2, and 0.49 mM MgCl2 Buffered phenol red solution: 1.0 ml 10X PBS (1.37 M NaCl, 26.8 mM KCl, 81.0 mM Na2HPO4, 15.0 mM KH2PO4, pH 7.2), 10 ml of 1.0 M MgCl2, 5.0 ml of 0.4% phenol red (Fisher Cat no. P-391), and 3.890 ml endotoxin-free water DNA plasmid preparation: 1:1 ratio of buffered phenol red solution and DNA plasmid (endotoxin-free water) for a final concentration of 2.5 mg/ml. Microinjection and electroporation are used to transfect cells with DNA plasmids encoding a fluorescent protein as early as HH stage 1. A large area of the ectoderm can be transfected using a flat cathode in conjunction with the anode (Fig. 5.4C and E). Alternatively, a very focal labeling occurs if the cathode is rotated so that the point of the cathode is directed downward (Fig. 5.4D). The electroporation chamber is tested for continuity by placing an embryonic culture insert (minus any embryo) into it and performing a mock electroporation. The chamber is filled with HBSS to a level so that the anode is covered, but such that embryos will not be submerged when

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present. The cathode is placed parallel to the anode and approximately 3 mm above it (Fig. 5.4B and C). The leads of the electrodes are attached to a CUY-21 Square Wave power supply (BEX Co., Tokyo), which is programmed with the following pulse sequence: 5V, 40 ms power-on and 900 ms power-off for a series of four pulses. Bubbles should be observed on the cathode and recorded amperage of at least 0.01 A displayed. A glass micropipette is used to introduce the plasmid DNA/phenol red solution into the space between the epiblast/embryo and the vitelline membrane and should be prepared just prior to starting the injections (Fig. 5.4B). A concentration of 2.5 mg/ml is general guideline for microinjection of fluorescent protein expressing plasmids used for cell labeling. However, if constructs for overexpression or knockdown are used, the toxicity of the plasmids must be determined empirically. An embryo (quail or chicken) is positioned ventral side down on an embryonic culture insert and then placed into the electroporation chamber. The insert is positioned so that the anode is below the site of interest. As with the antibody injection, the needle penetrates the vitelline membrane at a low degree of angle (

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