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 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

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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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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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

#

2008 Elsevier Inc. All rights reserved.

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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).

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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

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CD31 & podoplanin & DAPI

Vascular Remodeling during Inflammation

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.).

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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

* {

28 29 32 36 37 39 41 41 42 42 42 43 43 44 44 44 45 45 45 46

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

#

2008 Elsevier Inc. All rights reserved.

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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

46 47 47 47 47 48 48 49 49 50

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

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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

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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

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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-GFP^expressing 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-GFP^expressing 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-GFP^expressing 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-GFP^expressing 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-GFP^expressing 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

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Cell trafficking

Axillary lymph node

Inject into inguinal lymph node

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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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D

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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

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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

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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.

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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.

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T H R E E

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

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03003-6

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2008 Elsevier Inc. All rights reserved.

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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

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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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Methods

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.

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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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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

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03004-8

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2008 Elsevier Inc. All rights reserved.

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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).

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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 (<45 degrees). The DNA plasmid (20 to 100 nl) is then introduced by adjusting the Pico-Injector’s P-balance setting to expel the DNA/phenol red into the area of interest. The P-balance allows the solution to be injected slowly (5 s) and with very low pressure (<1.0 psi) so as to not damage the delicate embryonic tissue. The needle is slowly withdrawn from the embryo and the cathode placed parallel to the anode. The cathode should be positioned as close to the embryo as possible without making contact. After placing two drops of HBSS on the vitelline membrane, the cathode is lowered until a slight tissue deformation is observed. The cathode is slowly raised until the deformation has disappeared. With the cathode in the flat orientation, the CUY-21 power supply delivers four square electrical pulses of 4 V, 40 ms each at 900-ms intervals. If the cathode is positioned in the point orientation for a more focal labeling, the pulse sequence is as follows: 3V, 40 ms power-on and 900 ms power-off for four to six pulses. Check for the formation of bubbles at the cathode as well as a change in the color of the phenol red from pink to red. The embryo culture insert may now be removed from the electroporation chamber. At this point, the embryo is flipped over (dorsal side down) onto the embryonic culture insert, placed in a six-well dish, and incubated at 37
C for the appropriate time (up to 40 h). If the embryos are to be used for time-lapse imaging, they are screened at 1 to 2 h of incubation to check for the presence of fluorescence. The selected embryos, on the embryonic culture inserts, may then be placed directly into the 6-WNCC for imaging. Alternatively, if more than six embryos are to be imaged, the 12-WNCC may be used. Well-labeled embryos are floated free of the embryonic culture inserts by placing them in ePBS and then positioned dorsal side down into the 12-WNCC. After filling any unused wells with sterile water,

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the edge of the culture chamber is sealed shut with electrical tape and placed onto the stage within the microscope incubator. It may take a few hours before sufficient levels of fluorescent protein are produced for imaging depending on the strength of the promoters used. This will also depend on the sensitivity of the image detector being used. In addition to the orientation of the cathode (flat or point), the amount and placement of the DNA plasmid determines the area/number of cells transfected. In Fig. 5.4E, the interstitial space between the vitelline membrane and epiblast was flooded with a nuclear-localizing GFP expressing plasmid and then each side electroporated with the cathode in the flat position. In contrast, a small population of cells was labeled in Fig. 5.4F when a small bolus of plasmid was present. Another caveat that alters the efficiency of transfection is the voltage setting: The right side of the embryo in Fig. 5.4E was electroporated with 3 V and the left side with 4 V. The electroporation as described will label epiblastic/ectodermal cells. In order to label a population of endothelial cells, embryos are electroporated at very early stages (HH stage 3 and younger) prior to gastrulation; that is, prior to formation of the mesoderm, this will eventually result in labeled endothelial cells as well as other mesodermal derivatives (unpublished observations). Labeling cells in the center to lower portion of the primitive streak prior to HH stage 3 is optimal for ensuring that endothelial cells will be labeled. DNA plasmids with endothelial cell–specific promoters would obviously label only the early vascular progenitor cells and their daughters.

7.3. In ovo electroporation of fluorescent protein expressing constructs In ovo electroporation of quail embryos is difficult due to the size of the egg. Therefore, use of chicken eggs is recommended, at least while learning the technique. Eggs incubated to the appropriate stage are rinsed with 70% ethanol and allowed to dry. Using a 5-ml syringe with an 18-gauge needle, 2 to 3 ml of albumen are removed from the top of one end of the egg. A window is cut into the top of the shell using an angled iris scissors (Fine Science Tools). Plasmid DNA is introduced between the vitelline membrane and the embryo as described in the embryo whole-mount cultures. India ink, diluted 1:10 in ePBS, can be injected below the embryo to aid in visualizing the structures. For in ovo electroporation, it is recommended that tungsten needles be used instead of platinum as the former appear to yield better labeling results (unpublished observations). Two electrodes are prepared by bending the last 2 mm of a 0.25-mm diameter tungsten wire (A-M Systems, Calsborg, WA) at a 45-degree angle. The wire can be insulated using heat-shrink tubing found at an electrical supply store. For more stability, the straight end of the electrode can be soldered to a male pin connector. Alternatively, the

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tungsten wire may be fed through a glass microcapillary tube with one end bent and the other folded down the outside of the capillary and taped in place with electrical tape. A third alternative is to purchase premade 0.25mm–diameter Tungsten Epoxy-Insulated Microelectrodes with tapered tips (A-M Systems, Carlsborg, WA). The surface of the vitelline membrane is moistened with a small amount of ePBS. The anode is then gently placed below the area of interest by inserting the wire through the vitelline membrane, inserting through the extraembryonic region, and rotating it into position. To get the correct depth, pull the anode up until it is visible under the embryo and then lower it until just visible. As with the embryo whole-mount electroporation the cathode can be placed in two orientations (flat or point). To label a broader area, lower the cathode in the flat orientation parallel to the just visible anode. The anode must be close to, but not touching, the embryo. The CUY-21 Square Wave power supply is used to deliver four pulses of 7 V, 40 ms each at 900- ms intervals. To label a very discrete narrow population of cells, the point orientation is used with the following pulse sequence: 4 V, 40 ms power-on and 900 ms power-off for six pulses. After the embryos have been injected and electroporated, the egg is sealed shut using a rectangular stretched piece of Parafilm (American National Can, Greenwich, CT). The eggs are returned to a 37
C incubator for the appropriate time.

8. Post-Incubation Fixation and Processing In order to preserve embryos post-incubation and allow further analysis, embryos are fixed in a solution of paraformaldehyde (PFA) in PBS. To begin, each specimen is transferred to an individual well of a 24-well plate containing 300 ml of 3% PFA at 4
C. Gently pipette additional cold PFA into each well and carefully submerge the embryos. Allow the embryos to fix for 30 min with gentle agitation at 4
C. If embryos have been incubated on the embryonic culture inserts, the embryos can be floated off of the albumen/agar by the addition of PBS. After fixation and removal from the vitelline membrane, successive dehydration and rehydration is essential for removing lipids and making the specimens permeable to antibodies and other labeling agents. After gently removing the PFA, the specimens are washed twice with PBS þ 0.05% azide (PBSa) at 4
C for 10 min each. The wash is replaced with 50% methanol in PBSa and then returned to 4
C for 15 min. The solution is aspirated off and replaced with 100% methanol at 4
C for a minimum of 30 min. The embryos are rehydrated through a series of decreasing ethanol concentrations beginning with 90%, followed by 70, 50, and 30%. Each step occurs at 4
C for a minimum of 10 min. When the 30% incubation is complete, an equal amount of PBSa is added to

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the wells and incubated for an additional 10 min at 4
C. The embryos are then washed two times in PBSa. For storage, the embryos can be kept in PBSa at 4
C, in the dark, for up to 1 year. Once the embryos are fixed and rehydrated, they may undergo whole-mount immunolabeling and then be subsequently embedded in plastic and sectioned (see following protocols).

9. Whole-Mount Immunolabeling Using the microinjection techniques described above, when a ‘‘pulse’’ of antibody or plasmid is introduced, only cells present at that time (or their daughters in the case of plasmids) will be marked. The entire embryo is often immunolabeled at the end of an experimental protocol in order to visualize the entire vasculature structure, matrix scaffold, or any marker of interest. The first step is to fix and process the embryo as described above, leaving them in PBSa. The embryos are then incubated in bovine serum albumen (BSA) ‘‘blocking’’ solution (3% BSA in PBSa) at 4
C for a minimum of 1 h while gently shaking. A primary antibody is diluted in the blocking solution at the appropriate ratio determined for that antibody. The blocking solution is replaced with the diluted primary antibody and incubated for 2 h at 4
C. The embryos are then washed three times with PBSa for at least 30 min each. If the primary antibody used was directly conjugated with a fluorochrome, the embryos can now be mounted on a slide and imaged. However if a secondary antibody is necessary for visualization, select and dilute the appropriate one. The embryos should be exposed to the secondary at 4
C for 2 h in the appropriate amount of BSA blocking buffer. The washes are then repeated as before. The embryos are now ready for imaging and can be mounted on a slide in a glycerol-based, anti-bleaching mounting solution or in PBSa. If high levels of nonspecific binding occur with the secondary antibody, alternative methods of blocking may be required. An important item to point out is that dehydration/delipidation with methanol will result in the loss of any green fluorescent protein (GFP) or red fluorescent proteins (RFP) signal. A solution of Triton X-100 (SigmaAldrich, St. Louis, MO) can be used as an alternative to methanol. A method to permeabilize fixed cells is found in the book ‘‘Antibodies: A Laboratory Manual’’ by Harlow and Lane (1988), and involves incubating the cells for 2 to 15 min at room temperature in 0.2% Triton X-100. We have found that to permeabilize fixed cells within an embryo, a higher concentration (up to 3.0%) of Triton X-100 is needed. The embryos are not as durable when treated with Triton X-100 as they are with the methanol treatment. A second option is to use the methanol protocol of dehydration/ delipidation, and then reacquire the GFP or RFP signal by using an anti-GFP or anti-RFP antibody to localize the protein.

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10. Plastic Embedding and Sectioning To view vascular labeling in cross-section, embryos may be embedded in plastic and sectioned. The embryos must be dehydrated for the embedding process. The specimens are exposed to a series of increasing ethanol concentration starting at 30%, followed by 70, 80, 95, and 100% for a minimum of 10 min. During the dehydration, the infiltration solution is prepared by adding 0.25 g dry catalyst (benzoyl peroxide, plasticized) to 20 ml of JB-4 Solution A (Electron Microscopy Sciences, Hatfield, PA) and mixed on a stir plate for 15 min. The infiltration solution may be stored for up to 2 weeks in the dark at 4
C. The embryos are individually soaked overnight in 1 ml infiltration solution in the dark at either room temperature or at 4
C. The embedding solution is prepared by adding 60 ml of JB-4 Solution B to 1.5 ml of cold infiltration solution per embryo and vigorously mixing for 30 s. Mark the cranial portion of each paper ring with a tissuemarking dye. The specimens are than submerged (paper rings and all) into the embedding solution within the mold plate wells. A fiduciary marker, in the form of a stiff dark-colored paintbrush hair, is placed 2 mm to the right of the embryo axis on the ventral side of each specimen. The embryos are incubated at room temperature in a vacuum oven in the dark for 1 h to allow the plastic to harden. Once hardened, a fresh razor blade is used to cut out 3 mm  3 mm  6 mm blocks of plastic containing the specimens. The blocks are placed into fresh wells of a mold plate, oriented appropriately, and covered with 2 mm of fresh embedding solution. The specimens are mounted on plastic microtome chucks (EBH2 block holder, Electron Microscopy Sciences, Hatfield, PA). Make sure that the central hole of the chuck is filled with embedding solution and that the orientation of the specimen does not change. Placing the samples at room temperature in a vacuum oven in the dark for 1 h once again hardens the plastic. The specimens are then sectioned using a microtome to produce 1.0- to 15.0-micron thick samples that are mounted on slides for viewing and imaging.

REFERENCES Chapman, S. C., Collignon, J., Schoenwolf, G. C., and Lumsden, A. (2001). Improved method for chick whole-embryo culture using a filter paper carrier. Dev. Dyn. 220, 284–289. 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. 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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Czirok, A., Rupp, P. A., Rongish, B. J., and Little, C. D. (2001). Multi-field 3D scanning light microscopy of early embryogenesis. J. Microsc. 206, 209–217. Czirok, A., Rongish, B. J., and Little, C. D. (2004). Extracellular matrix dynamics during vertebrate axis formation. Dev. Biol. 268, 111–122. Czirok, A., Zamir, E. A., Filla, M. B., Little, C. D., and Rongish, B. J. (2006). Extracellular matrix macroassembly dynamics in early vertebrate embryos. Dev. Biol.Curr. Top. Dev. Biol. 73, 237–258. Filla, M. B., Czirok, A., Zamir, E. A., Little, C. D., Cheuvront, T. J., and Rongish, B. J. (2004). Dynamic imaging of cell, extracellular matrix, and tissue movements during avian vertebral axis patterning. Birth Defects Res. C Embryo Today 72, 267–276. Hamburger, V., and Hamilton, H. L. (1951). A series of normal stages in the development of the chick embryo. J. Morphol. 88, 49–92. Hamburger, V., and Hamilton, H. L. (1992). A series of normal stages in the development of the chick embryo: 1951. Dev. Dyn. 195, 231–272. Harlow, E., and Lane, D. (1988). ‘‘Antibodies: A Laboratory Manual.’’ Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Kulesa, P. M., and Fraser, S. E. (1999). Confocal imaging of living cells in intact embryos. Methods Mol. Biol. 122, 205–222. 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. 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. Zamir, E. A., Czirok, A., Rongish, B. J., and Little, C. D. (2005). A digital image–based method for computational tissue fate mapping during early avian morphogenesis. Ann. Biomed. Eng. 33, 854–865. Zamir, E. A., Czirok, A., Cui, C., Little, C. D., and Rongish, B. J. (2006). Mesodermal cell displacements during avian gastrulation are due to both individual cell-autonomous and convective tissue movements. Proc. Natl. Acad. Sci. USA 103, 19806–19811.

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Mouse Models to Investigate Anti-Cancer Effects of VEGF Inhibitors Yongping Crawford and Napoleone Ferrara Contents 125 127 129 130 131 132 133 134 135

1. Introduction 2. Xenografts 3. Genetic Models 3.1. Apcþ/min mice 3.2. MEN1 mice 3.3. RIP-Tag model 3.4. Humanized VEGF mouse model 4. Conclusions References

Abstract Angiogenesis, the growth of new blood vessels, is required for a variety of normal proliferative processes. Furthermore, it is well established that angiogenesis plays an important role also in neoplastic growth and metastasis. Numerous regulators of angiogenesis have been identified and characterized over the last decades. Among these, vascular endothelial growth factor (VEGF)-A appears especially important in several pathophysiological processes. Several VEGF inhibitors have been approved by the Food and Drug Administration for the treatment of tumors or age-related macular degeneration. This chapter examines the various mouse tumor models in which VEGF inhibitors have been tested and the lessons learned from these studies.

1. Introduction Angiogenesis results in the establishment of microvascular networks, required for pre- and post-natal normal development and for tissue repair in the adult (Carmeliet, 2003; Folkman and Klagsbrun, 1987; Red-Horse Genentech, Inc., South San Francisco, California Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03006-1

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2008 Elsevier Inc. All rights reserved.

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et al., 2007; Risau, 1997). Likewise, angiogenesis is crucial for the development of primary tumors and metastases (Algire et al., 1945; Ferrara and Kerbel, 2005; Folkman, 1995). Without the onset of angiogenesis, most tumors cannot grow beyond 1 to 2 mm due to diffusion limitations (Carmeliet, 2003; Gimbrone et al.). To activate and sustain angiogenesis, tumor cells appear to utilize developmental programs via upregulation of proangiogenic factors and, possibly, downregulation of inhibitory ones (Ferrara, 2002). One of the most important angiogenic factors is VEGF-A (hereafter, VEGF), which also may increase microvascular permeability (Ferrara and Kerbel, 2005; Red-Horse et al., 2007). Even a single allelic loss of VEGF results in defective vasculatures and embryonic lethality (Carmeliet et al., 1996; Ferrara et al., 1996). In healthy adult tissues (with the exception of the female reproductive system), VEGF expression is relative low. However, overexpression of VEGF has been frequently observed in human tumors (Dvorak, 2002; Kiselyov et al., 2007). In fact, increased VEGF expression was found to correlate with tumor progression, recurrences, and decreased survival (Poon et al., 2001; Zhang et al., 2007). VEGF belongs to a gene family that also includes VEGF-B, C, D, E, and PlGF (Kowanetz and Ferrara, 2006). Multiple isoforms of VEGF, ranging from 121 to 206 amino acids, can be generated by alternative exon splicing (Kowanetz and Ferrara, 2006). These isoforms differ in their ability to bind heparin, which determines their bioavailability, and may play distinct roles in angiogenesis during development (Red-Horse et al., 2007). In addition, extracellular proteolysis also may regulate VEGF activity. Plasmin is able to cleave heparin-binding VEGF isoforms at the COOH terminus to generate bioactive and diffusible fragments (Houck et al., 1992; Keyt et al., 1996). More recently, MMP3 has been also reported to generate VEGF proteolytic fragments, which are biologically and biochemically very similar to those resulting from plasmin action (Lee et al., 2005). All VEGF isoforms can bind to two receptors, VEGFR-1 and VEGFR-2. Despite the fact that VEGF binds to VEGFR1 with 10-fold higher affinity than VEGFR2, it is mainly VEGFR2 that mediates VEGF signaling in endothelial cells (Shibuya, 2006; Shibuya and Claesson-welsh, 2006). Hence, many efforts have been made toward targeting the VEGF/VEGFR2 pathway for the treatment of cancer and other disorders such as age-related macular degeneration. Several strategies have been employed to inhibit the VEGF/VEGFR pathway. These include VEGF function blocking antibodies, chimeric soluble receptors (VEGF-Trap), anti-VEGF synthetic aptamers, VEGFR2 blocking antibodies, a variety of small molecule VEGFR2 kinase inhibitors, and siRNAs targeting VEGF or its receptors (Ferrara et al., 2007; Kiselyov et al., 2007). Two VEGF receptor tyrosine kinase inhibitors (RTKI), Sorafenib (Bayer and Onyx Pharmaceuticals) and Sunitinib (Pfizer), an anti-VEGF RNA aptamer, Pegaptanib (Macugen, Pfizer), and

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two humanized VEGF function–blocking antibodies, bevacizumab (Avastin, Genentech) and ranibizumab (Lucentis, Genentech) have been approved by the Food and Drug Administration (FDA) for the therapy of cancer or age-related macular degeneration. These agents have different mechanisms of action. Sunitinib inhibits various RTKs such as PDGFRs or c-Kit in addition to VEGFRs (Faivre et al., 2007). Sorafenib was initially characterized as a raf kinase inhibitor and was later shown to inhibit VEGFR signaling as well (Wilhelm et al., 2004). Pegaptanib is an aptamer that targets the heparin-binding domain of VEGF165 (Ng et al., 2006), and thus does not neutralize alternatively spliced VEGF121 or plasmin- or MMPgenerated VEGF165 fragments. Bevacizumab and ranibizumab block all isoforms and bioactive proteolytic fragments of human VEGF (Ferrara et al., 2004). Bevacizumab is the first antiangiogenic agent to be approved by FDA and is currently approved as therapy for previously untreated and relapsed metastatic colorectal cancers, nonsquamous, non–small cell lung carcinoma, and most recently for metastatic breast cancer, in combination with chemotherapeutic agents. Details of various VEGF/VEGFR inhibitors and their mechanisms of action can be found elsewhere (Ferrara et al., 2007; Kiselyov et al., 2007). The ongoing clinical trials with such inhibitors are outlined at http://www. cancer.gov/clinicaltrials/developments/anti-angio-table. Here, we will focus on discussing mouse tumor models to test the efficacy of various VEGF inhibitors, with an emphasis on monoclonal antibodies. Animal models to assess the effects of anti-VEGF agents and other angiogenesis inhibitors in intraocular neovascularization and other pathological conditions are presented elsewhere (Adamis et al., 1996; Aiello et al., 1995; Campa et al., 2008; Saishin et al., 2003).

2. Xenografts Subcutaneous and orthotopic models have been employed to test the effects of inhibitors of the VEGF/VEGFR pathway on growth of a variety of tumor cell lines. Mab A4.6.1, the murine precursor of bevacizumab, was initially shown to suppress the growth of human rhabdomyosarcoma, glioblastoma, and leiomyosarcoma cells implanted in nude mice, while it had no direct effects on tumor cell proliferation (Kim et al., 1993). These data indicated that the primary effects of MAb A4.6.1 are on tumor endothelial cells, rather than on tumor cells. Since then, Mab A4.6.1/ bevacizumab has been tested on a wide range of human tumor cells implanted subcutaneously or orthotopically (Gerber and Ferrara, 2005). Collectively, these studies demonstrate that Mab A4.6.1/bevacizumab is effective in reducing tumor vessel density and suppressing tumor growth

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even as a single agent, regardless of tumor location and route of administrations. The tumor growth inhibition induced by bevacizumab (or Mab A4.6.1) ranged from 25 to 95%, depending on cell types and therapeutic regimens. Moreover, the growth inhibitory effects of bevacizumab or A4.6.1 are not limited to primary tumors. Several studies also documented similar inhibitory effects of bevacizumab/A4.6.1 on metastasis (Rowe et al., 2000; Warren et al., 1995). One early study involved the injection of HM7 human colorectal cancer cells in the spleen of nude mice (Warren et al., 1995). The tumor cells homed in the liver, creating a model of experimental liver metastasis. Mab A4.6.1 administration resulted in a substantial reduction of metastatic burden (Warren et al., 1995). While most xenograft studies focused on tumor growth inhibition, some studies indicated that bevacizumab or Mab A4.6.1 is able to prolong survival as well (Rubenstein et al., 2000). A confounding factor in assessing the efficacy of bevacizumab or Mab A.4.6.1 in human xenograft models is the species specificity and inability of these antibodies to neutralize host-derived murine VEGF (Yu et al., 2008). Interestingly, the extent of stromal cell recruitment is tumor dependent, and the VEGF produced by such host cells can be a major driver of tumor angiogenesis and growth. The A673 rhabdomyosarcoma is associated with a relatively modest stromal cell infiltration, whereas the CALU-6 lung carcinoma cell line induces a strong desmoplastic reaction (Gerber et al., 2000; Tejada et al., 2006). Accordingly, Mab A.4.6.1 had marked inhibitory effects on the in vivo growth of A673 cells, but it had little effect on CALU-6 growth (Gerber et al., 2000; Tejada et al., 2006). Flt(1-3)-IgG, a chimeric soluble receptor which binds both human and mouse VEGFs, showed greater antitumor efficacy than Mab A.4.6.1 or bevacizumab in both models (Gerber et al., 2000; Tejada et al., 2006). The availability of cross-reactive phage-derived antibodies, which can effectively neutralize mouse and human VEGF (Liang et al., 2006), has enabled more complete VEGF blockade not only in xenograft models but also in long-term studies in genetic mouse models. Using such cross-reactive antibodies, Shojaei et al. (2007) examined the differences among various syngeneic murine tumor cell lines in terms of responsiveness to VEGF blockade. They found that tumor cells that are relatively insensitive to VEGF blockade exhibit greater ability to recruit CD11bþGrþ myeloid cells compared to the sensitive ones. Additive or synergistic effects of bevacizumab with standard chemo- or radio-therapies have been observed in tumor models as well as clinical trials. Fujita et al. (2007) examined the antitumor effects of bevacizumab in combination with paclitaxel on head and neck squamous cell carcinomas (HNSCC) in murine models. They showed that bevacizumab was effective as a single agent in suppressing tumor growth, while paclitaxel was ineffective. A combination treatment was able to further suppress HNSCC xenografts. In a different study, Fox et al. (2002) reported a 98% growth

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inhibition when bevacizumab was combined with paclitaxel in treating androgen-independent xenograft model of prostate cancer. Similar additive or synergistic effects of bevacizumab in combination with paclitaxel were found in other xenograft studies (Hu et al., 2002). Additive and/or synergistic effects have also been reported between bevacizumab and several commonly used chemotherapeutic agents such as doxorubicin, topotecan, docetaxel, and 5-fluorouracil (5-FU) in a variety of human tumor xenografts (Gerber and Ferrara, 2005). Reductions in both primary tumor growth and metastases have been reported in the combinational studies (Hung, 2007; Soffer et al., 2001). An intensely debated issue is the mechanism of such synergistic interactions between bevacizumab and cytotoxic agents or radiotherapy. Since they act via different mechanisms of action, they may have additive antitumor activities by targeting both tumor cells and endothelial cells (Ferrara and Kerbel, 2005; Kerbel, 2006; Red-Horse et al., 2007). Some therapies such as radiation could increase VEGF expression (Gorski et al., 1999). Hence, blocking the increase in VEGF might explain additive or synergistic antitumor effects. Another hypothesis postulates that abnormal and leaky tumor vessels would result in increased interstitial fluid pressure (IFP), impaired flow, and hypoxia. ‘‘Normalization’’ of tumor vessels by bevacizumab or other antiangiogenic agents could improve flow and oxygenation, reduce IFP and consequently improve delivery of chemotherapeutics to tumor cells. Increased oxygenation would enhance the efficacy of radiation therapy ( Jain, 2005). Evidence supporting this hypothesis have been obtained in various xenograft studies (Ansiaux et al., 2006; Dickson et al., 2007; Dings et al., 2007). However, this model postulates the need to first ‘‘normalize’’ the vessels with an antiangiogenic agent during a critical time window, followed by delivery of the cytotoxic chemotherapy or radiation therapy. Considering that in most clinical trials no attempts have been made to perform such sequential treatments, one cannot rule out the possibility that alternative mechanisms are of greater significance to account for the observed additive or synergistic effects.

3. Genetic Models While studies conducted in xenograft models have provided important insights into the role of the VEGF/VEGFR signaling pathway in tumorigenesis, these studies suffer from some theoretical shortcomings. In most studies, tumor cells are implanted subcutaneously; hence, tumor progression occurs outside their native microenvironment. Also, the human tumor cell lines used for these studies have been maintained in culture for many generations and were adapted to grow in vivo, while

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primary cells directly isolated from patients have more limited growth potential. Furthermore, studies on human tumors are performed in an immunodeficient host. Hence, the contribution of the immune system cannot be adequately assessed. Therefore, it is useful to employ or develop appropriate genetic mouse models to investigate the efficacy of VEGF/ VEGFR blockers in more clinically relevant circumstances.

3.1. Apcþ/min mice APC plays a critical role in regulating the stability of b-catenin, a crucial intracellular mediator of the Wnt signaling pathway (Polakis, 2007). Somatic mutations in Apc are found in the majority of sporadic human colorectal cancers (Ilyas et al., 1999), whereas germline mutations are found in the autosomal-dominant familial adenomatous polyposis (FAP), a condition where individuals develop hundreds to thousands colorectal adenomas and early-onset colorectal carcinomas (Polakis, 2007). Similarly, mutations in murine germline Apc predispose mice to multiple intestinal neoplasia (Min) in an autosomal dominant fashion (Moser et al., 1990). Furthermore, intra-villus adenomas similar to those in Apcþ/min mice have been found in the duodenum of FAP patients (Preston et al., 2008). Thus, Apcþ/min mice represent a valuable model to investigate intestinal tumorigenesis. However, unlike human FAP, the majority of adenomas in Apcþ/min mice occur in the small intestine (Su et al., 1992). More recently, an Apc-mutant rat that more closely resembled human FAP has been described (Amos-Landgraf et al., 2007). Studies in xenograft models have confirmed the efficacy of anti-VEGF treatment in suppressing the growth of variety of cancer cells (Gerber and Ferrara, 2005). However, the effectiveness of anti-VEGF treatment in benign tumors remains to be fully tested. Korsisaari et al. (2007) took advantage of Apcþ/min mice to investigate the role of VEGF in the growth of intestinal benign tumors. In situ hybridization analysis demonstrated upregulation of VEGF in epithelial and stromal cells of adenomas compared to the adjacent normal mucosa.The authors initiated VEGF blocking treatment with Mab G6-31 (5 mg/kg once weekly), which blocks both murine and human VEGF, in Apcþ/min mice when the animals were 13 weeks old. They found that inhibition of VEGF reduced vascular densities and prevented further growth of intestinal adenomas of all size. The mean life span of animals that received Mab G6-31 treatment was extended by nearly 10 weeks when compared to control treated animals. Interestingly, despite the tumor growth arrest, there was no reduction in tumor numbers. To confirm these findings, Apcþ/min mice were crossed with VEGFlox: Villin-Cre mice. Specific deletion of VEGF in intestinal epithelial cells resulted in a reduction in the size of intestinal adenomas, but not in their numbers. Together, these data demonstrate that VEGF signaling is involved

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in tumor progression, but not tumor initiation. It is also noteworthy that pharmacologic inhibition using Mab G6-31 resulted in greater tumor inhibition than genetic deletion of VEGF in epithelial cells, suggesting that stroma-derived VEGF is actively involved in the disease progression (Korsisaari et al., 2007). In a different study, Goodlad et al. (2006) showed that a small molecule VEGFR-2 inhibitor, AZD2171, was able to reduce tumor number in Apcþ/min mice when the treatment was initiated at 6 weeks, whereas later intervention (at 10 weeks) only reduced tumor size. Surprisingly, AZD2171 did not result in any reduction in vascular density of adenomas, perhaps reflecting a complex mechanism of tumor suppression by an agent that inhibits several RTKs in addition to VEGFR-2.

3.2. MEN1 mice The MEN1 gene encodes a tumor suppressor protein, called ‘‘menin’’ (Yang and Hua, 2007). Individuals who possess mutations in MEN1 are predisposed to the development of multiple endocrine neoplasias (MEN) in tissues such as parathyroids, pituitary and pancreatic islets (Lakhani et al., 2007). Treatment for many patients often involves the physical removal of the endocrine tumors (Piecha et al., 2008). Recently, molecular therapies that target VEGF and mTOR have been developed, and encouraging results with bevacizumab and mTOR inhibitors have been observed in phase II clinical trials (Yao, 2007). A mouse model of MEN1 was established via targeted deletion of exons 3 to 8 of MEN1 (Crabtree et al., 2001). While homozygous MEN null mutations result in embryonic lethality, heterozygous mice are viable and display phenotypic changes remarkably similar to those associated with the human disorder. Hence, MEN1 mouse provide an interesting and relevant model for testing molecular therapies in vivo. Since angiogenesis is thought to be important for the growth of a broad variety of tumor types, Korsisaari et al. (2008) tested the efficacy of anti-VEGF treatment in Men1 mice. The tumors in animals that received anti-VEGF treatment were growth arrested, whereas those in control antibody-treated groups grew rapidly resulting in increased serum prolactin level and reduced life span. This finding demonstrates that pituitary tumors are accessible and sensitive to VEGF-blocking antibodies. Consistent with the reduction in tumor sizes, immunochemistry revealed a reduction in vascular density in anti-VEGF–treated pituitary tumors. Moreover, the reduction in vascular density was also observed in the pancreatic tumors within the same animals. This result suggests that antiVEGF treatment is likely to be effective for all types of endocrine tumors within MEN mice. Hence, VEGF neutralized a promising strategy for treating multiple tumor types within the same individual.

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3.3. RIP-Tag model RIP-Tag mice express SV40 T antigen under the control of the rat insulin promoter (Hanahan, 1985). The development of pancreatic islet carcinogenesis in such model (RIP-Tag2) has been well characterized, with hyperplastic and dysplastic islets appearing at 4 to 5 weeks of age. Angiogenic switch occurs in a subset of the hyperplastic and dysplastic islets during the next 5 weeks. A subset of angiogenic islets develops into adenomas during the next few weeks, of which a fraction will become invasive. All five members of VEGF family and VEGFR-1 and VEGFR-2 were found to be expressed in normal islets and during all stages of pancreatic islet tumorigenesis (Christofori et al., 1995; Inoue et al., 2002), in RIP-Tag mice. However, no marked upregulation of VEGF mRNA was observed during tumor progression. Both genetic and pharmacological approaches were subsequently utilized to directly probe the role of VEGF in RIP-Tag tumorigenesis (Casanovas et al., 2005; Inoue et al., 2002). Deletion of VEGF specifically in the b cells of RIP-Tag mice prior to the onset of hyperplasia resulted in reduced angiogenic switching and severely attenuated tumor growth at later stages (Inoue et al., 2002). MMP-9 was found to promote angiogenic switching in RIP-Tag mice and increase the bioavailability of VEGF, potentially accounting for the important role of VEGF in spite of the lack of significant mRNA upregulation (Bergers et al., 2000). Strikingly, no compensatory signaling pathway was able to substitute for VEGF deficiency in this model, despite the fact that all other members of VEGF family and other angiogenic factors such as FGF are expressed (Inoue et al., 2002). These data clearly point to a pivotal role of VEGF in pancreatic islet tumorigenesis, particularly at the early stages. To further investigate the role of the VEGF/VEGFR pathway in RIPTAG mice, Bergers et al. (2003) initially used small-molecule kinase inhibitors. They found that administration of SU5416, which primarily inhibits VEGFR signaling, has marked suppressive effects when tested at early but not at later stages of tumor progression. In contrast, SU6668, which also inhibits PDGFR signaling, was found to have inhibitory effects also on advanced tumors. The conclusion of these studies was that targeting PDGFR-B, which mediates pericyte recruitment, enhances the therapeutic effects resulting from suppression of VEGF signaling. Later, Casanovas et al. (2005) tested neutralizing monoclonal antibodies specific for VEGFR1 and/or VEGFR2 in RIP-Tag mice at 5 weeks of age, before the occurrence of the angiogenic switch, or at 10 weeks of age before the appearance of tumors. Consistent with the findings reported by Inoue et al. (2002), early administration of the anti-VEGFR2 monoclonal antibody DC101 resulted in reduced angiogenic switching and attenuated tumor growth. However, inhibition of VEGFR1 signaling using Mab MF1 had no effect on either angiogenic switching or subsequent tumor growth. These data demonstrate

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that VEGFR2, but not VEGFR1, is critical for angiogenic switching in the RIP-Tag model, even though both receptors are expressed throughout the premalignant and malignant stages. Since previous studies had established the critical role of inflammatory cells in angiogenic switching in this model (Bergers et al., 2000) and the involvement of VEGF/VEGFR1 signaling pathway in macrophages recruitment (Sawano et al., 2001), the authors investigated macrophage infiltration in the tumors. They found that macrophage recruitment was not affected by Mab MF1, suggesting that VEGFR1 signaling is dispensable for such event, at least in the RIP-Tag model. The RIP-Tag model has also been used to test the hypothesis that VEGF blocking agents may induce drug resistance. Casanovas et al. (2005) treated RIP-Tag mice with the VEGFR2 blocking antibody DC101, starting 12 weeks after the tumors had occurred. Administration of Mab DC101 led to a temporary stabilization of the tumors. However, tumor regrowth were observed after 4 weeks of treatment. The authors went on to show that FGF family members are upregulated in the tumors and may compensate for VEGF/VEGFR signaling deficiency. Neutralizing FGF signaling by adenovirus-delivered FGF-soluble receptor appeared to slow down the rate of tumor regrowth after DC101 administration, suggesting that VEGF- and FGF-mediated angiogenic pathways may be sequentially activated and thus they both represent potential therapeutic targets. Finally,the RIP-Tag model has been characterized as being particularly stringent, suggesting that the effects of various inhibitors obtained in this model may be particularly relevant to the design of clinical trials (Bergers et al., 1999). However, several anti-angiogenic agents (endostatin, angiostatin, TNP470, MMP inhibitor) were reported to have significant anti-cancer effects in RIP-Tag mice (Bergers et al., 1999), but none proved effective in cancer patients.

3.4. Humanized VEGF mouse model Preclinical studies in traditional xenografts and in genetic models were be useful in demonstrating the effectiveness of anti-VEGF treatment. However, several questions could not be answered by these models. As already pointed out, some antibodies such as bevacizumab only inhibit human VEGF (Yu et al., 2008), and thus the contribution of host stroma-derived VEGF cannot be determined (Dong et al., 2004; Tejada et al., 2006). Also, the toxicities associated with such treatments cannot be adequately investigated. To address these questions, a humanized mouse model was generated (Gerber et al., 2007). The mouse model that Gerber et al. (2007) engineered is hum-X VEGF, which expresses a humanized VEGF variant, employing gene knock-in technology. Based on prior crystal structure and site-directed mutagenesis studies (Muller et al., 1997, 1998), 10 amino acids (aa) of murine VEGF were replaced with the corresponding human residues, out of a total of 19 aa that are different between murine VEGF164 and human VEGF165.

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The 10 aa are encoded within exons 3, 4, and 5. Importantly, the ‘‘humanized’’ VEGF was similar to human VEGF in terms of biological activities in cultured endothelial cells and interaction with VEGF-blocking antibodies including bevacizumab. Furthermore, since the humanized VEGF gene is under the control of the endogenous murine promoter, the temporal and spatial expressions of the protein are not altered. Employing hum-X VEGF or hum-X VEGF:Rag2 KO mice, Gerber et al. (2007) also compared the pharmacokinetics and pharmacodynamic properties of various VEGF-blocking antibodies and tested their abilities to inhibit the growth of several human tumors. Interestingly, they found that there is no direct correlation between the affinities of the antibodies and their efficacy or potency in suppressing tumor growth in vivo, even though such a correlation was observed in in vitro assays. For example, bevacizumab was as potent and efficacious as Mab Y0317, an affinity-matured variant with 100-fold higher binding affinity for human VEGF. However, these studies revealed a correlation between VEGF binding affinity and toxicity. The most affected organs were kidney and liver. Considering that multiple monoclonal antibodies and other VEGF inhibitors, including small molecules, may induce such toxicities, it is likely that they truly result from suppression of VEGF signaling. For a discussion of the potential mechanisms underlying such toxicities, see Gerber et al. (2007).

4. Conclusions Murine models, both xenografts and genetic models, have generated much information on the biology of VEGF and the therapeutic potential of VEGF/VEGFR inhibitors. The findings obtained in xenografts were largely confirmed and extended in genetic models, and showed that VEGF-dependent angiogenesis frequently represents a rate-limiting step, particularly in early stages of tumor growth or tumor progression. Based on such promising findings, clinical trials were initiated with bevacizumab and other inhibitors of the VEGF pathway, in combination with standard chemotherapy. Results from phase III clinical trials demonstrated that bevacizumab in combination with chemotherapy had clinical benefits in patients with metastatic breast cancers, non–small cell lung cancers, or previously untreated metastatic colorectal cancers when compared with standard chemotherapy alone (Hurwitz et al., 2004; Miller et al., 2007; Sandler et al., 2006). In addition, Sorafenib and sunitinib exhibited clinical efficacy in metastatic renal cell carcinoma to warrant FDA approval and sorafenib is also FDA approved for hepatocellular carcinoma. It is noteworthy that the clinical studies resulting in FDA approval of bevacizumab and other VEGF blockers have been performed in

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combination with cytotoxic chemotherapy in patients with highly advanced malignancies. However, as already mentioned, the most impressive results in preclinical tumor models were obtained when VEGF inhibitors were administrated shortly after tumor inoculation or when the tumor burden was low. These circumstances resemble adjuvant therapy in which late-stage tumors are usually surgically resected, while less advanced neoplasia remains. Several adjuvant clinical trials with bevacizumab are ongoing to test the hypothesis that blocking VEGF may be beneficial in these settings.

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Rubenstein, J. L., Kim, J., Ozawa, T., Zhang, M., Westphal, M., Deen, D. F., and Shuman, M. A. (2000). Anti-VEGF antibody treatment of glioblastoma prolongs survival but results in increased vascular cooption. Neoplasia 2, 306–314. Saishin, Y., Saishin, Y., Takahashi, K., Lima e Silva, R., Hylton, D., Rudge, J. S., Wiegand, S. J., and Campochiaro, P. A. (2003). VEGF-TRAP(R1R2) suppresses choroidal neovascularization and VEGF-induced breakdown of the blood-retinal barrier. J. Cell Physiol. 195, 241–248. Sandler, A., Gray, R., Perry, M. C., Brahmer, J., Schiller, J. H., Dowlati, A., Lilenbaum, R., and Johnson, D. H. (2006). Paclitaxel-carboplatin alone or with bevacizumab for non– small-cell lung cancer. N. Engl. J. Med. 355, 2542–2550. 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. Shibuya, M. (2006). Differential roles of vascular endothelial growth factor receptor-1 and receptor-2 in angiogenesis. J. Biochem. Mol. Biol. 39, 469–478. Shibuya, M., and Claesson-Welsh, L. (2006). Signal transduction by VEGF receptors in regulation of angiogenesis and lymphangiogenesis. Exp. Cell Res. 312, 549–560. Shojaei, F., Wu, X., Malik, A. K., Zhong, C., Baldwin, M. E., Schanz, S., Fuh, G., Gerber, H. P., and Ferrara, N. (2007). Tumor refractoriness to anti-VEGF treatment is mediated by CD11bþGr1þ myeloid cells. Nat. Biotechnol. 25, 911–920. Soffer, S. Z., Moore, J. T., Kim, E., Huang, J., Yokoi, A., Manley, C., O’Toole, K., Stolar, C., Middlesworth, W., Yamashiro, D. J., and Kandel, J. J. (2001). Combination antiangiogenic therapy: increased efficacy in a murine model of Wilms tumor. J. Pediatr. Surg. 36, 1177–1181. Su, L. K., Kinzler, K. W., Vogelstein, B., Preisinger, A. C., Moser, A. R., Luongo, C., Gould, K. A., and Dove, W. F. (1992). Multiple intestinal neoplasia caused by a mutation in the murine homolog of the APC gene. Science 256, 668–670. Tejada, M. L., Yu, L., Dong, J., Jung, K., Meng, G., Peale, F. V., Frantz, G. D., Hall, L., Liang, X., Gerber, H. P., and Ferrara, N. (2006). Tumor-driven paracrine plateletderived growth factor receptor alpha signaling is a key determinant of stromal cell recruitment in a model of human lung carcinoma. Clin. Cancer Res. 12, 2676–2688. Warren, R. S., Yuan, H., Matli, M. R., Gillett, N. A., and Ferrara, N. (1995). Regulation by vascular endothelial growth factor of human colon cancer tumorigenesis in a mouse model of experimental liver metastasis. J. Clin. Invest. 95, 1789–1797. Wilhelm, S. M., Carter, C., Tang, L., Wilkie, D., McNabola, A., Rong, H., Chen, C., Zhang, X., Vincent, P., McHugh, M., Cao, Y., Shujath, J., et al. (2004). BAY 43-9006 exhibits broad spectrum oral antitumor activity and targets the RAF/MEK/ERK pathway and receptor tyrosine kinases involved in tumor progression and angiogenesis. Cancer Res. 64, 7099–7109. Yang, Y., and Hua, X. (2007). In search of tumor suppressing functions of menin. Mol. Cell Endocrinol. 265–266, 34–41. Yao, J. C. (2007). Neuroendocrine tumors. Molecular targeted therapy for carcinoid and islet-cell carcinoma. Best Pract. Res. Clin. Endocrinol. Metab. 21, 163–172. Yu, L., Wu, X., Cheng, Z., Lee, C. V., Lecouter, J., Campa, C., Fuh, G., Lowman, H., and Ferrara, N. (2008). Interaction between Bevacizumab and Murine VEGF-A: A Reassessment. Invest. Ophthalmol. Vis. Sci. 49, 522–527. Zhang, J., Jia, Z., Li, Q., Wang, L., Rashid, A., Zhu, Z., Evans, D. B., Vauthey, J. N., Xie, K., and Yao, J. C. (2007). Elevated expression of vascular endothelial growth factor correlates with increased angiogenesis and decreased progression-free survival among patients with low-grade neuroendocrine tumors. Cancer 109, 1478–1486.

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Molecular Imaging of Tumor Vasculature Weibo Cai,*,† Sanjiv S. Gambhir,* and Xiaoyuan Chen* Contents 1. Introduction 2. Structural/Functional Imaging of Tumor Vasculature 2.1. Structural imaging of tumor vasculature 2.2. Functional imaging of tumor vasculature with MRI 2.3. Functional imaging of tumor vasculature with other modalities 3. Molecular Imaging 4. Imaging Integrin avb3 in Tumor Vasculature 4.1. Single-modality imaging 4.2. Dual-modality imaging 5. Non-Radionuclide–Based Imaging of VEGFR in Tumor Vasculature 5.1. Targeted ultrasound 5.2. Optical imaging 6. Radionuclide-Based Imaging of VEGFR in Tumor Vasculature 6.1. SPECT imaging 6.2. PET imaging 7. Experimental Section 7.1. Production of VEGF121 protein 7.2. DOTA conjugation 7.3. Cell-binding assay of VEGF121 and DOTA-VEGF121 7.4. Cell lines and animal models 7.5. 64Cu-labeling of DOTA-VEGF121 7.6. Micro-PET imaging 7.7. Biodistribution studies 7.8. Immunofluorescence staining 8. Summary Acknowledgments References

* {

142 143 145 146 146 148 149 150 152 153 154 156 156 156 158 161 162 162 163 163 164 164 166 167 167 168 168

Stanford University School of Medicine, Stanford, California Departments of Radiology and Medical Physics, University of Wisconsin, Madison, Wisconsin

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03007-3

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2008 Elsevier Inc. All rights reserved.

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Abstract Cancer, with more than 10 million new cases a year worldwide, is the third leading cause of death in developed countries. One critical requirement during cancer progression is angiogenesis, the formation of new blood vessels. Structural and functional imaging of tumor vasculature has been studied using various imaging modalities such as magnetic resonance imaging (MRI), computed tomography (CT), and ultrasound. Molecular imaging, a key component of the 21st-century cancer-patient management strategy, takes advantage of these traditional imaging techniques and introduces molecular probes to determine the expression of indicative molecular markers at different stages of cancer development. In this chapter, we will focus on two tumor vasculature– related targets: integrin avb3 and vascular endothelial growth factor receptor (VEGFR). For imaging of integrin avb3 on the tumor vasculature, only nanoparticle-based probes will be discussed. VEGFR imaging will be discussed in depth, and we will give a detailed example of positron emission tomography (PET) imaging of VEGFR expression using radio-labeled VEGF121 protein. Future clinical translation will be critical for maximum patient benefit from these agents. To achieve this goal, multidisciplinary approaches and cooperative efforts from many individuals, institutions, industries, and organizations are needed to quickly translate multimodality tumor vasculature imaging into multiple facets of cancer patient management.

1. Introduction Cancer is the third leading cause of death (after heart disease and stroke) in developed countries and the second leading cause of death (after heart disease) in the United States (http://www.cdc.gov). Studies have shown that there were 10 million new cases, 6 million deaths, and 22 million people living with cancer in the year 2000 (Parkin, 2001). The most common cancer types in terms of new cases were lung (1.2 million), breast (1.05 million), and colorectal (0.95 million) cancer. These numbers represent an increase of about 22% in incidence and mortality from that of the year 1990 (Parkin et al., 1999; Pisani et al., 1999). The most common causes of cancer mortality are lung (17.8%), stomach (10.4%), and liver (8.8%) cancer. It is projected that the number of new cases for all cancers will be 12.34 million and 15.35 million in the year 2010 and 2020, respectively (Parkin, 2001). One of the key requirements during cancer progression is angiogenesis, the formation of new blood vessels, without which the tumor may not grow beyond a few millimeters in diameter (Bergers and Benjamin, 2003; Folkman, 1995). Tumor angiogenesis differs significantly from physiological angiogenesis, including the aberrant

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vascular structure, altered endothelial cell–pericyte interactions, abnormal blood flow, increased permeability, enlargement of the vessel diameter, basement membrane degradation, thin endothelial cell lining, increased number of endothelial cells, decreased number of pericytes, and delayed maturation (Bergers and Benjamin, 2003; Hanahan and Folkman, 1996). Cancer cells can spread throughout the body by metastasis (Bogenrieder and Herlyn, 2003; Friedl and Wolf, 2003). The leaky tumor vasculature can allow for intravasation of tumor cells, which can then exit the bloodstream (extravasatin) and undergo expansive growth within the parenchyma of other organ(s). The fact that tumor progression is dependent on angiogenesis has inspired scientists to search for antiangiogenic molecules and design antiangiogenic strategies for cancer treatment and prevention of cancer recurrence/metastasis (Cai and Chen, 2006; Folkman, 2007; Kerbel and Folkman, 2002). These antiangiogenic agents typically lead to a stop of tumor progression rather than to tumor shrinkage; thus, the traditional approach of measuring the therapeutic response by a reduction of tumor size is no longer applicable. Imaging techniques can be used as biomarkers to provide an indicator of effectiveness at the functional or molecular level. In this chapter, we will first briefly describe the structural/functional imaging techniques commonly employed for imaging the tumor vasculature. We will then focus on two of the most well-studied tumor vasculature–related molecular targets: integrin avb3 and vascular endothelial growth factor receptor (VEGFR). Lastly, we will give a detailed example of positron emission tomography (PET) imaging of tumor vasculature VEGFR expression using radio-labeled vascular-endothelial-growth-factor (VEGF) protein.

2. Structural/Functional Imaging of Tumor Vasculature Because antiangiogenic therapies are designed to affect the abnormal blood vessels within the tumor, morphological changes of the tumor vasculature and changes in hemodynamic parameters such as blood flow, blood volume, or vessel permeability may be promising biomarkers for evaluating therapeutic effects. Structural and functional imaging of blood vessels has been studied using various imaging modalities (Table 7.1) (McDonald and Choyke, 2003). In most cases, they were performed to evaluate the vasculature of certain organs and/or diseases besides cancer. However, these techniques can also be applied to structural/functional imaging of the vasculature in tumors.

Table 7.1 Techniques employed to investigate structural and functional characteristics of tumor vasculature noninvasively or ex vivo Structural/ Functional

In vivo/ Ex vivo

References

Vascular cast Immunohistochemical staining Fluorescent transgenic mice Intravital labeling CT angiography MR angiography Doppler ultrasonography Intravital microscopy Dynamic contrastenhanced MRI

Structural Structural

Ex vivo Ex vivo

Konerding et al., 1995; Less et al., 1991 Miettinen et al., 1994; Schlingemann et al., 1991

Structural

Both

Motoike et al., 2000; Schlaeger et al., 1997

Both Structural Structural Both

Both In vivo In vivo In vivo

Both Functional

In vivo In vivo

Arterial spin labeling Blood oxygen level– dependent MRI Dynamic contrastenhanced CT Positron emission tomography Fluorescence-mediated tomography

Functional Functional

In vivo In vivo

Hashizume et al., 2000; Trotter et al., 1990 McDonald and Choyke, 2003 Fink et al., 2003; Kobayashi et al., 2001 Fleischer et al., 1999; Forsberg et al., 2004; Foster et al., 2000; Stride and Saffari, 2003 Tozer et al., 2005 Choyke et al., 2003; Demsar et al., 1997; Haroon et al., 2007; Lankester et al., 2007; O’Connor et al., 2007; Orth et al., 2007; Preda et al., 2004; Roberts et al., 2006 Liu and Brown, 2007; Williams, 2006; Wolf and Detre, 2007 Baudelet et al., 2006; Hsu et al., 2004

Functional

In vivo

Bisdas et al., 2007; Haider et al., 2005

Functional

In vivo

Functional

In vivo

Dimitrakopoulou-Strauss et al., 2001; Mullani et al., 2000; Ter-Pogossian and Herscovitch, 1985 Montet et al., 2007; Montet et al., 2005

Techniques

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2.1. Structural imaging of tumor vasculature Several techniques can be used for tumor vasculature imaging ex vivo. Vascular casts, after intravascular injection of colored gelatin, latex, or plastic casting material followed by tissue clearing or corrosion (Konerding et al., 1995; Less et al., 1991), have been prepared for examining the 3D architecture of blood vessels and vascular networks. This approach enables quantification of the vessel dimensions, intervascular distances, branching order, and luminal surface features that cannot be readily assessed in tissue sections. Both immunohistochemical staining of endothelial cell markers such as CD31, CD34, and von Willebrand factor (Miettinen et al., 1994; Schlingemann et al., 1991), and labeling the endothelial cells by fluorescent reporters expressed in transgenic mice (Motoike et al., 2000; Schlaeger et al., 1997) can allow for visualization of the tumor blood vessels. During such histological examination, the appearance of blood vessels in the sections is greatly influenced by section thickness. Only short segments of the vessels are visible in thinner sections and the vascular network has to be studied in thicker sections. Intravital labeling with an agent that can reach vascular targets only through the circulation can help identify the functional blood vessels in the tumor (Hashizume et al., 2000; Trotter et al., 1990). Tumor macrovasculature imaging can be performed clinically by various noninvasive imaging modalities such as magnetic resonance imaging (MRI), computed tomography (CT), and ultrasound (McDonald and Choyke, 2003). However, visualization of the microvasculature is very challenging. X-ray, CT, and MR angiography, even after administration of intravascular contrast agents, do not have sufficient resolution to resolve the microvasculature. Even under optimized conditions, only vessels down to diameters of 100 to 200 mm can be imaged with clinical MR scanners (Fink et al., 2003; Kobayashi et al., 2001). The resolution of color Doppler ultrasonography depends on the imaging frequency (usually limited to
200 mm), and images are confined to superficial parts of the body (Fleischer et al., 1999). Scanners dedicated to small animal imaging studies, such as micro-CT, have better spatial resolution in preclinical models but with poor temporal resolution and substantial radiation exposure ( Jiang et al., 2000). Intravital microscopy of tumors growing in animal models with window chambers can directly enable the visualization of tumor angiogenesis and vascular response to treatment, in terms of both the morphology of the vascular networks and the function of individual vessels (Tozer et al., 2005). This technique allows for repeated measurements of the same tumor with very high resolution (down to the submicrometer level). Multi-photon fluorescence microscopy techniques have also been applied to these model systems to obtain 3D images of the tumor vasculature (Tozer et al., 2005).

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2.2. Functional imaging of tumor vasculature with MRI Several different MRI contrast mechanisms can be used for evaluating the functional properties of tumor vasculature, such as dynamic contrastenhanced MRI (DCE-MRI), arterial spin labeling (ASL), and blood oxygen level–dependent (BOLD) MRI. DCE-MRI works by tracking the pharmacokinetics of injected contrast agents as they pass through the tumor vasculature, which represents a complex summation of vascular permeability, blood flow, vascular surface area, and interstitial pressure (Choyke et al., 2003; O’Connor et al., 2007). In T2*-weighted DCEMRI, the regional blood flow, regional distribution volume, and the mean transit time of the contrast agent can be estimated using a deconvolution method (Haroon et al., 2007; Lankester et al., 2007). Alternatively, concentration-time courses derived from DCE-MRI can be analyzed by pharmacokinetic modeling with two or more compartment models (Orth et al., 2007; Roberts et al., 2006). Some macromolecular MR contrast agents, such as gadolinium-labeled albumin (Demsar et al., 1997; Preda et al., 2004), have long circulation half-lives that can be used to more accurately measure the blood volume and estimate the vascular permeability in tumors. In ASL, water molecules can be labeled for MRI by inverting the nuclear spin of their hydrogen atoms with a radiofrequency pulse directed at the arterial blood before it enters the region of interest (ROI) (Liu and Brown, 2007; Williams, 2006). An absolute value of blood flow is determined by the change in the MR signal as the labeled water in the arterial bloodstream arrives in the ROI (Wolf and Detre, 2007). Because the spin label is very short-lived, the signal-to-noise ratio of ASL MRI is relatively low with limited sensitivity and spatial resolution, especially when the rate of blood flow is low. BOLD-MRI can detect the changes in oxygen saturation of the blood (Baudelet et al., 2006; Hsu et al., 2004). By measuring signal changes in response to hypercapnia and hyperoxia, the vascular maturity can be detected since only mature vessels react to hypercapnia. The BOLD effect can be enhanced by increasing the amount of oxygen in the breathed air.

2.3. Functional imaging of tumor vasculature with other modalities DCE-CT is analogous to DCE-MRI (Bisdas et al., 2007; Haider et al., 2005). Contemporary CT scanners can permit rapid repeated image acquisitions at low radiation doses. The linear relationship between signal attenuation and contrast agent concentration in CT is advantageous over DCE-MRI. However, repeated exposure to ionizing radiation and the potential nephrotoxicity of CT contrast agents are two major concerns.

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To minimize these risks, DCE-CT studies are typically quite brief using only a low dose of contrast agent. PET imaging has also been used to assess blood flow and blood volume in tumors. Blood volume within tumors can be determined by inhaling carbon monoxide labeled with either 11C or 15O (Ter-Pogossian and Herscovitch, 1985). When inhaled, trace amounts of labeled carbon monoxide irreversibly bind to red blood cells and distribute in accordance with the vascular volume. Most PET perfusion measurements are performed with 15O-H2O (Dimitrakopoulou-Strauss et al., 2001; Mullani et al., 2000), using either static or dynamic PET imaging. The first pass of 15OH2O can be used to calculate the blood flow within tumors because water is completely diffusible and redistributes quickly. The washout of the activity curve for a tumor is directly proportional to the blood flow. The major limitation of these techniques is that the short decay half-lives of the radioisotopes (20 min and 2 min for 11C and 15O, respectively) make such studies very difficult, and it is only possible when there is an on-site cyclotron available. Ultrasound is well established for measuring blood flow or, more precisely, blood velocity using the Doppler principle (Fleischer et al., 1999; Forsberg et al., 2004). Innovations in ultrasound imaging have made it possible to measure blood flow in microvasculature smaller than 100 mm. Microbubble contrast agents, often 1 to 10 mm in diameter, can reveal the blood flow down to the microcirculation level by increasing the signal from small vessels (Stride and Saffari, 2003). At higher frequencies, such as 50 MHz, Doppler processing can allow the direct assessment of flow dynamics in real time in arterioles as small as 15 mm (Foster et al., 2000). Fluorescence-mediated tomography (FMT) has been applied to measure angiogenesis by using fluorescent dye–containing nanoparticles (Montet et al., 2005). Results from such FMT measurements were shown to correlate well with CD31 microvascular–density measurements and MRI of vascular volume fraction. Quantification of the vascular volume fraction with FMT using long-circulating, fluorescent blood-pool agents has also been reported (Montet et al., 2007). The major limitation of such optical imaging techniques is that it is very difficult to image tumor tissue deeper than 1 cm because of significant tissue absorption and scattering of light (Massoud and Gambhir, 2003). Even though structural/functional imaging of the tumor vasculature can reveal potentially useful information before, during, and after therapeutic intervention, they do not convey enough knowledge about the biological changes upon therapy at the molecular level that may occur long before any structural/functional changes can be detected. Molecular imaging, a recently emerged field, can be a powerful tool in cancer diagnosis, cancer treatment, and anticancer drug development (Massoud and Gambhir, 2003).

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3. Molecular Imaging The field of molecular imaging, recently defined by the Society of Nuclear Medicine (SNM) to be ‘‘the visualization, characterization and measurement of biological processes at the molecular and cellular levels in humans and other living systems’’ (Mankoff, 2007), has flourished over the last decade. In general, the predominant molecular imaging modalities include molecular MRI (mMRI), magnetic resonance spectroscopy (MRS), optical bioluminescence, optical fluorescence, targeted ultrasound, single-photon-emission computed tomography (SPECT), and PET (Massoud and Gambhir, 2003). Many hybrid systems that combine two or more modalities are also commercially available and others are under active development (Beyer et al., 2000; Catana et al., 2006; Even-Sapir et al., 2003). Continued development and wider availability of scanners dedicated to small animal imaging studies (Fig. 7.1), which can provide a similar in vivo imaging capability in mice, primates, and humans, can enable smooth transfer of knowledge and molecular measurements between species, thereby facilitating clinical translation. Molecular imaging (except MRS based on endogenous contrast) takes advantage of the traditional diagnostic imaging techniques and introduces molecular probes to determine the expression of indicative molecular

micro MRI

micro SPECT

Ultrasound

Optical

micro PET

Figure 7.1 The molecular imaging instruments dedicated for small-animal imaging and representative images.

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markers at different stages of diseases. It can provide a whole-body readout in an intact system, which is much more relevant and reliable than in vitro/ ex vivo studies; help decrease the workload and speed up the drug development process; provide more statistically accurate results since longitudinal studies can be performed in the same animal, which serves as its own control; aid in lesion detection in cancer patients and patient stratification; and help in individualized anticancer treatment monitoring and dose optimization (Cai et al., 2006b; Massoud and Gambhir, 2003). Noninvasive detection of molecular markers can allow for much earlier diagnosis, earlier treatment, better prognosis, and improved staging and management, which can eventually lead to personalized medicine. Molecules regulating angiogenesis include, but are not limited to, growth factor receptors, tyrosine kinase receptors, G-protein–coupled receptors for angiogenesis-modulating proteins, integrins, and matrix metalloproteinases (MMPs) (Ferrara, 2002). During tumor angiogenesis, these molecules interact closely with each other. For example, integrin avb3 can recruit and activate MMP-2 which degrades the components of basement membrane and interstitial matrix to facilitate tumor progression (Brooks et al., 1996). It can also regulate the production of VEGF in certain tumor cells (De et al., 2005). The VEGF/VEGFR signaling pathway plays a pivotal role in both normal and tumor vasculature development (Ferrara, 2002, 2005). Overexpression of VEGF and/or VEGFRs has been implicated as a poor prognostic marker in many clinical studies (Ferrara, 2004). In this chapter, we will summarize the progress to date on molecular imaging of two key tumor vasculature–related targets: integrin avb3 and VEGFR.

4. Imaging Integrin avb3 in Tumor Vasculature Integrins are a family of cell adhesion molecules playing key roles during tumor angiogenesis and metastasis (Hood and Cheresh, 2002; Hynes, 2002). Integrins expressed on endothelial cells modulate cell migration and survival during tumor angiogenesis while integrins expressed on carcinoma cells potentiate metastasis by facilitating invasion and movement across blood vessels. The avb3 integrin, which binds to arginine-glycine-aspartic acid (RGD)–containing components of the extracellular matrix, is significantly upregulated on tumor vasculature but not on quiescent endothelium (Hood and Cheresh, 2002; Xiong et al., 2002). Currently, most reports on integrin avb3 imaging use RGD peptides as the targeting ligand, and many comprehensive reviews have been published (Cai et al., 2005, 2006b; Liu, 2006). In the majority of cases, the RGD peptide-based probes bind to integrin avb3 expressed on both tumor vasculature and tumor cells. Therefore, these probes are not truly targeting/imaging the tumor vasculature alone.

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Gas

Liposome

Perfluorocarbon

Microbubble

Adenovirus

CdSe ZnS

Carbon nanotube

Quantum dot

Iron oxide

Figure 7.2 Many nanoparticles have been tested for imaging integrin avb3 on the tumor vasculature.

A number of nanoparticle-based probes have been reported for integrin avb3 imaging (Fig. 7.2). Due to the large overall size (from a few nanometers to a few micrometers in diameter), these probes either do not extravasate at all or do not extravasate well from the vasculature. Therefore, they almost exclusively target integrin avb3 on the tumor vasculature. Comparing with the vast number of literature reports on integrin avb3 imaging using RGD peptide or protein-based probes, nanoparticle-based imaging of integrin avb3 is relatively rare and has been recently reviewed (Cai and Chen, 2007b, 2008a). Here we will briefly summarize the current state of the art on imaging tumor-vascular integrin avb3 using nanoparticlebased agents.

4.1. Single-modality imaging Each molecular imaging modality has been employed for imaging tumor vascular integrin avb3 expression (Cai and Chen, 2008a; Cai et al., 2005, 2006b). For mMRI, Gd3þ-containing paramagnetic liposomes (Sipkins et al., 1998), peptidomimetic–integrin avb3 antagonist–conjugated magnetic nanoparticles (Schmieder et al., 2005; Winter et al., 2003), and avb3 integrin–targeted ultrasmall superparamagnetic iron oxide (USPIO) nanoparticles (Zhang et al., 2007) have been investigated. Due to the poor sensitivity of MRI, which can only be partially compensated by working at higher magnetic fields (4.7 to 14 T) and acquiring data for much longer time periods, mMRI is so far mostly a proof-of-principle study, and whether it can significantly improve cancer patient management remains unclear.

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For optical imaging of tumor-vascular integrin avb3, RGD peptide– conjugated quantum dot (QD) has been reported (Cai et al., 2006c). Spectral imaging techniques, where the fluorescence signals of different fluorophores can be separated based on their emission spectra (Levenson, 2004; Mansfield et al., 2005), were employed to better interpret the imaging results. QD-based optical imaging may play an important role in imageguided surgery upon further development of smaller, less-toxic, multifunctional QDs, and further improvement of the conjugation strategy (Cai et al., 2007d; Li et al., 2007). Ultrasound imaging of integrin avb3 during tumor angiogenesis has been reported exclusively with microbubble-based agents (Ellegala et al., 2003; Hughes et al., 2006). Very few reports are available for SPECT imaging of integrin avb3 using a nanoparticle-based tracer. In one study, 111In-labeled perfluorocarbon nanoparticles were tested for integrin avb3 imaging in New Zealand white rabbits implanted with Vx-2 lung carcinoma tumor (Hu et al., 2007). With either bioluminescence imaging (BLI) or PET imaging, depending on the gene incorporated inside the virus, we have noninvasively monitored the re-direction of adenovirus tropism in tumor-bearing mice through chemical modification of the fiber knobs with PEGylated RGD peptides (Niu et al., 2007; Xiong et al., 2006). The biodistribution of radio-labeled, single-walled carbon nanotubes (SWNTs) in mice has been studied using PET, biodistribution, and ex vivo Raman spectroscopy (Liu et al., 2007). Efficient targeting of integrin avb3– positive U87MG tumors in mice (
15% injected dose per gram of tissue) (% ID/g), among the highest of any nanoparticles ever reported, was achieved with RGD peptide–conjugated SWNTs. The unique Raman signatures of SWNTs enabled direct measurement of SWNTs in various mouse tissue, which confirmed the radionuclide-based results. Additionally, tumor accumulation of this SWNT-based construct in living mice was also noninvasively detected using a prototype Raman-imaging system (Zavaleta et al., 2007). Radio-labeled nanoparticles represent a new class of probes that has enormous potential for clinical applications. The radio-label enables quantitative measurement of tumor-targeting efficacy and pharmacokinetics, provided that the radio-label on the nanoparticle is stable under physiological conditions. However, dissociation of the radionuclide (typically metal) from the chelator, and/or the radionuclide-containing polymer coating from the nanoparticle, may occur, which can cause significant differences between the nanoparticle distribution and radionuclide distribution. In several other studies, measuring the radio-label only has led to the conclusion that these radio-labeled SWNTs clear mainly through the renal pathway in mice (McDevitt et al., 2007; Singh et al., 2006), which is very unlikely, unless severe kidney damage has occurred. Direct measurement of the nanoparticle in various tissues using its intrinsic signal (e.g., Raman

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for SWNT), as well as rigorous validation of the stability of the radio-label on the nanoparticle, should always be carried out to obtain reliable results.

4.2. Dual-modality imaging Among all the molecular imaging modalities, no single modality is perfect and sufficient to obtain all the necessary information for any questions of interest. For example, it is difficult to accurately quantify fluorescence signal in living subjects based on fluorescence imaging alone, particularly in deep tissues; MRI has high spatial resolution and exquisite soft tissue contrast, yet it suffers from very low sensitivity; and radionuclide-based imaging techniques are very sensitive but they have relatively poor spatial resolution. Combining multiple molecular imaging modalities can offer synergistic advantages over any single modality alone. Nanoparticles are ideal platforms for dual- and multi-modality imaging and potentially therapeutic applications (Cai and Chen, 2007b). The most desirable property of a nanoparticle is multifunctionality in that multiple targeting ligands, imaging labels, and therapeutic drugs can be attached to each nanoparticle, which can provide enormous sensitivity, throughput, and flexibility. We recently developed a QD-based probe for optical and PET imaging of integrin avb3, in which QD surface modification with RGD peptides allows for integrin avb3 targeting and DOTA (1,4,7,10-tetraazacyclododecanetetraacetic acid) conjugation enables PET imaging after 64Cu-labeling (Cai et al., 2007b). We quantitatively evaluated the tumor-targeting efficacy of this dual-modality probe with PET imaging, which was not possible based on optical imaging alone (Cai et al., 2006c). This PET/optical dualmodality probe can confer sufficient tumor contrast at much lower concentration than that required for in vivo optical imaging, thus significantly reducing the potential toxicity and facilitating future clinical translation/biomedical applications of QD-based agents. In another report, MR-detectable and fluorescent liposomes carrying RGD peptides have been evaluated for in vivo tumor imaging (Mulder et al., 2005). These two studies demonstrated the feasibility of dual-modality imaging in vivo but the two modalities were not equally effective. One imaging modality was mainly used for the ex vivo validation of the in vivo results obtained from the other modality. Although only single-modality noninvasive imaging was achieved with these dual-modality probes, which does not take full advantage of the nanoparticle-based approach, the capability of detecting the probe with another imaging modality did provide a convenient/reliable way for ex vivo probe detection after harvesting the targeted tissue, which greatly facilitated the validation of such dual-modality probes than the conventional single-modality probe. PET/MR, acquired in a single measurement, has the potential to become the imaging modality of choice for various clinical applications in

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neurological studies, certain types of cancer, stroke, and the emerging field of stem cell therapy. The future of PET/MR scanners will greatly benefit from the use of dual-modality PET/MR molecular imaging agents. We recently developed an iron oxide (IO)–based probe for PET/MR imaging of tumor integrin avb3 expression (Lee et al., 2008). Poly(aspartic acid)– coated IO nanoparticles (PASP-IO) were synthesized and the surface amino groups were coupled to cyclic RGD peptides for integrin avb3 targeting and DOTA chelators for PET imaging (after labeling with 64Cu), respectively. Both micro-PET and T2-weighted MR imaging showed integrin-specific delivery of RGD-PASP-IO nanoparticles to the U87MG human glioblastoma tumor, also confirmed by ex vivo histology. This study is the first example of in vivo dual-modality PET/MR imaging using a single agent. Adding a fluorescent label to this conjugate that forms a multimodality PET/MR/optical probe may provide even more information regarding the molecular mechanisms of cancer. One major component of molecular medicine is nanomedicine, where the ultimate goal is that multifunctional nanoparticles containing both therapeutic components and multimodality-imaging labels can allow for efficient, specific in vivo delivery of drugs as well as accurate, quantitative assessment of the therapeutic efficacy noninvasively over time (Cai and Chen, 2007b). Most of the integrin avb3-targeted nanoparticles to date are far from optimal for clinical applications. Much research effort will be needed in the future before such nanoparticle-based tumor angiogenesis– targeted agents enter clinical trials. A variety of factors need to be optimized in parallel, among which are biocompatibility, in vivo kinetics, tumortargeting efficacy, evasion from the reticuloendothelial system (RES), acute/chronic toxicity, and cost effectiveness.

5. Non-Radionuclide–Based Imaging of VEGFR in Tumor Vasculature The VEGF/VEGFR signaling pathway is a major moderator of both normal vasculature development and many disease processes (Ferrara, 2002, 2005). The VEGF family is composed of seven members with a common VEGF homology domain: VEGF-A, VEGF-B, VEGF-C, VEGF-D, VEGF-E, VEGF-F, and placenta growth factor (Cai and Chen, 2007a; Ferrara, 2004). VEGF-A is a homodimeric, disulfide-bound glycoprotein existing in several isoforms with different numbers of amino acid residues, e.g. VEGF121 and VEGF165, which also differ in their biological properties including the ability to bind to cell surface heparin sulfate proteoglycans (Ferrara, 2004). The angiogenic actions of VEGF are mainly mediated via two endothelium-specific receptor tyrosine kinases, VEGFR-1 (Flt-1/FLT-1)

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and VEGFR-2 (Flk-1/KDR) (Hicklin and Ellis, 2005). VEGFR-1 is critical for physiologic and developmental angiogenesis whose function varies with the stages of development, states of physiologic/pathologic conditions, and cell types in which it is expressed (Ferrara, 2004, 2005). VEGFR-2 is the major mediator of the mitogenic, angiogenic, and permeability-enhancing effects of VEGF. Many clinical studies have shown that overexpression of VEGF/ VEGFR implied a poor prognosis (Ferrara, 2004). Thus, VEGFR-targeted molecular imaging can serve as a new paradigm for assessing the efficacy of antiangiogenic therapeutics, improving cancer patient management, and elucidating the role and modulation of VEGF/VEGFR signaling during cancer development/intervention. Direct measurement of VEGFR expression with mMRI has not been achieved. However, targeted ultrasound, optical, SPECT, and PET imaging of VEGFR has been reported over the past several years.

5.1. Targeted ultrasound Ultrasonography is the most commonly used clinical imaging modality because of its safety, low cost, ease of use, and wide availability (Bloch et al., 2004). Almost all contrast-enhanced ultrasound imaging uses microbubbles, which can only target the tumor vasculature since they are too large to extravasate (Korpanty et al., 2005). In a mouse model of pancreatic adenocarcinoma, targeted microbubbles were used to image and quantify the vascular effects of anticancer therapies in both subcutaneous and orthotopic pancreatic tumors (Korpanty et al., 2007). Tumor-bearing mice were treated with an anti-VEGF monoclonal antibody and/or gemcitabine (a chemotherapy drug). The localization of antibody-conjugated microbubbles to VEGFR-2 or VEGF-activated blood vessels (the VEGF-VEGFR complex) was monitored by contrast-enhanced ultrasound (Fig. 7.3A). Significant signal enhancement of the tumor vasculature was observed when compared with untargeted or control IgG–conjugated microbubbles. The video intensity from targeted microbubbles also correlated with the expression level of the target (either VEGFR-2 or the VEGF-VEGFR complex), as well as with the microvessel density in tumors under therapy. We have also imaged VEGFR-2 expression in murine tumor models using anti-VEGFR-2-monoclonal-antibody–conjugated microbubbles (Willmann et al.). Targeted microbubbles gave significantly higher average video intensity than control microbubbles in both tumor models tested. The video intensity was significantly lower when blocked by anti–VEGFR-2 antibodies, thus demonstrating target specificity in vivo. These two studies demonstrated that targeted microbubbles can be a novel and attractive tool for noninvasive imaging of tumor angiogenesis and monitoring of vascular effects after therapy. Further development of targeted ultrasound imaging will involve the expansion of targeted disease

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Figure 7.3 Nonradionuclide-based imaging of VEGFR expression. (A) Contrastenhanced ultrasound images obtained withVEGFR-2 targeted microbubbles in control and treated tumor-bearing animals. The tumor video intensity of mice under anti-VEGF treatment is significantly lower than that of the control animals. Arrows indicate the periphery of the tumor and‘‘Ctr’’denotes the center of the tumor. (B) Bioluminescence (BLI) (after injection of D-luciferin), fluorescence (after injection of Cy5.5-VEGF), and merged images of a 4T1 tumor-bearing mouse. (Adapted from Backer, M. V., Levashova, Z., Patel, V., Jehning, B. T., Claffey, K., Blankenberg, F. G., and Backer, J. M. (2007). Molecular imaging of VEGF receptors in angiogenic vasculature with single-chain VEGF-based probes. Nat. Med. 13, 504^509; and Korpanty, G., Carbon, J. G., Grayburn, P. A., Fleming, J. B., and Brekken, R. A. (2007). Monitoring response to anticancer therapy by targeting microbubbles to tumor vasculature. Clin. Cancer Res.13, 323^330.)

states, improvements in technology for ligand attachment to microbubbles, better characterization of the acoustic behavior of targeted contrast agents, and development of more sensitive/accurate imaging methods such as second harmonic strategies. Acoustic destruction of ‘‘payload-bearing’’ microbubbles has been used to deliver drugs or to augment gene transfection (Shohet et al., 2000). Therefore, tumor vasculature–targeted microbubbles may also have future applications in site-specific cancer (gene) therapy.

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5.2. Optical imaging Optical imaging is a relatively low-cost method suitable primarily for small animal studies. In fluorescence imaging, excitation light illuminates the subject and the emission light is collected at a shifted wavelength (Massoud and Gambhir, 2003). Fluorescence imaging has many disadvantages in that it is not quantitative, the image information is surface-weighted due to tissue absorption, and significant background signal is observed due to tissue autofluorescence (Weissleder and Mahmood, 2001). Human VEGF has been conjugated to a fluorescent dye Cy5.5 (maximum emission 696 nm) and tested for in vivo imaging (Backer et al., 2005, 2006). Although tumor contrast was observed after administration of the probe, no information was reported about the whole-body distribution of Cy5.5-VEGF in these studies. Interestingly, the BLI signal of the luciferase-transfected tumor cells does not overlap well with the fluorescence signal of Cy5.5-VEGF in some cases (Fig. 7.3B) (Backer et al., 2007). Studies have shown that cyanine dyes can nonspecifically accumulate in the tumor even without conjugation to a targeting ligand (Licha et al., 2000; Wu et al., 2006), which indicates that extra care needs to be taken when interpreting the experimental findings of fluorescent dye–based imaging. Whenever feasible, blocking experiment should always be carried out to confirm the receptor specificity of the probe in vivo. The near-infrared (700 to 900 nm) fluorescence (NIRF) imaging approach, in which the absorbance spectra for all biomolecules reach minima, thus revealing a clear window for in vivo optical imaging (Frangioni, 2003), should provide better opportunities for visualizing tumor angiogenesis in the future in both small animal tumor models and the clinical setting.

6. Radionuclide-Based Imaging of VEGFR in Tumor Vasculature Radionuclide-based imaging has been routinely used in the clinic over the last decades (Rohren et al., 2004; Spencer et al., 1995). Because of the wider availability of gamma-cameras and SPECT scanners in the past (Peremans et al., 2005), VEGFR imaging was achieved with SPECT earlier than with PET. A number of radioisotopes, such as 123I, 111In, 99mTc, and 64Cu, have been used for SPECT/PET imaging of VEGFR in the tumor vasculature.

6.1. SPECT imaging To explore the possibility of VEGFR scintigraphy of primary tumors and their metastases, the binding properties of 123I-VEGF165 and 123I-VEGF121 to human umbilical vein endothelial cells (HUVECs), several human cancer

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cell lines, and a variety of primary human tumors were evaluated (Li et al., 2001). Besides HUVECs, some tumor cells were also found to express high levels of VEGFR in this study. 123I-VEGF165 was then evaluated for tumor localization in gastrointestinal cancer patients (Li et al., 2003). However, the primary and metastatic lesions were identified only in some patients. In another study, the biodistribution, safety, and absorbed dose of 123IVEGF165 were investigated in nine patients with pancreatic carcinoma (Li et al., 2004). Although the majority of primary pancreatic tumors and their metastases were visualized by 123I-VEGF165 scans (Fig. 7.4), the organ with the highest absorbed dose was the thyroid due to severe deiodination. Another report also demonstrated that 123I-VEGF165, despite the high receptor-binding affinity, gave poor tumor-to-background ratio in tumorbearing mice due to low metabolic stability (Cornelissen et al., 2005). Recently, 125I-labeled VEGF121 and VEGF165 were tested for biodistribution and autoradiography studies in tumor-bearing mice (Yoshimoto et al., 2006). As with most other radio-iodinated tracers, prominent activity accumulation in the stomach was observed due to deiodination. Interestingly, 125I-VEGF121 accumulation in tumors decreased with increasing tumor volume, suggesting that small tumors have higher VEGFR expression than large tumors. It was also found that 125I-VEGF165 uptake was higher than that of 125I-VEGF121 in some organs (e.g., the kidneys, heart, and lung), but lower in many other organs (Yoshimoto et al., 2006). However, the reason for such difference in the accumulation pattern between the two tracers remains unclear.

CT

123I-VEGF 165

Figure 7.4 A transverse CT image of a pancreatic adenocarcinoma patient and a transverse SPECT image of the same patient at 1.5-h postinjection of 123I-VEGF165. (Adapted from Li, S., Peck-Radosavljevic, M., Kienast, O., Preitfellner, J., Havlik, E., Schima,W.,Traub-Weidinger,T., Graf, S., Beheshti, M., Schmid, M., Angelberger, P., and Dudczak, R. (2004). Iodine-123^vascular endothelial growth factor-165 (123I-VEGF165). Biodistribution, safety and radiation dosimetry in patients with pancreatic carcinoma. Q. J. Nucl. Med. Mol. Imaging 48,198^206.)

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Besides radioiodine, VEGF121 has also been labeled with 99mTc (Backer et al., 2007; Blankenberg et al., 2004). In a 4T1 murine mammarycarcinoma model, the tumor uptake of 99mTc-VEGF121 was found to be about 3% ID/g. It was suggested that 99mTc-labeled VEGF121, stable for about 1 h in vivo, can be used to image mouse tumor neovasculature in lesions as small as several millimeters in soft tissue. Recently, this tracer was also employed for imaging the tumor vasculature before and after different types of chemotherapy (Blankenberg et al., 2006). A recombinant protein composed of VEGF165 fused through a flexible polypeptide linker, (GGGGS)3, to the n-lobe of human transferrin (hnTfVEGF) was constructed for angiogenesis imaging (Chan et al., 2005). The molecular weight of hnTf-VEGF is 65 kDa and 130 kDa for the monomeric and dimeric form, respectively. 111In-hnTf-VEGF accumulation in the U87MG human glioblastoma tumors was about 6.7 %ID/g at 72 h postinjection (p.i.). The tumor uptake decreased when coinjected with 100-fold excess of VEGF but not with apotransferrin. This fusion protein, hnTfVEGF, represents a new class of proteins that can be labeled with 111In without the need to introduce metal chelators. All abovementioned reports used radio-labeled VEGF isoforms for SPECT imaging of VEGFR expression. Although the VEGF isoforms used in these studies all exist in nature and should have very high binding affinity and specificity to VEGFRs, much research effort is still needed in the future to improve the in vivo stability, target affinity/specificity, and pharmacokinetics of these radiopharmaceuticals. The major disadvantage of SPECT imaging is its very low detection efficiency (typically <10–4 times the emitted number of gamma rays) because of the use of lead collimators to define the angle of incidence (Chatziioannou, 2005). Another imaging modality, PET, offers many advantages over SPECT, and the increasing popularity of the clinical PET and PET/CT scanners can facilitate clinical translation of promising new tracers (Beyer et al., 2000; Townsend and Beyer, 2002).

6.2. PET imaging We have labeled VEGF121 with 64Cu for PET imaging of VEGFR expression (Cai et al., 2006a). DOTA-VEGF121 has nanomolar VEGFR-2–binding affinity (comparable to VEGF121) in vitro (Fig. 7.5A and B). Micro-PET imaging revealed rapid, specific, and prominent uptake of 64Cu-DOTAVEGF121 (
15% ID/g) in highly vascularized small U87MG tumors (high VEGFR-2 expression) but significantly lower and sporadic uptake (2 to 3% ID/g) in large U87MG tumors (low VEGFR-2 expression) (Fig. 7.5C). Western blotting of the tumor tissue lysate, immunofluorescence staining (Fig. 7.5C), and blocking studies with unlabeled VEGF121 all confirmed that the tumor uptake of 64Cu-DOTA-VEGF121 is VEGFR-specific. This study demonstrated the dynamic nature of VEGFR expression during

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Figure 7.5 PET imaging of VEGFR expression. (A) Schematic structure of 64CuDOTA-VEGF121. (B) Cell-binding assay of VEGF121 and DOTA-VEGF121 using PAE/ VEGFR-2 cells. IC50 values are 1.02 nM and 1.66 nM for VEGF121 and DOTA-VEGF121, respectively. (C) PET imaging using 64Cu-DOTA-VEGF121 revealed much higher tumor uptake in the small U87MG tumor (
15% ID/g) than in the large tumor (2 to 3% ID/g). Immunofluorescence staining revealed that VEGFR-2 expression is much higher in the small tumor than in the large tumor. (Adapted from Cai,W., Chen, K., Mohamedali, K. A., Cao, Q., Gambhir, S. S., Rosenblum, M. G., and Chen, X. (2006a). PETof vascular endothelial growth factor receptor expression. J. Nucl. Med.47, 2048^2056.).

tumor progression in that even for the same tumor model, VEGFR expression level is dramatically different at different sizes/stages. Our recent study revealed that the tumor uptake of 64Cu-DOTA-VEGF121 peaked during a narrow size range (100 to 250 mm3 for U87MG tumors) when high VEGFR-2 expression was present based on histology (Chen et al., 2008).

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Most importantly, the tumor uptake value obtained from PET imaging had good linear correlation with the relative tumor-tissue VEGFR-2 expression as measured by Western blot (r2 equals 0.68 based on the PET uptake at 4 h p.i.). Such finding underscores the importance of imaging VEGFR expression in anti-VEGFR cancer therapy, since the therapeutically efficacious window may be quite narrow. Noninvasive imaging to evaluate the VEGFR expression level before the initiation of treatment can help decide whether, and when, to start antiangiogenic treatment, thus improving cancer patient management in the future. The linear correlation between tracer uptake and target expression level will also significantly facilitate future treatment planning and treatment monitoring of cancer as well as other angiogenesis-related diseases. In a follow-up study, we tested a VEGFR-2–specific fusion protein VEGF121/rGel (VEGF121 linked to a recombinant plant toxin, gelonin) for treating orthotopic glioblastoma in a mouse model (Hsu et al., 2007). Before initiation of treatment, micro-PET imaging with 64Cu-labeled VEGF121/rGel was carried out to evaluate its tumor-targeting efficacy and pharmacokinetics, thus determining the dose interval. Such study with a radio-labeled drug clearly demonstrated the power of molecular imaging, where cancer patients can also be selected for molecular cancer therapy based on pretreatment screening using a radio-labeled drug (analog). During and after treatment, the therapeutic efficacy of VEGF121/rGel was successfully monitored by multimodality imaging using 18F-FLT PET, MRI, and BLI, all of which were validated by ex vivo histology. This study suggested that future clinical multimodality imaging and therapy with VEGF121/rGel may provide an effective means to prospectively identify patients who will benefit from VEGF121/rGel therapy, and then stratify, personalize, and monitor the treatment to obtain optimal survival outcomes. VEGF121 has also been labeled with 64Cu site-specifically (Backer et al., 2007). Compared with the 99mTc-labeled analog where the tumor uptake (
3% ID/g) was lower than most of the normal organs and the kidney uptake was about 120% ID/g, the 64Cu-labeled PEGylated VEGF121 gave similar tumor uptake (
2.5% ID/g) and a lower kidney uptake of about 65% ID/g. Whether such site-specific labeling through an extra cysteine tag is advantageous over direct labeling at the lysine side-chain amino groups, which has been shown to not affect the VEGFR-2–binding affinity (Cai et al., 2006a), remains to be determined. Direct comparison of the two labeling methods, as well as potentially other strategies, is needed to determine which tracer is more suitable for clinical evaluation. All VEGF-A isoforms bind to both VEGFR-1 and VEGFR-2 (Ferrara, 2004). In most of the imaging studies discussed above, specificity to either VEGFR-1 or VEGFR-2 was not achieved as most of the tracers are based on VEGF-A isoforms. Due to high VEGFR-1 expression, the rodent kidneys can take up significant amount of VEGF-A–based tracer, which

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often makes it the dose-limiting organ (Backer et al., 2007; Cai et al., 2006a; Simon et al., 1998). Since VEGFR-2 is generally accepted as more functionally important than VEGFR-1 in cancer progression (Ferrara, 2004; Underiner et al., 2004), the ability to image VEGFR-2 expression by PET can be a valuable tool for evaluating patients with a variety of malignancies, particularly those undergoing anti–VEGFR-2 therapies. Alanine-scanning mutagenesis has revealed that the residues Arg82, Lys84, and His86, located in a hairpin loop of VEGF165, were critical for VEGFR-2 binding, and some negatively charged residues, Asp63, Glu64, and Glu67, were associated with VEGFR-1 binding (Keyt et al., 1996). As VEGF121 is a soluble, non–heparin-binding variant containing the full biological and receptor-binding activity of the larger variants (Ferrara, 2004), we recently developed a VEGFR-2–specific PET tracer based on mutated VEGF121 (Wang et al., 2007). The D63AE64AE67A mutant of VEGF121 (denoted as VEGFDEE) was generated by recombinant DNA technology. Compared to VEGF121, VEGFDEE has 20-fold lower binding affinity to VEGFR-1 without significant reduction in VEGFR-2–binding affinity. MicroPET imaging demonstrated that 64Cu-DOTA-VEGF121 and 64Cu-DOTA-VEGF DEE had comparable activity accumulation in VEGFR-2–expressing 4T1 tumors. Meanwhile, the renal uptake of 64Cu-DOTA-VEGF 64Cu-DOTADEE was much lower than that of VEGF121 due to the much lower VEGFR-1–binding affinity of DOTAVEGFDEE. Significantly lower kidney uptake of 64Cu-DOTA-VEGFDEE, conferring much lower renal toxicity than other VEGF-A–based tracers, makes it highly amenable for clinical translation. Further improvement in VEGFR-2–binding affinity/specificity, in vivo pharmacokinetics, and tumor-targeting efficacy by generating other VEGF121 mutants is currently being investigated.

7. Experimental Section A molecular imaging agent, defined as ‘‘a probe used to visualize, characterize, and measure biological processes in living systems’’ (Mankoff, 2007), is typically composed of an imaging label, a carrier that contains (a) targeting ligand(s) or is a targeting ligand, and a linker between the carrier and the label. Both endogenous molecules and exogenous agents can be molecular imaging agents and in some cases not all three components are present. In this section, we will describe in detail the entire process of micro-PET imaging of tumor vasculature in living mice. The molecular target is VEGFR expressed on the tumor vasculature, the targeting ligand is VEGF121, and the imaging label is 64Cu, which can be complexed through a DOTA chelator.

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7.1. Production of VEGF121 protein Detailed procedures for preparing VEGF121 have been reported earlier using various methods (Cai et al., 2006a; Wang et al., 2007). Here we will describe one of them. A pair of primers were designed and synthesized for VEGF121: CATATGGCACCCATGGCAGAAGGAGGA (sense) and CTCGAGTCAGTGGTGATGATGGTGATGGGATCCCCGCCTCG GCTTGTCAC (antisense). An NdeI and XhoI restriction site was incorporated into each respective flanking primer. The two polymerase chain reaction (PCR) products were then annealed and extended. After confirming the DNA sequence, the gene encoding VEGF121 was released by NdeI and XhoI (New England Biolabs) double digestion and ligated into a pRSF-Duet1 expression vector (pretreated with the same restriction endonuclease enzymes [Novagen]). The plasmid containing the gene was transfected into Escherichia coli BL21 (DE3). The single clone was cultured in 5 ml of Luria broth (LB) medium (Invitrogen) containing 30 mg/ml of kanamycin at 37  C with constant shaking. The overnight cultures were inoculated into 2 l of LB medium and cultured until the OD600nm was between 0.6 and 1.0. Protein expression was then induced by adding isopropyl-b-D-thiogalactopyranoside (IPTG) (QiaGen) to a final concentration of 0.5 mM and cultured for 4 h. The bacteria were harvested and the pellet was resuspended in lysis buffer (20 mM NaH2PO4, 10 mM tris-HCl, 10 mM imidazole, 300 mM NaCl, pH ¼ 8.0) for sonication. After centrifuging at 12,000 rpm for 20 min, the supernatant was collected and incubated with Ni-NTA agarose (QiaGen) at 4  C for 1 h. The His-tagged recombinant protein was eluted from the agarose by elution buffer (20 mM NaH2PO4, 10 mM tris-HCl, 300 mM imidazole, 300 mM NaCl, pH ¼ 8.0), dialyzed against PBS, and stored at –80  C for later use. Alternatively, a slightly different protein based on VEGF121 is commercially available from SibTech, which can also be used for the following experiments.

7.2. DOTA conjugation Detailed procedures for DOTA conjugation of peptides or proteins have been reported earlier (Cai et al., 2006d; Wu et al., 2005). DOTA was activated by 1-ethyl-3-[3-(dimethylamino)-propyl] carbodiimide (EDC) and N-hydroxysulfonosuccinimide (SNHS) at pH 5.5 for 30 min with a molar ratio of DOTA:EDC:SNHS at 10:5:4. Without purification, the DOTA-OSSu was cooled to 4  C and added to VEGF121 at a molar ratio (DOTA-OSSu: VEGF121) of 50:1. The reaction mixture was adjusted to pH 8.5 with 0.1 N NaOH solution and allowed to incubate overnight at 4  C. The DOTAVEGF121 conjugate was then purified using PD-10 column (GE Healthcare) and concentrated by Centricon filter with 10-kDa molecular-weight cut-off (MWCO) (Millipore). The final concentration of DOTA-VEGF121 was

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measured based on UV absorbance at 280 nm using unconjugated VEGF121 of known concentrations as the standard. The average number of DOTA chelators per VEGF121 can be determined using the isotope-dilution method (Cai et al., 2006a; Meares et al., 1984). Typically, we found that there are about one to two DOTA chelators per VEGF121 molecule.

7.3. Cell-binding assay of VEGF121 and DOTA-VEGF121 DOTA conjugation at the amino groups in VEGF121 protein may affect the VEGFR-binding affinity. Therefore, we recommend that cell-binding assays be carried out using both VEGF121 and DOTA-VEGF121 before in vivo imaging studies. It is imperative to obtain appropriate training from the institutional radiation safety office and have all related protocols approved before ordering radioactivity. Abide by all relevant regulatory rules and use appropriate protection when handling radioactivity (125I and 64Cu for this experiment). Dispose of all radioactive waste according to the institutional radioactive waste disposal guidelines. For detailed procedure of a cell-binding assay, see Cai and Chen (2008b). In brief, VEGFR-1– and VEGFR-2–binding affinity of VEGF121 and DOTA-VEGF121 can be evaluated by PAE/VEGFR-1 (porcine aorta endothelial cells transfected with VEGFR-1) and PAE/VEGFR-2 (PAE cells transfected with VEGFR-2) cell-binding assays using 125I-VEGF165 (GE Healthcare) as the radio-ligand. The best-fit IC50 values can be calculated by fitting the data by nonlinear regression using GraphPad Prism (GraphPad Software, Inc.). We typically found the IC50 values of VEGF121 and DOTA-VEGF121 to be about 1 to 5 nM (Fig. 7.5B).

7.4. Cell lines and animal models Obtain appropriate training from the institution regarding biosafety and animal handling. Animal protocols must be in place before performing any animal studies. We have used two tumor models for VEGFR imaging studies—the U87MG human glioblastoma and 4T1 murine breast cancer tumor models (Cai et al., 2006a; Wang et al., 2007). The cell lines were obtained from American Type Culture Collection (ATCC) and cultured under standard conditions. The cells need to be tested as free of murine virus before inoculation into the mice. The U87MG tumor model was generated by subcutaneous injection of 5  106 cells in 50 ml of PBS into the front flank of female athymic nude mice (4 to 6 weeks old) (Harlan). The mice were subjected to micro-PET imaging studies when the tumor volume reached about 60 to 150 mm3 (small tumor, 2 to 3 weeks after inoculation) or
1000 mm3 (large tumor, 5 to 6 weeks after inoculation). For the 4T1 tumor model, female BALB/c

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mice (4 to 6 weeks old) (Harlan) were injected with 2  106 4T1 cells in 100 ml of PBS into the front flank of each mouse. Two weeks after inoculation when the tumor volume reached about 100 to 200 mm3, the mice were subjected to in vivo studies.

7.5.

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of DOTA-VEGF121

Water and all buffers should be treated with Chelex 100 resin (Sigma) before use in radio-labeling procedures to ensure heavy metal–free conditions. Otherwise, the trace amount of metal ions in the solution will compete with 64Cu for DOTA complexation and results in low radiolabeling yield. Perform the radio-labeling behind appropriate lead shielding (Fig. 7.6A). Two mCi of 64CuCl2 were added to a solution of DOTAVEGF121 in 300 ml of 0.1 M sodium acetate buffer (pH ¼ 6.5; 10 mg of DOTA-VEGF121 per mCi of 64Cu). The reaction mixture was incubated for 1 h at 40  C with constant shaking. The 64Cu-DOTA-VEGF121 conjugate can then be purified by PD-10 column using PBS as the mobile phase. The radioactive fractions (typically between 3.5 and 4.5 ml, total of 1 ml) containing 64Cu-DOTA-VEGF121 were collected for further in vitro and in vivo experiments. A typical size exclusion column chromatography profile should look like that in Fig. 7.6B. After 6 ml of PBS, the unreacted 64Cu may start to elute from the column.

7.6. Micro-PET imaging Under anesthesia with 2% isoflurane in oxygen (Fig. 7.7A and B), the mice were injected with 200 to 300 mCi of 64Cu-DOTA-VEGF121 (about 2 to 3 mg of VEGF121) via the tail vein (Fig. 7.7C). PET imaging of tumor-bearing B

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Figure 7.6 64Cu-labeling of DOTA-VEGF121. (A) An L-block with lead glass shield for 64 Cu-labeling experiments. (B) Size-exclusion column chromatography profile for 64 Cu-DOTA-VEGF121 purification.The unreacted 64Cu elutes after 6 ml.

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D

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Figure 7.7 The setup for micro-PET imaging of tumor-bearing mice. (A) An anesthesia system that uses isoflurane. (B) A knockdown chamber on a warm-water bed is used for anesthetizing the mice. (C) A tumor-bearing mouse (front right flank) under anesthesia before tail vein injection of 64Cu-DOTA-VEGF121. (D) A micro-PETscanner. (E) Position of the mouse before the micro-PETscan. (F) A tumor-bearing mouse during a micro-PETscan.

mice was performed on a micro-PET-R4 rodent model scanner (Siemens Medical Solutions; Fig. 7.7D). At various time points p.i., the mice were anesthetized again and placed in prone position near the center of the field of view where the highest image resolution and sensitivity are available (Fig. 7.7E and F). Typically, 3- to 10-min static scans were performed depending on the radioactivity left in each mouse. The images were reconstructed by a two-dimensional, ordered-subsets expectation maximum (OSEM) algorithm, with no attenuation or scatter correction. To quantify the tracer uptake in the tumor and each organ, a conversion factor should be determined using a 50-ml centrifuge tube filled with 64CuCl2 as a mouse phantom. For each micro-PET scan, three-dimensional ROIs were drawn over the tumor and major organs by using the vendor software on decaycorrected whole-body coronal images (Fig. 7.8). Values in several adjacent slices (depending on the size of the tumor or organ) were averaged to obtain a reproducible value of activity concentration in the ROI drawn. Assuming a tissue density of 1 g/ml, the measured values of the ROIs were converted to mCi/g/min and then divided by the administered radioactivity to obtain image ROI-derived % ID/g values. To obtain statistically meaningful results, each group should have at least three mice. Several control experiments are suggested to confirm the in vivo VEGFR specificity of 64Cu-DOTA-VEGF121. For example, mice bearing large tumors with low VEGFR expression can be a control group; in vivo receptor-blocking experiments (co-injection of unconjugated VEGF121 and

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Figure 7.8 A screenshot of the software used for region-of-interest analysis.The image is the same as that shown in Fig.7.5C. 64Cu-DOTA-VEGF 121)

can serve as another control; mice bearing other low VEGFR-expressing tumors may be used as controls; and imaging the high VEGFR-expressing tumor-bearing mice with a 64Cu-DOTA-VEGFmutant (which does not bind to VEGFRs) may also be carried out to further validate the conclusion.

7.7. Biodistribution studies After the last micro-PET scans, the mice can be euthanized for biodistribution studies to confirm the in vivo imaging results. Collect and wet weigh the blood samples, tumor, major organs and tissues (contents in the intestines should be removed before weighing). The radioactivity in each tissue was measured using a g-counter and the results were calculated as % ID/g. For each mouse, the radioactivity in the tissue sample was calibrated against a known aliquot of the injectate and normalized to a body weight of 20 g. Typically we found that the data of most tissues obtained from micro-PET and biodistribution studies correlate very well (Cai et al., 2006a, 2006d, 2006e, 2006c, 2007e; Liu et al., 2007).

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7.8. Immunofluorescence staining To further validate the VEGFR expression level in the tumor tissue, we recommend immunofluorescence staining of frozen U87MG tumor tissue slices. Most of the needed antibodies are available from Jackson Immuno Research Laboratories, Inc. Typically, 5-mm-thick tumor slices were fixed with cold acetone for 10 min and dried in the air for 30 min. The slices were rinsed with PBS for 2 min and blocked with 10% donkey serum for 30 min at room temperature. After incubating with a rat anti-mouse VEGFR-2 antibody overnight at 4  C, the slices were visualized using a Cy3-conjugated donkey anti-rat secondary antibody (1:200) (Fig. 7.5C). For VEGFR-1 staining, the tissue slices were incubated with rabbit anti-mouse VEGFR-1 antibody (1:50) at room temperature for 1 h and visualized with Cy3-conjugated donkey antirabbit secondary antibody (1:200). Alternatively, Western blotting of the tissue lysates can also be performed to examine the VEGFR expression level in the tumor and other organs of interest.

8. Summary Molecular imaging of tumor vasculature has clinical applications in many aspects: lesion detection, patient stratification, new drug development/validation, treatment monitoring, and dose optimization. Integrin avb3 and VEGFRs are the key regulators of tumor angiogenesis and many reports are available in the literature. Imaging VEGF and VEGFR expression are both important for cancer diagnosis and monitoring the therapeutic efficacy against cancer. Due to the dynamic nature of VEGF proteins, imaging VEGF expression is much more challenging than imaging VEGFR, and it has not been well studied (Collingridge et al., 2002; Jayson et al., 2002; Nagengast et al., 2007). Examining the tumor in the same animals or cancer patients with both VEGF- and VEGFR-targeted tracers may give important insights about the expression kinetics of VEGF and VEGFRs during cancer development and cancer therapy. Nanotechnology can have many applications in imaging VEGFR expression in the future. Surprisingly, no such study has been reported. Other proteins involved in tumor angiogenesis, such as endoglin (CD105) (Fonsatti et al., 2003), should also be investigated in the future to enlarge our arsenal of tumor-vasculature imaging probes. Quantitative imaging of tumor angiogenesis is highly desirable in the future. Although it is generally assumed that noninvasive imaging results correlate with the target expression level in vivo, such assumption has not been extensively validated. In most reports, two tumor models are studied where one acts as a positive control and the other as a negative control.

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Quantitative correlation between the target expression level in vivo and the noninvasive imaging data is rare (Cai et al., 2007a, 2007c; Chen et al., 2008; Zhang et al., 2006). Such correlation is critical for future treatment monitoring applications, as it would be ideal to be able to monitor the changes in the target expression level quantitatively, rather than qualitatively, in each individual patient. Upon further development of new tracers with better tumor targeting efficacy and desirable pharmacokinetics, clinical translation will be critical for the maximum patient benefit of these imaging probes. Clinical translation of new molecular imaging agents has been very slow in the past. However, the situation has started to change recently. Continued development and wider availability of scanners dedicated to small animal imaging studies has enabled smooth transfer of knowledge and molecular measurements among species, which significantly facilitates clinical translation. The Food and Drug Administration (FDA) has recently developed the exploratory Investigational New Drug (IND) mechanism to allow faster first-inhuman studies. To foster the continued discovery and development of tumor vasculature–targeted imaging probes, interdisciplinary and cooperative efforts from cellular/molecular biologists, chemists/radiochemists, engineers, medical physicists, and mathematicians are needed. Close partnerships among academic researchers, clinicians, pharmaceutical industries, the National Cancer Institute (NCI), and the FDA are also desired to quickly apply tumor vasculature imaging to multiple facets of anticancer clinical trials and cancer patient management, thus paving the way to personalized molecular medicine.

ACKNOWLEDGMENTS Research carried out in the authors’ laboratory was supported by the National Cancer Institute (R01 CA119053, R01 CA082214, R01 HL078632, R01 NIBIB BRP, R21 CA121842, R21 CA102123, P50 CA114747, U54 CA119367, and R24 CA93862), the Department of Defense (DOD) (W81XWH-07-1-0374, W81XWH-04-1-0697, W81XWH-06-1-0665, W81XWH-06-1-0042, and DAMD17-03-1-0143), Doris Duke Foundation, Canary Foundation, General Electric Medical Systems, Bayer-Schering, Genentech Inc., and a Benedict Cassen Postdoctoral Fellowship from the Education and Research Foundation of the Society of Nuclear Medicine.

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Proteomic Mapping of the Vascular Endothelium In Vivo for Vascular Targeting Noelle M. Griffin and Jan E. Schnitzer Contents 1. Introduction 2. Proteomics: An Overview 3. Proteomic Methodologies 3.1. Two-dimensional electrophoresis 3.2. Liquid chromatography 3.3. Mass spectrometry 3.4. Bioinformatics 4. Databases 5. Quantitative Proteomics 5.1. Label-free approaches 6. Application of Proteomics to the Vasculature 7. Summary 8. Protocols 8.1. Isolation and subfractionation of plasma membranes 8.2. Mass spectrometric analysis of endothelial cell membrane proteins References

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Abstract The mapping and characterization of the vasculature using proteomics offers the opportunity to better understand the steps and molecular mechanisms involved in vascular development, and angiogenesis in particular. Proteomics has many key advantages over genomics, especially in directly determining protein expression. Such an approach offers researchers the opportunity to discover the proteins that make up the vasculature in an a priori manner facilitating the generation of hypothesis which can subsequently be validated

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by other methods. This chapter focuses on proteomic principles and methods, with a particular focus on their applications to characterizing the vascular endothelium (both tumor and normal), as it exists in vivo.

1. Introduction Angiogenesis is a physiological process involving the growth of new blood vessels from pre-existing vessels. It is a normal process in growth and development, as well as in wound healing. In many serious diseases states, the body loses control over angiogenesis. Angiogenesis-dependent diseases result when new blood vessels either grow excessively or insufficiently. Excessive angiogenesis occurs in diseases such as cancer, diabetic blindness, age-related macular degeneration, rheumatoid arthritis, psoriasis, and more than 70 other conditions. In these conditions, new blood vessels feed diseased tissues and destroy normal tissues, and in the case of cancer, the new vessels allow tumor cells to escape into the circulation and lodge in other organs (tumor metastases). Excessive angiogenesis occurs when diseased cells produce abnormal amounts of angiogenic growth factors, overpowering the effects of natural angiogenesis inhibitors. Insufficient angiogenesis occurs in diseases such as coronary artery disease, stroke, and delayed wound healing. In these conditions, inadequate blood vessels grow, and circulation is not properly restored, leading to the risk of tissue death. Insufficient angiogenesis occurs when the tissue cannot produce adequate amounts of angiogenic growth factors. Therefore, by better understanding the steps and molecular mechanisms involved in angiogenesis, better, more targeted therapies may be developed in the treatment of angiogenesis-related disorders. Our interest is in tumor angiogenesis. Tumors cannot grow beyond a certain size, generally 1 to 2 mm3, due to a lack of oxygen and other essential nutrients. Therefore, the subsequent growth and survival of such a tumor is dependent on acquiring its own blood supply. Tumor angiogenesis starts with cancerous tumor cells releasing molecules that send signals to surrounding normal host tissue. This signaling activates certain genes in the host tissue that, in turn, make proteins to encourage growth of new blood vessels. Various growth factors, such as bFGF (basic fibroblast growth factor) and VEGF (vascular endothelial growth factor) are secreted, which can induce new vascular growth into the tumor, thus supplying the required nutrients to allow tumor expansion. Angiogenesis is also required for the spread of a tumor, or metastasis. Single cancer cells can break away from an established solid tumor, enter the blood vessel, and be carried to a distant site, where they can implant and begin the growth of a secondary tumor. The subsequent growth of such metastases will also require a supply of nutrients, oxygen, or a waste disposal pathway. The inner surface of blood vessels is lined by a thin monolayer of endothelial cells that form a critical interface and barrier between the

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circulating blood and the cell within any tissue. Endothelial cells are normally quite quiescent and do not proliferate rapidly. However, when stimulated by angiogenic factors, they rapidly proliferate and migrate to form new blood vessels. Endothelial cells have long been considered genetically more stable than cancer cells. This genomic stability confers an advantage to targeting endothelial cells using antiangiogenic therapy compared to chemotherapy directed at cancer cells, which rapidly mutate and acquire ‘‘drug resistance’’ to treatment. It has been suggested that traditional therapies, such as radiation therapy, may actually work in part by targeting the genomically stable endothelial cell compartment, rather than the genomically unstable tumor cell compartment. Tumor cells evolve resistance rapidly due to rapid generation time (days) and genomic instability (variation), whereas endothelial cells are a good target because of a long generation time (months) and genomic stability (low variation). For this reason, endothelial cells are thought to be an ideal target for therapies directed at them. In addition, most cancer-associated proteins are expressed by malignant cells inside the tumor compartment not readily accessible to intravenously (i.v.) injected agents. With this in mind and due to the fact that a tumor must develop its own blood supply in order to thrive, the benefit to be gained from specifically targeting the vasculature of the tumor itself for drug delivery becomes readily apparent. This approach is very attractive as the endothelial surface lining the vasculature shows molecular heterogeneity induced by its specific tumor microenvironment, allowing the vasculature of the tumor to be distinguished from the ‘‘normal’’ vasculature (Oh et al., 2004b). As the plasma membrane of the luminal surface of the vasculature supplying the underlying tumor cells is in constant contact with the blood, it represents an ideal surface for targeted delivery of drug and therapeutic agents, especially as it is readily accessible to i.v. injected agents. Furthermore, it has been proposed that specific delivery into cells and tissues underlying the vascular endothelium can be improved by targeting caveolae (plasmalemmal invaginations abundant in endothelium) and exploiting their transcytotic function and membrane-trafficking properties (Carver and Schnitzer, 2003; Schnitzer, 1993, 2001; Schnitzer et al., 1995a). By generating a molecular fingerprint of the vascular endothelium, in particular the plasma membrane and caveolae, of various organs, tissues and tumors, the door is opened to discovering tissue- and tumor-specific drugable targets. Although the genome provided the basis for a better understanding of cellular and molecular mechanisms, neither the transcriptional profile nor the genomic sequence can be directly correlated with protein expression; thus, the importance of measuring protein levels becomes increasingly clear. Because proteins are involved in virtually every cellular function, most activities performed by cells will be reflected in their proteomes. The most commonly used and appropriate techniques for such experiments are proteomics methodologies. This chapter focuses on

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proteomic principles and methods, with a particular focus on their applications to characterizing the vascular endothelium (both tumor and normal), as it exists in vivo.

2. Proteomics: An Overview The proteome is the set of all expressed proteins in a cell, tissue, or organism (Pennington et al., 1997). Proteomics differs from genomics in both complexity and dynamic variability. Whereas the genome is relatively constant, the proteome is constantly changing according to the moment-tomoment interactions between the genome and the environment. It is often conceptualized that one gene encodes one protein; it is known that the expressed products of a single gene in reality represent a protein population with a high degree of heterogeneity. The DNA sequence alone cannot be used to build a molecular description of a cell. Proteins comprise a major portion of cell structural molecules and are the effecter molecules for all cell functions, either directly or indirectly through their actions on metabolites and other nonprotein substrates. Biological function appears to be carried out by the dynamic population of proteins determined by an interaction of gene and protein regulation with extracellular influences. There is no strict linear relationship between genes and the protein complement or ‘‘proteome’’ of a cell. Although each gene clearly encodes a specific ‘‘protein core’’ through transcription and RNA translation, alternative splicing greatly expands protein diversity beyond a one gene—one protein concept. Using gene expression, specifically mRNA levels, to infer protein expression can be quite misleading because the regulatory mechanisms mediating gene and protein expression can differ and can be uncoupled. Newly expressed protein can remain long after shutting down the gene encoding process. The ‘‘protein core’’ encoded by a specific gene expands into many distinct proteins through a wide variety of post-translational modifications, including, but not limited to, glycosylation, phosphorylation, ubiquitination, oxidation, alkylation, and biotinylation. This effectively increases the number of different proteins present. For example, a protein that has three potential modification states (glycosylation, phosphorylation, and ubiquitination) has eight potential protein forms if each modification only occurs at a single site and is not mutually exclusive. Each additional state (e.g., another phosphorylation, acetylation, protease cleavage, etc.) or modification site adds diversity to the expression profile of that protein (64 potential forms for just six modifications) (Peng and Gygi, 2001). These posttranslational modifications can greatly alter the function of a protein, and thus even the complete sequence of a genome is insufficient to elucidate biological function.

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Advances in the use of mass spectrometry (MS) over the last 5 to 10 years, in conjunction with DNA/protein database—searching algorithms, have opened the door for the identification and quantification of proteins with unprecedented speed. However, there are still analytical challenges in accurate quantification of proteins and the detection of site-specific modifications. Current mass spectrometry—based strategies for protein identification and quantification can be classified into two main streams: two-dimensional polyacrlyamide gel electrophoresis (2D-PAGE) followed by MS analysis has been a widely used approach. A second, more recent strategy is based on multidimensional chromatography coupled to MS to separate, identify and quantify tryptic peptides derived from complex mixtures (Gygi et al., 1999a; Washburn et al., 2001). Both approaches have advantages and disadvantages, which are outlined below.

3. Proteomic Methodologies 3.1. Two-dimensional electrophoresis The technique of two-dimensional (2D) protein separation was first described by O’Farrell et al.(1975). In this method, proteins are separated in the first dimension according to isoelectric point (pI), followed by separation in the second dimension according to relative molecular mass (Mr). Protein molecules are very diverse biophysically, with each protein having a particular charge, either positive or negative, that in turn is dependent on the chemical environment. This charge is exploited in isoelectric focusing (IEF). Proteins are separated or ‘‘focused’’ along a pH gradient until each individual protein reaches the pH at which it has no charge. This pH is known as the isoelectric point. The proteins are further separated according to their molecular weight by sodium dodecyl sulfate PAGE (SDS-PAGE), thus giving a second dimension of separation. The resulting 2D gel consists of a map of separated proteins or spots that can be visualized by a number of staining techniques, such as silver, coomassie, and fluorescent stains. Theoretically each spot represents an individual protein, but this is not always the case, due to co-migratory proteins. Two-dimensional gels have the ability to display a large number of resolved proteins (a few thousand) from specific samples on a single gel. The actual visualization of protein spots not only has an aesthetic appeal but an obvious pragmatic benefit when comparing samples to identify and even quantify differential expression. Unfortunately, there are significant limitations in using 2D electrophoresis (2DE) for large-scale comprehensive mapping, including its poor ability to resolve membrane and hydrophobic proteins. An estimated 30% of proteins are membrane proteins and only approximately 1% of them can be resolved properly on 2D gels with current

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techniques (Santoni et al., 2000). Some improvement with technical advances can be seen. The increased use of denaturing reagents such as thiolurea (Rabilloud, 1998) and zwitterionic detergents (Chevallet et al., 1998) can improve 2DE analysis of membrane proteins. Other ‘‘tricks’’ include prewashing of the sample with alkaline buffer prior to solubilization with the detergent C8phi, which can assist in hydrophobic protein entry into the gel (Santoni et al., 1999). Another challenge for 2DE is a limited dynamic range of the proteins that can be displayed (Gygi et al., 2000). Poor detection levels are, in part, due to the practical limitations of the imaging technique used– silver staining cannot visualize spots representing less than 0.5 ng of protein. Furthermore, co-migratory proteins can also affect protein visualization. This is especially important, as visualization of proteins of low abundance, including signaling proteins and transcription factors, can become masked by larger, more abundant proteins such as actin, which may migrate to the same spot on the gel. Even the best 2D gel can routinely resolve no more than 1000 different proteins, with highly basic and acidic proteins–that is, those with extreme isoelectric points and extreme molecular weights–being excluded (Gorg et al., 2004; Gygi et al., 2000). However, a more fundamental problem is the mismatch between the dynamic range of protein expression and the capacity of gel-based IEF. The quantity of sample that must be loaded onto a 2D gel to detect a low-level protein reliably would significantly exceed the gel’s capacity. This problem has led to a systematic bias in 2DE protein profiling toward more abundant proteins (Gygi et al., 1999b). Therefore, only a small subset of complex eukaryotic proteomes can be visualized by 2DE. A number of enrichment techniques have been used to get around this problem. For example, immunodepletion of the more abundant proteins can be employed prior to 2DE (Pieper et al., 2003). In addition, prefractionation of the sample into different cellular components prior to 2DE application can facilitate the identification of less abundant proteins. Also, the advent of zoom gels or narrow pH range gels have helped ease this problem, with reports of better capacity to separate different protein species and isoforms (O’Neill et al., 2002; Westbrook et al., 2001). Another important consideration for 2DE experiments is throughput. A 2DE experiment typically takes 3 to 5 days to complete. Although gels can be run simultaneously in batches, the process is still time consuming and technically demanding. Lastly, the MS analysis of each spot has been automated robotically, but may still lack sufficient throughput and robustness to meet the needs of large-scale comprehensive profiling.

3.2. Liquid chromatography Liquid chromatography coupled to tandem MS, called LC-MS/MS, is a powerful technique for the analysis of peptides. Proteins are initially digested into peptides by specific proteases such as trypsin. The peptide

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sample is cleaned, fractionated, and concentrated in a single step prior to elution into the ESI ion source (detailed in the following section). For onedimensional liquid chromatography (1D-LC), the peptides are separated according to their hydrophobicity by a reverse phase (RP) column. This is the most commonly employed method of separation of peptides obtained from in-gel digests of 1D or 2D SDS-PAGE or by in-solution digests of low complexity samples (immunoprecipitates) (Patterson and Aebersold, 2003). Exploitation of more than one property of a peptide is a powerful chromatographic technique for protein identification in complex samples. For example, multidimensional protein identification technology (MudPIT) separates peptides first by electrostatic charge using strong cation exchange (SCX) and then by hydrophobicity by RP high-performance liquid chromatography (HPLC). Coupled to online MS/MS, MudPIT has proved to be a very successful means of identifying proteins in complex mixtures without the need for gel electrophoresis (Coppinger et al., 2004; Link et al., 1999; Washburn et al., 2001). However, there are some major limitations with this approach for the identification of membrane proteins, which is discussed in detail later in this chapter. Additionally, in 2002, Gevaert and colleagues reported a novel gel free proteomic technology called combined fractional diagonal chromatography (COFRADICTM). Central to this method is a modification reaction that alters the retention behavior of specific peptides on RP columns. COFRADICTM was initially developed for the isolation of methionine-containing peptides from a tryptic peptide mixture, prior to protein identification by MS, and applied to investigate the Escherichia coli proteome (Gevaert et al., 2002). More recently, the same group has also described a diagonal method to isolate N-terminal peptides (Gevaert et al., 2003). This reduces the complexity of the peptide sample, because each protein has one N terminus and this is represented by one peptide.

3.3. Mass spectrometry Mass spectrometry for protein identification relies on the digestion of protein samples into peptides by a sequence-specific protease such as trypsin. Trypsin specifically cleaves proteins on the carboxyl-terminal side of arginine and lysine residues. This creates peptides both in the preferred mass range for sequencing and a basic residue at the carboxyl terminus of the peptide. Peptides are much more amenable to MS analysis because proteins cannot easily be eluted from gels. In addition, the molecular weight of whole proteins is usually not sufficient information for protein identification because proteins are heterogeneous and thereby usually posses no single molecular weight. Most importantly, if the purpose is to identify proteins, sequence information is needed (except when using peptide mass fingerprinting [PMF], generated by matrix-assisted laser desorption/ionization [MALDI]).

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In addition, the mass spectrometer is most efficient at obtaining sequence information from peptides that are up to
20 residues long rather than from whole proteins. After proteins are digested, the peptides are introduced into the mass spectrometer for analysis. Mass spectrometers consist of a series of components: a means of sample introduction; an ionization mechanism; a section to separate, select, and fragment peptides; a mass analyzer (with a vacuum system), and an ion detector. Mass spectrometers differ vis-a`-vis their mechanism of ionization and type of mass analyzer. In electrospray ionization (ESI), peptides enter the mass spectrometer by elution from a microscale, capillary, high-performance liquid chromotography (HPLC) column that is directly coupled to, or is ‘‘on-line’’ with the mass spectrometer. When a peptide arrives at the end of the column, it flows through a needle. At the needle tip, the liquid is vaporized and the peptide is subsequently ionized by the action of a strong electrical potential. This ESI process was developed for use in biological MS (Fenn et al., 1989). For MALDI, the peptide is mixed with a large excess of ultraviolet-absorbing matrix, which is normally a low-molecular-weight aromatic acid. On irradiation with a focused laser beam of the appropriate wavelength, the excess matrix molecules sublime and transfer the embedded nonvolatile protein or peptide into the gas phase. After numerous ion— molecule collisions in the plume of ions and molecules, singly protonated peptide ions are formed, which are accelerated by electrical potentials into a mass analyzer of choice (see below) (Steen and Mann, 2004). Once the peptides are within the vacuum system, they are guided and manipulated by electrical fields that aid in determining the mass of the peptides. The three main types of mass spectrometers used in proteomics are quadrupole MS, time of flight (TOF) MS, and quadrupole ‘‘ion traps.’’ Each of these differs in how they ascertain the mass or mass/charge ratio (m/z) of the peptides. Each instrument generates a mass spectrum, which is a recording of the signal intensity of the ion at each value of the m/z scale (which has Dalton [Da] units per charge) (Steen and Mann, 2004). Once the m/z values and peak intensities are generated, the MS then obtains primary structure (sequence) information about these peptides. This is called tandem MS, as it couples two stages of MS. In tandem MS, a particular peptide ion is isolated, energy is imparted by collision with an inert gas (such as nitrogen molecules or argon or helium atoms), and this energy causes the peptide to break apart. Often referred to as collision-induced dissociation (CID), this process results in the production of a tandem MS, MS/MS, or MS2 spectrum, which can be seen as a peptide fragmentation map. The resulting fragmentation pattern is dependent on the collision energy where lowenergy CID results in dissociation of the peptide along the backbone bonds, and at high energy, fragments are generated by amino-acid side chains. Low-energy CID, which is normally used in triple quad, Q-tof, and ion-trap mass spectrometers, fragments the peptide sufficiently to

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generate a ladder of peptides from the parent peptide that are detected by the mass spectrometer to produce spectra containing mostly b- and y-ions. A b-ion occurs when the charge is retained by the amino-terminal fragment (N-terminal), and a y-ion, when it is retained by the carboxy-terminal fragment (C-terminal). Thus, the mass gaps between each fragment peptide represent amino acid changes with an exact molecular mass difference that enables sequencing (Fig. 8.1).

b-ion

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Figure 8.1 (A) Peptide fragmentation is induced by collisions with an inert gas, and bond breakage occurs mainly through the lowest energy pathways, that is, cleavage of amide bonds.This leads to the generation of b-ions when the charge state is retained by the amino terminal fragment, or y-ions when retained by the carboxy terminal fragment. (B) Simplified representation of an MS/MS spectrum showing the b-ion ladder in blue and the y-ion ladder in red. Distances between peaks on the m/z axis can be used to infer partial sequences of the peptide.

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3.4. Bioinformatics Bioinformatics involves the use of techniques from applied mathematics, informatics, statistics, and computer science to solve biological problems. A common theme in bioinformatics is the use of mathematical tools to extract useful information from the noisy data produced by high-throughput techniques such as MS for proteomics analysis. Thus, bioinformatics is a useful tool in proteomics for the generation of protein identification from complex mass spectra data. Proteins are identified by comparing the PMF generated by a MALDI MS (i.e., the tryptic peptide masses in the spectrum) with the calculated tryptic peptide masses for each protein in the database. Alternatively, a fragmentation spectrum generated by tandem MS is compared to an in silico-generated fragmentation spectrum. A scoring method is then used to rank various peptide candidates, based on similarity between theoretical and observed spectra, where positive identifications are accepted above a certain score (Resing and Ahn, 2005). Several different algorithms are used to search sequence databases with tandem MS spectra data and include X!-Tandem, PeptideSearch, Sequest, Mascot, and ProteinProspector, some of which are outlined below. Current MS/MS search-algorithm scoring functions can be classified into two categories–heuristic and probabilistic. Heuristic algorithms correlate the acquired experimental MS/MS spectrum with a theoretical spectrum and calculate a score based on the similarity between the two. These search algorithms are often based on the ‘‘shared peak count’’ (SPC) approach, which counts the number of peaks common between to the two spectra. Examples of these algorithms include X!Tandem and Sequest. These programs use mathematical methods such as a correlation analysis to assess the quality of the match. X!Tandem and Sequest can often match various spectra. Scores are often generated by a given algorithm and deemed significant can be below the threshold for identification by another algorithm. However, good identifications are never below the threshold for both algorithms. Probabilistic algorithms model to some extent the peptide fragmentation process and calculate the probability that a particular peptide sequence produced by the observed sequence by chance. Mascot is an example of such a search algorithm. This algorithm uses a statistical approach to determine the probability of peptide identification and its significance. A recent analysis of the various database searching algorithms by Kapp et al. (2005) showed that Sequest performed well in terms of sensitivity (number of correctly identified proteins), but was inferior to Mascot and X!Tandem in terms of specificity (ability to discriminate between correct and incorrect peptide hits). Inherent with the use of such algorithms for large-scale identification of proteins from tandem MS is the generation of false positives, that is, incorrectly assigned identifications. False positives arise for a number of

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reasons. The abovementioned algorithms do not discriminate between peptide ions and other types of ions that may be present in the spectra, such as chemical noise or non peptide molecules. Also, peptides are often present at a wide variety of concentrations in a sample, and peptides present at the limit of detection can produce poor-quality fragmentation. Incomplete fragmentation patterns or poor signal-to-noise ratios may lead to incorrect identification. Another factor complicating definitive identification is that sequence conservation can lead to confusing results. If the same peptide exists in multiple proteins, all the proteins will be identified. Without additional peptide data, it would be impossible to determine which protein produced the peptide that generated the tandem mass spectrum. However, most algorithms are very accurate for peptides that follow general rules of fragmentation. A subset of amino acid sequences and more highly charged peptide ions deviate from these rules. Thus, by better understanding the relationship between amino acid or peptide sequence and fragment ion intensity, it will be possible to design better models for matching spectra to sequences. Furthermore, not all peptides are detected equally in the mass spectrometer. This fact can therefore influence the protein being identified in the first place. Peptide detection is highly dependent on the ease with which the peptide can be ionized. Ionization efficiency can be thought of as the tendency of the peptide to ionize and contribute to a mass spectrum. This is influenced mainly by the inherent structural properties of the peptide, such as length, mass, amino acid composition, and various biochemical properties, such as hydrophobicity, pH under the experimental condition, and so on. With this in mind, many groups are developing methods to identify prototypic peptides (peptides that are repeatedly and consistently identified for any given protein in a complex mixture) in order to gain increased confidence in protein identification (Craig et al., 2005; Mallick et al., 2007).

4. Databases As well as using the powerful bioinformatics tools described above to generate protein and peptide information from MS experiments, proteomics requires the usage of high-quality databases containing previously characterized proteins for comparison and identification. The objective of a database search is to find an exact match, or near-exact match between peptide fragments generated by MS experiment and sequences encoded by the genome. Existing databases, such as UniProtKB/Swiss-Prot, UniProtKB/TrEMBL, and International Protein Index (IPI), share common objectives including maximization of annotation, minimization of redundancy, and integration or at least linkage to other databases.

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UniProt knowledgebase ((http://www.uniprot.org)) is a database of protein sequences with functional and sequence annotations and consists of two parts–Swiss-Prot and TrEMBL. Swiss-Prot contains fully manually annotated entries, while TrEMBL consists of computer-annotated translations of the coding sequences proposed by the submitters of the sequences incorporated in the public nucleic acid databases. This database includes protein sequences for human as well as many other model organisms, such as mouse, rat, yeast, E. coli, and so on. An advantage to this database is that if your sequence is not matched to a corresponding protein from your organism of interest, a best match is generally found for this sequence in a lower organism, which can potentially infer function of the sequence. A disadvantage is that a true match for your sequence may be missed due to a ‘‘better match’’ to a protein from a different organism. This may occur because of alternative splicing or incomplete sequence coverage of the original peptide. IPI (http://www.ebi.ac.uk/IPI/IPIhelp.html) is assembled from protein sequence information taken from the following data sources: UniProtKB/ Swiss-Prot, UniProtKB/TrEMBL, RefSeq, Ensembl, TAIR, H-InvDB, and Vega. Each of these databases contains the results of different gene predication algorithms, and experimentally determined (mRNA and protein) sequences. IPI provides a species-specific, complete, and nonredundant data set. An advantage to this human database is that all identified proteins will be human proteins, and this avoids the assigning of the sequence to a similar protein from a lower organism, as mentioned above.

5. Quantitative Proteomics Quantitative proteomics centers on both the identification and quantification of proteins within complex mixtures. It is often used to examine differential protein expression between two cellular states and involves methodologies such as ICAT, O18, and iTRAQ. In order to differentiate proteins derived from two experimental states, certain chemical modifications or heavy-isotope—labeling techniques are introduced in the early stages of sample preparation. This may occur before or during the experiment (e.g., use of isotopic amino acids in growth media), or after the experiment is complete (e.g., ICAT). Although the modifications and techniques vary, the end goal is the same. The ultimate, and to date unrealized, goal of quantitative proteomics is the identification of every protein expressed in a cell or tissue, and the determination of that protein’s abundance, state of modification, and possible involvement in multiprotein complexes. Until recently, MS was not considered to be suitable for quantitative proteomics. This was mainly due to the variability of peptide detection in

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the mass spectrometer. Peptide detection is highly dependent on the efficiency of peptide ionization, and can be confounded by ionization suppression effects often observed when analyzing complex mixtures (Staes et al., 2004). Nevertheless, MS allows relative quantification (within a dynamic range that depends on the instrument) of compounds by comparing ions that are identical, apart from the presence of different stable isotopes. This allows them to be discriminated by a known mass difference. This approach has been widely used for differential proteomics (Goshe and Smith, 2003; Kislinger and Emili, 2003; Lill, 2003). In a typical quantitative proteomics analysis, cells are grown under at least two different conditions, which would be expected to alter some protein expression levels. The proteins from each cell growth condition are labeled with a ‘‘heavy’’ or ‘‘light’’ tag ( Washburn et al., 2002), resulting in the differential modification of the peptides in each sample. The types of proteins and peptide modifications used in quantitative proteomics can generally be placed in four classes: metabolic labeling such as with N15 (Conrads et al., 2001; Oda et al., 1999), post-growth amino-acid labeling such as ICAT (Griffin et al., 2001; Gygi et al., 1999a), digestion labeling such as O18 (Yao et al., 2001), and post-digestion labeling (Goodlett et al., 2001; Munchbach et al., 2000). Each of these methods introduces ‘‘heavy’’ and ‘‘light’’ labels into a protein or peptide at distinct points in an experimental system.

5.1. Label-free approaches Currently, the two widely used but fundamentally different label-free quantification strategies are (1) spectral counting and (2) ion intensity measurements. Regarding the former, a recent study by John Yates and colleagues and previous studies by other groups (Cagney et al., 2005; Liu et al., 2004; Washburn et al., 2001, 2003) suggest that a relationship exists between the level of sampling observed (as reflected in the frequency of MS/MS events recorded for that protein in the run) for a protein in a tandem mass spectrometer and the relative abundance of that protein in a mixture. Liu et al. (2004) developed a statistical model that predicts the level of sampling expected for mixtures of specific complexity. They validated this theory by spiking a yeast-soluble protein mixture with specific quantities of six different proteins. Four mixtures were created with decreasing amounts of the six proteins, but the same quantity of total protein. Each protein mixture was independently digested using trypsin and then analyzed three times using LC/LC/MS/MS. All six proteins showed a linear correlation between spectral copy number and their abundance from 4.17 to 0.0417% of total protein content, a two-order linear dynamic range. R2 values were between 0.9967 and 0.9995 for the linear correlation between spectral count and the amount of each protein added to the yeast cell lysate. Spectral

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count refers to the total number of MS spectra identified from a protein. Their model accurately predicts that higher-abundance proteins are sampled more frequently, but that greater coverage of lower-abundance proteins can be achieved by increasing the number of experiments performed on a single sample. However, saturation in protein identifications from a complex mixture can be reached. Although very intuitive and attractive in practical terms, the spectrumcounting approach is still controversial because it does not measure any direct physical property of a peptide. It further assumes that the linearity of response is the same for every protein. However, this caveat has been taken into account by a number of groups in their attempt to refine the method. This includes an approach by Rappsilber et al. (2002), in which a protein abundance index (PAI) was calculated by dividing the number of observed peptides by the number of all possible tryptic peptides from a particular protein that are within the mass range of the instrument used. This group then extended the PAI into an exponentially modified form (emPAI), which showed a better correlation to known protein amounts (Ishihama et al., 2005). Other adaptations include using computer models to predict which peptides of a given protein are likely to be detected by the mass spectrometer, thus potentially providing more reliable means for quantification (Craig et al., 2005; Lu et al., 2007; Tang et al., 2006). In the second approach, peptide ion intensities belonging to a particular protein are measured and compared. The ion chromatograms for every peptide are extracted from an LC-MS/MS run and their mass spectrometric peak areas are integrated over the chromatographic time scale. For low-resolution mass spectra, this is typically done by creating extracted ion chromatograms (XICs) for the mass-to-charge ratios determined by each peptide (Bondarenko et al., 2002). More recently, this concept has been extended to high-resolution data to include contributions of C13 isotopes to the overall signal intensities (Ono et al., 2006). The intensity value for each peptide in a single experiment can then be compared to the respective signals in one or more other experiments to yield relative quantitative information (Bondarenko et al., 2002; Chelius and Bondarenko, 2002; Higgs et al., 2005; Wang et al., 2006; Wang et al., 2003; Wiener et al., 2004). For proteomics analysis of very complex peptide mixtures, three important experimental parameters affect the analytical accuracy of quantification by ion intensities. 1. Using a high-mass-accuracy mass spectrometer is beneficial because the influence of interfering signals of similar but distinct masses can be minimized. 2. The peptide chromatographic profile should be optimized for reproducibility to ease finding corresponding peptides between different experiments. This is not a trivial task and special software has been developed to

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align LC-runs prior to identifying corresponding peptides (Bylund et al., 2002; Jaitly et al., 2006; Strittmatter et al., 2003; Wang et al., 2007). 3. The right balance between acquisition of parent and fragment spectra has to be found. While extensive peptide sequencing by tandem MS is required to identify as many proteins as possible in complex mixtures, a robust quantitative reading by ion intensities requires multiple sampling of the chromatographic peak by survey mass spectra. Typically, multiple fragment spectra are acquired for every survey spectrum at acquisition rates ranging from 0.2 s/spectrum (ion traps) to 1 to 3 s/spectrum (Q-TOFs). However, as chromatographic peak widths are in the order of 10 to 30 s for nano-LC separations, ion traps have an inherent advantage over QTOFs, as many more MS to MS/MS cycles can be performed within the available chromatographic time. Nevertheless, even for fast sampling instruments, better quantification accuracy will inevitably mean poorer proteome coverage and vice versa. We recently described and extensively validated a novel label-free quantitative method termed the ‘‘spectral index’’ (SI), which incorporates the key MS features of abundance, spectral count, peptide number, and ms/ms ion intensity (Yu, 2008). However, unlike previous studies (Lukas et al., 2006; Silva et al., 2006), we generated our intensity values by extracting the total intensity of all detected fragment ions (ms/ms) for a specific peptide, as opposed to intensity values generated by integrated peak areas from the chromatogram. Other benefits may have been gained by using peak areas, but this method can be particularly difficult when trying to integrate poorly resolved chromatogram peaks. We clearly demonstrate the benefit to be gained from combining these features in SI over using them independently in isolation, such as with spectral count (SC). SI consistently outperformed SC regardless of the measurements used or the normalization methods applied. A major challenge still lies in the computational and statistical analysis of quantitative proteomic results ( Wong et al., 2007). For example, we previously discovered that multiple MS measurements of a sample are required for large-scale ‘‘shotgun’’ proteomic platforms to achieve statistically significant comprehensiveness in protein identifications, even when using a single analytical technique (Durr et al., 2004). However, this requirement for repetitive measurements makes comparative quantitative analysis inherently more difficult, especially when multiple analytical techniques are used with multiple replicates to significantly improve statistical comprehensiveness. For example, discovering differences in distinct biological samples based on a single shotgun measurement with any confidence becomes futile when false positive rates soar, simply because any second replicate ‘‘shotgun’’ MS measurement will identify 30 to 40% of proteins not found in a single MS measurement of an identical sample (Durr et al., 2004). This generation of partially overlapping data sets from identical samples suggests poor reproducibility of shotgun proteomic analysis, which contributes to

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the impression of proteomics data being soft and of dubious quality. The statistical edict of ‘‘absence of evidence is not evidence of absence’’ holds true for shotgun proteomic data. Hence, a higher statistical stringency reaching 95% analytical completeness is not optional, but required, for identifying meaningful protein distinctions between different biological samples; however, doing so requires 5 to 10 MS measurements of each distinct sample. Unfortunately, MS measurements contain inherent biases and variations, so that signals can frequently be corrupted by either systematically or even apparently random changes. Thus, replicate samples, regardless of the abundance feature used, will usually show variation in protein abundance, which is not likely a reflection of biological change. This highlights the need to normalize measurements in order to minimize inherent experimental biases and variability so that the ‘‘sameness’’ of the sample is preserved and the real changes in protein abundance between distinct samples can be reliably determined. Such statistical quantification becomes especially critical when single or even multiple high-throughput, shotgun proteomics methods, including replicate MS measurements for each technique are used to increase comprehensiveness and thus generate enormous volumes of data. In order for label-free quantitative proteomics to become more widely used, methods of normalizing and quantifying the data must be made available with full transparency and in a user-friendly format. We recently described such an approach, SIN, a normalized version of the SI described above, and applied it to multiple data sets generated by various MS methodologies using multiple sample types. This approach was found to be extremely effective in reducing variation and noise between replicate measurement data sets. Comparisons to other methods published in the literature highlight the superiority of the normalized SI method in reducing noise and day-to-day variations as well as controlling for different sample loads. Furthermore, application of the method to biologically distinct data sets highlighted differential protein expression between distinct samples, which we heavily validated and shows excellent correlation with Western blot analysis (Yu, 2008).

6. Application of Proteomics to the Vasculature As mentioned in the introduction, exploiting the angiogenic response in a developing tumor is a good starting point for targeted drug delivery. The tissue microenvironment surrounding the blood vessels appears to control the endothelial cell phenotype in vivo (Aird et al., 1997; Janzer and Raff, 1987; Madri and Williams, 1983; Stewart and Wiley, 1981). This idea of molecular heterogeneity of the endothelium is supported by the reported

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ability of certain cells and peptide sequences to home to specific tissues after intravenous injection (Pasqualini and Ruoslahti, 1996; Rajotte et al., 1998). Furthermore, the identification of tissue-specific markers on the surface of the endothelium may provide a means of ‘‘targeting’’ therapies to tumors in specific organs, thus circumventing the adverse effects associated with systemic drug delivery. Thus, mapping the proteins or ‘‘proteome’’ that line the endothelium of various tumors, organs, and tissues would provide us with insight into normal physiological and pathological states of the endothelium. In addition, tissue- and disease-specific biomarkers, which are inherently accessible in vivo (i.e., lining the vascular endothelium), constitute key targets for molecular medicine by directing pharmacodelivery as well as molecular and functional imaging. To reach its potential, vascular targeting requires the discovery of tissuespecific (i.e., vascular bed–specific), accessible endothelial targets. Many attempts have been made to identify such targets and develop probes (Arap et al., 1998, 2002; Hagemeier et al., 1986; Horoszewicz et al., 1987; Hughes et al., 1989; Kolonin et al., 2004; Ohizumi et al., 1998; Pasqualini and Ruoslahti, 1996; St Croix et al., 2000); thus far, without technologies for comprehensive analysis, most directed delivery in vivo has not met theoretical expectations (Schnitzer, 1998). A number of angiogenic markers have been identified, including Roundabout4 (Robo4), which is expressed at sites of active angiogenesis in tumor endothelial cells (ECs) in adults (Huminiecki et al., 2002). Delta 4, a Notch ligand located in the plasma membrane, is induced by hypoxia on the endothelium at sites of active angiogenesis, including tumor ECs (Mailhos et al., 2001). Nucleolin is a well-known nuclear protein that can be expressed on tumor cells and on the EC surface in tumor blood vessels but not generally normal vessels (Christian et al., 2003; Oh et al., 2004b). CD44 is a highly heterogeneous cell-surface receptor with specific variants highly expressed in tumor ECs as well as in other tissues (Taniguchi et al., 2000). ICAM-1 (intercellular adhesion molecule-1) is an adhesion molecule expressed normally on ECs, where it functions in leukocyte attachment and migration to inflammation sites (Smith et al., 1988). During tumor angiogenesis, cytokines produced by the tumor cells activate ECs to produce increased amounts of ICAM-1 and to shed the soluble form (sICAM-1) into the blood (Giavazzi et al., 1992). sICAM-1 stimulates EC migration, differentiation, and vessel sprouting in vitro as well as angiogenesis and tumor growth promotion in vivo (Gho, et al., 2001,1999). Most of these vascular targets are expressed by or proximal to ECs in tissue and, in fact, have been identified by studying isolated ECs and tissue sections with genetic-, ligand-, and/or antibodybased probes. More direct MS-based proteomic mapping of vascular ECs in vivo is desirable, and may, at least theoretically, be even more fruitful, but it is technically complex and difficult.

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Although it appears logical to study vascular endothelia in vivo (due to the morphological and molecular heterogeneity of endothelial cells [ECs] among tissues), it has been quite difficult to analyze ECs as they exist in their native state in tissue. ECs constitute a small percentage of cells that make up a tissue, even in highly vascularized tissue. Therefore, in order to tease out the EC molecular signal from the surrounding tissue, it becomes critical to isolate the ECs from the tissue; otherwise, the EC signal will be lost in the sea of heterogeneous cells that make up all organs and tissues. In addition, the ECs constitute a very thin layer lining the blood vessels that are dispersed throughout every organ, thereby making their microdissection or relatively pure isolation very difficult. Some of the many attempts to isolate ECs from tissues have proved successful, including the EC separation from the tissue by enzymatic digestion and sorting the released cells (Auerbach et al., 1985; Bruneel et al., 2003; Obermeyer et al., 2003; St Croix et al., 2000). This was made possible through the discovery of EC markers such as von Willebrand factor and angiotensin-converting enzyme (ACE), which allowed validation and confirmation of EC isolation (Auerbach et al., 1982; Jaffe et al., 1973). The ability to isolate and grow ECs in culture permitted many of the pathways involved in the various EC functions to be characterized in detail. However, as mentioned previously, ECs are highly plastic cells that clearly alter their phenotype according to microenvironment cues. This included the transition from growing in an in vivo to an in vitro state. EC change considerably even when rapidly removed from their native tissue environment and even more when isolated and grown in culture. Such cells de-differentiate and adapt their metabolism and protein expressions to ex vivo and cell culture conditions (Madri and Williams, 1983; Schnitzer, 1997a; Thum et al., 2000). Many distinctive characteristics found in vivo are lost, including expression of tissue-specific proteins and the usual abundance of caveolae, which decrease 30- to 100-fold in cultured ECs and even in ECs freshly and rapidly isolated from tissue (Durr et al., 2004; Oh et al., 2004b; Schnitzer, 1997b). Furthermore, comparison of the cell-surface proteome of ECs existing natively in vivo versus rat lung microvasculature ECs (RLMVECs) grown under standard cell culture conditions has revealed significant differences in expression, including proteins apparently induced by the tissue microenvironment (Durr et al., 2004). Of the 450 proteins identified in vivo, only 263 were found to be expressed in vitro (58%), suggesting modulation by the unique tissue microenvironment not reproduced in cell culture. Of the 73 known EC marker proteins, 65 were found at the EC surface in the rat lung versus only 32 in cultured RLMVECs (Durr et al., 2004). Thus, the in vivo tissue microenvironment, which cannot yet be duplicated ex vivo, appears to significantly regulate distinct protein expression in ECs. Due to this phenotypic drift that occurs when EC are grown ex vivo, one can clearly see the advantages to be gained by profiling the vascular

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endothelium as it exists in its native state in vivo. Drug development is often hampered by the inaccessibility of the tissues that harbor protein targets. By profiling the inherently accessible plasma membrane (PM) of the vasculature of the organ of interest for molecular signals indicative of disease or processes of interest such as angiogenesis, one may not only learn more about the process, but potentially uncover novel targets for therapeutic drug delivery. This concept is supported by the fact that more than two-thirds of known existing drug targets are PM proteins ( Josic and Clifton, 2007). Furthermore, the proteins expressed on the PM of these endothelial cells can provide valuable insight into the physiological state of the cell and thus the underlying tissue. In addition to providing a fundamental physical interface between the inside and outside of any cell, the PM also mediates a wide variety of basic biological functions including signal transduction, molecular transport, membrane trafficking, cell—cell interactions and even drug resistance. PM-associated proteins, especially integral membrane proteins (IMP) traversing the lipid bilayers, are key elements in mediating these biological processes. It is clear why the PM of the vasculature is being extensively targeted for biomarker discovery. The advantages to be gained by performing a comprehensive proteomic analysis on the vascular endothelium as it exits in vivo become readily apparent, especially as it offers the potential to uncover vast numbers of new targets that otherwise may not be discovered by traditional biochemical and molecular methods. Thus, in order to exploit the vascular endothelium as it exists in vivo, we developed a novel method to analyze EC surface proteins by physically isolating only the luminal aspect of the EC plasma membranes from tissues (Oh et al., 2004a). To isolate the luminal EC plasma membrane directly from tissue, the tissue is first perfused with polycationic colloidal silica nanoparticles (NPs) that selectively coat this luminal surface to increase membrane density. Then, after electrostatic cross-linking and cation quenching, tissue/cell homogenization is then used to break the cells or tissue into subcellular components. Because the NP has a density greater than any component of the cell or tissue, a series of centrifugations through high-density gradients yields silica-coated plasma membranes as a pellet. Electron microscopy shows large silica-coated membrane sheets in these pellets. Intracellular membranes or membranes of other cell types (when processing tissue) are not NP-coated, so they float in the gradient and are nearly eliminated from the pellet. This procedure can also be performed on cultured cells (Schnitzer and Oh, 1996), even as a cell monolayer where the top membrane surface opposite to the plastic can be coated and purified. Using this method, the PMs are enriched to levels greatly exceeding standard subcellular fractionation techniques (e.g., Percoll) (Oh and Schnitzer, 1999) with minimal contamination from marker proteins of various intracellular organelles (nuclei, endoplasmic reticulum, Golgi, and mitochondria), as well as non-ECs in the tissue (Schnitzer et al., 1995b), including the endoplasmic reticulum and Golgi, that are commonly present in other

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published preparations (Chang et al., 1994; Lisanti et al., 1994; Oh and Schnitzer, 1999; Smart et al., 1995). The isolated luminal EC membranes can then be subjected to proteomic analysis. Consequently, the major advantage of mapping the protein profile of isolated luminal plasma membranes of the EC is the reduction in tissue complexity and the potential to define the set of proteins in direct contact with the circulating blood, which are therefore inherently accessible to biological agents injected intravenously (Durr et al., 2004; Oh et al., 1998, 2004b; Schnitzer et al., 1995b, 1995c, 1996). In our initial proteomic survey of normal rat lung EC plasma membranes as isolated above (Durr et al., 2004), we used LC-LC-MS/MS (MudPIT) and identified 332 proteins in these luminal EC membranes. Furthermore, the endothelial plasma membranes isolated from tumors expressed multiple angiogenesis markers (Oh et al., 2004b), such as VEGF R1, VEGF R2, Tie 2, CD13, C-CAM1 (carcino-embryonic-antigen—related cell-adhesion molecule 1), endoglin, neuropilin 1, and amino peptidase-N (Ferrara, 2002; Kerbel, 2002). In addition, seven new tumor-induced vascular proteins were also identified: annexin A1, annexin A8, ephrin A5, myeloperoxidase, nucleolin, Transferrin receptor, and vitamin D—binding protein. Expression profiling showed that almost all of these proteins exist at the endothelial cell surface of at least one major organ, but at much lower levels than expressed in the tumors. One promising tumor candidate target was annexin A1 (annA1), which was found only in tumor endothelial plasma membrane. Whole-body g-scintigraphy of live, lung tumor—bearing rats injected with radiolabeled annA1 antibodies rapidly validated annA1 as a lung-tumor target readily accessible to the circulating antibody. Lung tumor—bearing rats were also subjected to radioimmunotherapy. Survival studies of these rats showed that a single injection of radio-labeled annA1 antibody produced significant remission even in advanced disease. Thus, the tumor microenvironment appeared to induce distinct protein expression on the EC surface. Despite this significant improvement in membrane protein identification over previous studies and the success of the study in terms of identifying a lung-tumor— specific target, we failed to detect several well-known EC markers, many of which are lipid-embedded proteins, as well as a muchlower-than-expected number of IMPs. We subsequently carried out an extensive analysis and comparison of other commonly used MS-based proteomics methods to determine the optimal technique for EC plasma membrane protein, especially IMP identification. These methods included SDS-PAGE protein separation coupled to RP-MS/MS using either an LCQ or LTQ mass spectrometer; and 2DLC or ‘‘MudPIT,’’ as well as a newer SDS-PAGE-2DLC method, both using LCQ mass spectrometers (Li, 2008). Because the insolubility and resistance to extraction of IMP among other possible factors might contribute to their under-representation in this and other past proteomic analyses, we introduced a prefractionation

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step, SDS-PAGE, which is quite conducive to resolving and separating IMP prior to MS analysis (Li, 2008). After SDS-PAGE separation, each gel lane was cut into 50 to 80 gel slices before in-gel tryptic digestion and the gel-extracted peptides were pooled into seven fractions (
10 gel slices each fraction) prior to analysis using 2DLC/MS/MS (strong cation exchange (SCX), and reversed-phase separation with 17-step elution directly into a LCQ MS. We performed rigorous analysis of each method in terms of sensitivity, specificity, and dynamic range, as well as the number of integral membrane proteins identified by each method. We used the normalization and quantification methods developed in our previous paper (Yu, 2008), and determined that SDS-PAGE prefractionation followed by 2DLC-MS/MS (G2DC) is the superior method, outperforming the current popular standards of 2D-LC/MS/MS without SDS-PAGE and SDS-PAGE with 1D-LC-MS/MS, in each tested criteria. In particular, we detected a five-fold enhancement in the number of integral membrane proteins identified and a seven-fold increase in number of proteins with predicted multiple transmembrane helices compared to the standard MudPIT method. A minimum 10-fold improvement in the sensitivity and dynamic range of detection was also seen with the extra SDS-PAGE separation. This gel-2DLC approach resulted in the identification of 1361 proteins from the EC plasma membrane, including 397 proteins with one or more predicted transmembrane helices (IMP), which was a dramatic increase from the 332 total proteins and 160 IMP proteins identified in our initial study (Durr et al., 2004). Our study clearly demonstrated that prefractionation by SDS-PAGE prior to 2DLC selectively enhanced IMP detection by MS/MS analysis. This work has huge implications for membrane proteomics, as hitherto these proteins were extremely difficult to isolate and detect by traditional proteomics methods. In a different approach, Mustafa et al. (2007) identified a number of angiogenesis-related proteins in glioma. The vasculature of surgically removed tissue samples of gliomas was compared with normal brain vessels using a laser capture microdissection and MALDI-FTMS approach. The hypertrophied vessel walls of glioma vasculature consist of endothelial cells, pericytes, and cells expressing smooth-muscle actin. These vessels may also contain glial tumor cells (mosaic vessels), so to eliminate proteins derived from these tumor cells, they also microdissected glial tumor tissue for comparison. Comparison of the various microdissected tissues was essential for targeting structure-specific proteins. Any peptide present in the blood vessels that was also found in the glioma tissue was eliminated from the list of differentially expressed peptides. Using this approach, the authors identified and validated the expression of fibrinogen-b chain and colligin 2. However, both the fibrinogen b chain and colligin 2 are also seen in non-neoplastic tissues in which angiogenesis takes place. Therefore, colligin 2 and

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fibronectin should be considered as participants in the process of neovascularization in general without specificity for tissue type. The characterization of the EC plasma membrane proteins identified in our two studies is ongoing work in our lab. This characterization is critical in the era of biomarker discovery and drug development as the vascular EC membrane continues to be extensively targeted due to its fundamental importance in normal cellular functions as well as pathophysiological states. It also has important implications for understanding the process that underlies normal and pathological angiogenesis.

7. Summary Elucidating the molecular topography of the EC surface in multiple tissues and tumor types constitutes the first step in gaining a better fundamental understanding of functional differences across organ and tumor systems and forms the basis for future studies into how microenvironments alter EC gene/protein expression and physiology. The information gained from such analysis should provide valuable insight into blood vessel structure, function, and formation, which has critical importance in angiogenesis research. This information is also vital for tissue engineering where there is a significant lack of understanding of the properties of ECs in the context of normal tissues. By knowing EC expression in a given organ type, researchers gain markers to discover the factors leading to a particular phenotype and thus will be able to re-create the tissue environment in culture with higher fidelity. Finally, defining this vascular proteome is expected to uncover new functions and to discover clinically significant tissue-modulated and possibly even organ-specific molecules that may be useful as targets for site-directed delivery of drugs, genes, or imaging agents. This has important implication in the study tumor angiogenesis for vascular targeting. The definition of EC molecular expression in vivo is still in its infancy. The next decade should yield very significant progress toward defining the molecular topography and diversity of endothelium in many normal and diseased tissues.

8. Protocols 8.1. Isolation and subfractionation of plasma membranes Important note: Unless otherwise specified in the protocol, all procedures should be performed at 4  C and all solutions must be kept and used at ice-cold temperature.

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8.1.1. Solutions Buffer PM: 0.25 M sucrose, 1 mM EDTA, 20 mM tricine in ddH2O pH to 7.8 with NaOH (filter through 0.45-ml bottle-top filter (can be stored at 4  C). Colloidal Silica solution: From 30% positively charged colloidal silica stock solution, dilute to 1% with MBS pH 6.0 (pH to 6.0 with NaOH if necessary and filter through 0.45-ml bottle-top filter (can be stored at 4  C). Cytosolic buffer: 25 mM potassium chloride, 2.5 mM magnesium acetate, 5 mM EGTA, 150 mM potassium acetate, 25 mM HEPES in ddH2O pH to 7.4 with KOH, and filter through 0.45-ml bottle-top filter (can be stored at 4  C). Mammalian Ringer’s solution (w/o calcium): 114 mM sodium chloride, 4.5 mM potassium chloride, 1 mM magnesium sulfate, 11 mM glucose, 1.0 mM sodium phosphate (dibasic), 25 mM sodium bicarbonate in ddH2O pH to 7.4 with HCl filter through 0.45-ml bottle-top filter (can be stored at 4  C). MES (2-[N-morpholine]ethanesulfonic acid) buffered saline (MBS): 20 mM MES, 135 mM NaCl in ddH2O pH to 6.0 with NaOH filter through 0.45-ml bottle-top filter (can be stored at 4  C). Sucrose/HEPES: 250 mM sucrose, 25 mM HEPES, 20 mM potassium chloride in ddH2O pH to 7.4 with NaOH and filter through 0.45-ml bottle-top filter (can be stored at 4  C). Nitroprusside: 20 mg/ml in ddH2O, this is 200 stock (always make fresh). Nycodenz 102% (v/w): 102 g Nycodenz in 100 ml of ddH2O. Polyacrylic acid (PAA): From 25% stock solution, dilute to 0.1% with MBS pH 6.0 (pH to 6.0 with NaOH and filter through 0.45-ml bottle-top filter (can be stored at 4  C). Protease inhibitors (200): 2 mg/ml leupeptin in ddH2O, 2 mg/ml pepstatin A in ddH2O, 10 mg/ml O-phenanthroline in ddH2O, 2 mg/ml E-64 in 50% EtOH, 2 mg/ml pefabloc in ddH2O, 4M potassium phosphate in 2% polyacrylic acid, pH 11. 8.1.2. Purification of silica-coated plasma membranes 8.1.2.1. Perfusion procedure 1. Anesthetize rats with a cocktail of ketamine (60 mg/kg) and xylazine (1.6 mg/kg). Perform a tracheotomy to ventilate the lungs using a respirator. 2. Insert the tubing from the perfusion apparatus into the pulmonary artery via the right ventricle and fasten by tying a 3-0 silk suture around the artery. Cut the left atrium to allow the flow to exit.

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3. Perfuse with Ringer’s/nitroprusside for 5 min starting at room temperature. After 1 to 1.5 min, begin lowering the temperature to
10  C by placing the stainless steel loop into an ice cold bath. Gently drip ice-cold PBS over the lung to prevent drying. The remainder of the procedure should be performed at this cool temperature. 4. Perfuse MBS for 1.5 min. 5. Perfuse 1% colloidal silica solution for 1.5 min. At this point, keep the lungs inflated by replacing the tube attached to the respirator with a syringe and inflate with 3 to 5 ml of air. 6. Flush with MBS for 1.5 min. 7. Perfuse 0.1% PAA for 1.5 min. 8. Flush lungs with 8 ml of sucrose/HEPES with protease inhibitors (1). 9. Excise lungs from the animal and keep cold by immersion in ice-cold sucrose/HEPES with protease inhibitors (1). 8.1.2.2. Processing of the lung

1. Finely mince the excised lung mince with a ‘‘new’’ razor blade on a cold aluminum block embedded in ice. 2. Add 20 ml of sucrose/HEPES with protease inhibitors and place in a type C homogenizer vessel. 3. Homogenize for 12 strokes at 1800 rpm in a cold (4  C) room. 8.1.2.3. Purification of silica-coated luminal endothelial cell plasma membrane

1. Filter the homogenate through a 53-ml Nytex filter, followed by a 30 ml Nytex filter. 2. Remove 200 ml from the filter solution, label as ‘‘homogenate,’’ and store at —20  C. Adjust the volume of the remaining solution to 20 ml with cold sucrose/HEPES with protease inhibitors. 3. Add an equal volume of 102% Nycodenz and mix (this is enough for two SW 28 tubes). Layer onto a 70 to 55% continuous Nycodenz sucrose/ HEPES gradient (form by placing 3 ml of 70, 65, 60, and 55% Nycodenz sucrose/HEPES and carefully swirl the solution holding the tube at a 45-degree angle about 5 to 10 times). 4. Top sucrose/HEPES with protease inhibitors. Spin at 15,000 rpm for 30 min at 4  C in an SW 28 rotor. Aspirate off the supernatant. Resuspend the pellet in 1 ml MBS. Add equal volume of 102% Nycodenz and mix. 5. Layer onto an 80-to-60% continuous Nycodenz gradient (form by placing 350 ml of 80, 75, 70, 65, and 60% Nycodenz and twirling tube about 5 to 10 times). Top with 20 mM KCl. Spin at 30,000 rpm for 30 min at 4  C in a SW55 rotor. Aspirate and discard the supernatant. Resuspend the pellet in 1 ml MBS and label as ‘‘P.’’ Store at —20  C.

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8.1.2.4. Isolation of silica-coated plasma membranes from cultured cells (monolayer) Protocol is written for T75 flasks; alter proportionately for larger or smaller flasks. All parts of the procedure must performed on ice or in a 4  C room.

1. Wash cell monolayer three times with MBS. 2. Overlay with 1% ice-cold colloidal silica solution and incubate for 10 min. Wash cell monolayer three times with MBS. 3. Overlay with 0.1% PAA and incubate for 10 min. Wash cell monolayer three times with MBS. 4. Add 5 ml of sucrose/HEPES with protease inhibitors (1). Scrape cells and place in 15-ml centrifuge tube. Centrifuge 1000 g for 5 min at 4  C. 5. Bring to 1 ml with cold sucrose/HEPES with protease inhibitors (1). (You can use up to 6 T75 flask per 1 ml of cold sucrose/HEPES with protease inhibitors.) 6. Place in type AA homogenizer vessel. Homogenize with corresponding grinder for 20 strokes at 1800 rpm. 7. Remove 100 ml, label as ‘‘homogenate,’’ and store at –20  C. 8. Add an equal volume of 102% Nycodenz and mix (SW55 tube). Layer onto a 70 to 55% continuous Nycodenz sucrose/HEPES gradient (form by placing 350 ml of 70, 65, 60, and 55% Nycodenz sucrose/HEPES, and carefully twirling tube as above). 9. Top with sucrose/HEPES with protease inhibitors. Spin at 30,000 rpm for 30 min at 4  C in an SW 55 rotor. Aspirate off the supernatant. Resuspend the pellet in 1 ml MBS and label as ‘‘P.’’ Store P at —20  C.

8.2. Mass spectrometric analysis of endothelial cell membrane proteins Gel-2DLC: Membrane (P) proteins (40 mg) isolated from rat lung vasculature were separated by SDS-PAGE (PAGEr gel, 8 to 16% T, 10 10 cm, Cambrex Bio Science, Inc., Rockland, ME), and visualized with colloidal coomassie blue staining (Invitrogen, Carlsbad, CA). Gel lanes were cut into 50 to 72 slices. Each slice was cut into approximately 1 mm2 pieces for ingel proteolytic digestions carried out manually or robotically (MassPrep Station II, Waters) according to manufacturer’s directions. Digested peptides were extracted from the gel slices three times with 20% ACN and 10% formic acid solution. The extracted peptide fractions were lyophilized. The dried digests were resuspended with 30 ml of buffer A, and then loaded manually into a 2D (strong cation exchange, SCX and Reverse Phase, RP), self-packed microcapillary column under a helium pressure cell with approximately 600 psi. The loaded samples were directly introduced into the mass spectrometer (LCQ DecaXP equipped with ESI nanospray ion

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source) by eluting the bound peptides with a 2DLC/MS/MS scheme controlled by Agilent 1100 HPLC quaternary pumps. Briefly, 17 salt steps (ammonium acetate) were applied stepwise as 0, 5, 7, 10, 15, 20, 25, 30, 35, 40, 45, 50, 80, 100, 250, 350, and 500 mM over 2 min to elute the bound peptides from the SCX column. Each salt step was followed by a 5 to 80% ACN gradient containing 0.1% formic acid to elute the peptides on the C18 column over a 110-min period. The flow rate was maintained at 200 to 250 nl/min by a pre-column flow splitter of a 50-mm fused silica capillary. Data acquisition was carried out in data-dependent mode. Full MS scan of data acquisition was performed at the range of 400 to 1400 m/z and one MS scan followed by three MS/MS scans on the most abundant ions. The temperature of the ion transfer tube of both mass spectrometers was set at 180  C and the spray voltage was 2.0 kv. The normalized collision energy was set at 35% for both LCQ and LTQ. A dynamic exclusion window was applied for a 3-min duration.

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C H A P T E R

N I N E

Development of Coronary Vessels Xiu Rong Dong,*,† Colin T. Maguire,*,§ San-Pin Wu,} and Mark W. Majesky*,†,‡,§ Contents 1. Introduction 1.1. Origins of coronary vessels in the proepicardium 1.2. What is the PE? 1.3. Coronary vasculogenesis in the subepicardium 1.4. Coronary artery formation and smooth muscle cell recruitment 2. Microdissection and Explant Culture of the Proepicardium 2.1. Avian embryos 2.2. Mouse Embryos 3. Isolation of Total RNA from Individual PEs for Gene Expression Studies 3.1. Isolation of total RNA from single PEs 3.2. Analysis total RNA from individual PEs by RT-PCR 4. Explant Culture of the Epicardium 5. Analysis of PE and Epicardium In Vivo by Scanning Electron Microscopy 6. Methods to Study Coronary Vessel Development In Vivo 6.1. Whole-mount immunostaining of avian embryos 6.2. Whole-mount PECAM1 immunostaining of mouse embryos 6.3. Alternate antigen-retrieval methods 6.4. Confocal microscopy 6.5. Image processing 6.6. Whole-mount immunostaining for smooth muscle–marker proteins 6.7. Whole mount b-galactosidase staining 6.8. Preparation of coronary vascular casts Acknowledgments References * { { } }

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Carolina Cardiovascular Biology Center, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina Department of Medicine, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina Department of Genetics, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina Cardiovascular Sciences Graduate Program, Baylor College of Medicine, One Baylor Plaza, Houston, Texas Department of Molecular and Cellular Biology, Baylor College of Medicine, One Baylor Plaza, Houston, Texas

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03009-7

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2008 Elsevier Inc. All rights reserved.

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Abstract This chapter summarizes experimental techniques used to study coronary vessel development from its origins in the proepicardium (PE) to the final assembled network of arteries, veins, and capillaries present in the mature heart. Methods are described for microdissection and culture of the PE and embryonic epicardial cells, isolation of total RNA from single PE primordia and analysis by RT-PCR, imaging of the epicardium and coronary vessels by whole-mount confocal microscopy and by scanning electron microscopy, and the preparation of coronary vascular corrosion casts to visualize the entire coronary artery network structure. These techniques form the basic tools to study the cellular and molecular pathways that guide development and remodeling of coronary vessels.

1. Introduction 1.1. Origins of coronary vessels in the proepicardium Coronary vessels arise from the proepicardium (PE), a primordium that contains precursors for the epicardium, coronary endothelial cells (CoECs), smooth muscle cells (CoSMCs), and interstitial fibroblasts (Mikawa and Fishman, 1992) (for review, see Majesky, 2004). Proepicardial cells arise outside the heart and extend villus-like projections that attach to the myocardium around Hamburger-Hamilton (HH) stage 17.5 in avian embryos (chick, quail) and E9.5 in the mouse (Hiruma and Hirakow, 1989; Manner et al., 2001; Viragh et al., 1993). PE cells then migrate over the surface of the heart to form an epicardial covering ( Manner, 1992; Viragh and Challice, 1981; Viragh et al., 1993). In response to signals from the myocardium, some epicardial cells undergo an epithelial to mesenchymal transition (EMT) and move into the subepicardium (Dettman et al., 1998; Gittenberger-de Groot et al., 1998; Perez-Pomares et al., 1998). From this position, coronary angioblasts form a subepicardial coronary plexus that surrounds the ventricles and encircles the aortic root (for review, see Tomanek, 2005). Capillary-like vessels then invade the wall of the aorta and make contact with the aortic lumen usually at two points within the aortic valve sinuses (Ando et al., 2004; Bogers et al., 1989; Eralp et al., 2005). These contacts initiate unidirectional blood flow through the coronary plexus, which promotes remodeling of the plexus into arteries, veins and capillaries, and stimulates recruitment of pericytes and CoSMCs.

1.2. What is the PE? In avian embryos, the PE arises from mesothelial cells lining the pericardial cavity where the sinus venosus (SV) joins the inflow tract of the heart. In mouse embryos, the PE arises from mesothelium covering the septum

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transversum beginning around E8.5. In both cases, the PE initiates development as bilateral primordia on either side of the ventral midline (Schulte et al., 2007). In mouse embryos, the bilateral primordia continue to develop as such until they fuse at the midline around E9.5, whereas in avian embryos the left side regresses while the right side continues to develop (Schulte et al., 2007). Formation of the PE is initiated by signals from the underlying liver bud that induce competent mesothelium to express PE markers and proliferate to form villus-like projections that extend into the pericardial cavity (Ishii et al., 2007). Each villus is composed of a simple squamous mesothelium covering a proteoglycan- and hyaluronic acid–rich extracellular matrix (ECM) core, which contains occasional clusters of mesenchymal cells and angioblasts embedded within ( Kalman et al., 1995; Kuhn and Liebherr, 1988; Munoz-Chapuli et al., 2002; Nahirney et al., 2003; Viragh and Challice, 1981). Together, the mesothelial and mesenchymal cells of the PE comprise the progenitor cells that are delivered to the heart to form the epicardium and coronary vessels (Dettman et al., 1998; Gittenberger-de Groot et al., 1998; Mikawa and Gourdie, 1996; Mikawa and Fishman, 1992; Perez-Pomares et al., 1998).

1.3. Coronary vasculogenesis in the subepicardium After contacting the heart, PE cells migrate as a continuous mesothelial sheet of cells in avian embryos to cover the surface of the myocardium and form the epicardium. In the mouse, aggregates of PE cells attach to the myocardium either as free-floating cell clusters or as fragments of villi that transfer to the myocardium after contact with the beating heart (Rodgers et al., 2008). This process is complete by E10.5 in the mouse embryo and HH21 in the chick embryo. The next critical step in coronary vessel formation is an epithelial to mesenchymal transition (EMT) that produces epicardiumderived mesenchymal cells (EPDCs) that enter the subepicardium and migrate into the myocardium. EPDCs are forerunners for CoSMCs, adventitial fibroblasts, and cardiac interstitial cells (Dettman et al., 1998; Landerholm et al., 1999; Mikawa and Gourdie, 1996). CoECs appear to arise from two sources. One source is from angioblasts formed in the region of the liver primordium and septum transversum, and carried to the heart by proepicardial villi (Kattan et al., 2004; Mikawa and Fishman, 1992; Poelmann et al., 1993; Vrancken Peeters et al., 1997). A second source is from the epicardium itself. A variety of studies suggest that at least some epicardial cells are initially multipotential, and that following EMT they are guided to various cell fates, including endothelium, by instructive signals present in the subepicardium or produced by myocardial cells. Coronary vasculogenesis occurs in a local environment that is rich in FGF2, FGF9, FGF16, VEGF-A, VEGF-B, and angiopoietin-1 (Lavine et al., 2005; Tomanek et al., 1999, 2006; Ward and Dumont, 2004). Genetic analysis

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suggests important roles for FGF receptor 1 (FGFR1)– and FGFR2-mediated signaling in the myocardium, which triggers a wave of sonic hedgehog (Shh) activation that is essential for VEGF-A, B, and C, and angiopoietin 2 expression during coronary vasculogenesis (Lavine et al., 2006). Furthermore, loss of function mutations for Friend of Gata-2 (FOG2) in the myocardium result in the absence of subepicardium formation and a failure of coronary vessels to form (Crispino et al., 2001; Tevosian et al., 2000).

1.4. Coronary artery formation and smooth muscle cell recruitment Around E13.5 in the mouse, or HH32 (day 7.5) in the chick, paired coronary ostia are formed by ‘‘controlled invasion of the aorta’’ by individual capillary-like vessels that surround the aortic root (Bogers et al., 1989); (Waldo et al., 1990). This remarkable process usually produces two stable connections with the aortic lumen, and these are almost always found in the opposed aortic valve sinuses that face away from the pulmonary artery. Contact with the aortic lumen initiates unidirectional blood flow through the coronary plexus. This directs a plexus remodeling process that involves the enlargement of main channels of blood flow into major distributing coronary arteries, pruning and regression of vascular channels that receive little or no flow, and the formation of coronary veins. Initiation of coronary blood flow also marks the onset of recruitment of CoSMCs and pericytes that will provide important survival and maturation signals for the endothelium, and will build a pressure-bearing artery wall. The first appearance of SMC differentiation markers in the coronary vasculature is around E16.0 in the rat at the level of the coronary stems (Ratajska et al., 2001). Smooth muscle a-actin was the first marker detected in the coronary vessel wall at E16.0, followed by SM-myosin heavy chain on E17, the 1E12 antigen (a smooth muscle–specific isoform of a-actinin) on E18, and finally smoothelin in the early postnatal period (Ratajska et al., 2001). This timing suggests that CoSMC differentiation is dependent on blood flow through the coronary vessels. CoSMC differentiation marker expression initiates in the coronary stems and proceeds in an orderly and continuous downstream sequence (Hellstrom et al., 1999; Hood and Rosenquist, 1992). An important role for PDGF signaling is suggested by a study from Tallquist et al. (2003), who used gene-targeting approaches to produce an allelic series of tyrosine to phenylalanine mutation in the intracellular domain of the PDGF receptor-b. Their analysis showed a strong correlation among the extent of CoSMC investment, the amount of PDGF-Rb expressed by CoSMCs, and the number of signal-transduction pathways that are activated by the receptor intracellular domain (Tallquist et al., 2003). A critical source of PDGFBB for CoSMC recruitment is from CoECs (Bjarnegard et al., 2004). Finally, there is a rapid expansion of the coronary vascular network shortly

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after birth triggered by rapid growth of the heart and increased demands for cardiac perfusion (Tomanek, 2005). In the mouse, capillary density increases three- to four-fold, and the number of coronary vessels that acquire a coating of SMCs increases at least 10-fold during the first 3 weeks after birth (Carmeliet et al., 1999).

2. Microdissection and Explant Culture of the Proepicardium 2.1. Avian embryos 2.1.1. Obtaining the embryo Fertilized quail (Cortunix cortunix) (Northwest Gamebirds, Kennewick, WA) or SPF chick (Charles River Laboratories, Wilmington, MA) eggs are incubated at 37
C in a moisture-controlled egg incubator (GQF Manufacturing, Inc., Savannah, GA) with rotation for 60 to 66 h. Under these conditions, the embryo should be at Hamburger-Hamilton (HH) stage 17.0 to 18.0 (Hamburger and Hamilton, 1951). In our experience, stage HH17.5 is optimal to visualize and microdissect the developing PE prior to its contact with the heart. To begin, clean the eggshell with 70% ethanol, and sterilize dissecting instruments in a 250-ml beaker containing water at a slow boil. To open the egg, use the blunt side of a pair of scissors and tap the small end of the egg to break the shell and then carefully make a radial cut in the shell and slide the yolk and egg contents out onto a 100-mm Petri dish. The embryo should be positioned on top of the yolk. Prepare a small piece of filter paper (1 cm  1 cm, size 3, 1003185, Whatman International Ltd., Maidstone, England) with a center window consisting of two overlapping holes made by a standard-sized, hand-held paper punch (7-mm diameter). Carefully position the autoclaved filter paper carrier over the embryo such that the entire embryo is visible within the central window of the paper carrier, and then lower the carrier onto the yolk surface. Using the edges of the filter paper as a guide, carefully cut the membrane covering the yolk, lift the filter paper carrying the embryo with sterile forceps, and place the embryo in cold 1  PBS, 2 antibiotic-antimycotic solution (AB/AM, Gibco/BRL) on ice until ready for further dissection. 2.1.2. Microdissection of avian PE Three steps to successful microdissection of the PE from avian embryos are identify, expose, and cut. The first step is to identify the PE in HH-stage17.5 embryos under a dissecting stereomicroscope (Leica MZ6, or equivalent). At this stage, the PE looks like a cauliflower-shaped cluster of villi located adjacent to the sinus venosus (SV) of the developing heart (Fig. 9.1).

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A Quail PE HH17

B

HT SV

PE

HH17 C

D HT

HT

PE E9.5

Mouse PE

Figure 9.1 Coronary vessels are formed from progenitor cells that reach the heart via a transient structure called the proepicardium (PE). (A) A stage-HH17 quail embryo oriented with the ventral side up.The PE (arrow) is seen posterior to the looped heart tube (HT) and overlying the junction of the sinus venosus (SV) and the inflow tract of the heart. (B) Scanning electron micrograph of an HH17 quail embryo PE oriented similarly to that shown in A. Note the irregular surface of the PE, and the specialized cell^ cell junctions with numerous and interdigitated villi (inset). (C) An E9.5 mouse embryo with the PE (arrow) located just ventral to the heart tube (HT). (D) Scanning electron micrograph of mouse PE at E9.5. Note the less well-organized structure of the mouse PE (thick arrow) compared to the quail PE shown in (B). Note also the numerous small clusters of PE cells that have attached to the surface of the myocardium (thin arrow). The inset shows the numerous filopodial-type extensions that PE clusters exhibit when they initially attach to the myocardium.

To visualize the PE, use sterile forceps and a tungsten needle microscalpel (Conrad et al., 1993; Le Douarin et al., 1996), and expose the heart field by peeling off all membranes that cover the embryo body and the heart. In most cases, the PE will be found on the left side of the embryo. To microdissect the PE, make one cut between the PE and the SV then make a second half-moon–like cut on the bottom of the ‘‘cauliflower-like’’ PE cell cluster. Making the latter cut will free the PE from the embryo. After rinsing the dissected PE in PBS, the PE is drawn into a pipette tip using a volume of about 8 ml. Then transfer the PE in the pipette tip to 0.7 ml of culture medium (M199 medium containing 30 mM D-glucose, 5 mM L-glutamine, 1.25 mM putrescine, 1 AB/AM, and 10% fetal calf serum) in a 24-well plate. Position the PE in the center of the well with the pipette tip, and make sure it settles down on the bottom of the dish before

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placing into a 5% CO2/95% air–humidified, cell-culture incubator at 37
C as described (Landerholm et al., 1999; Lu et al., 2001).

2.2. Mouse Embryos The mouse PE differs from the avian PE in at least four ways (Figs. 9.1 and 9.2). First, the mouse PE spans the entire width of the embryo at the level of the looped heart owing to the continued growth of both the right and left sides of the PE primordia. In contrast, in avian embryos the left side regresses while only the right side continues to develop (Schulte et al., 2007). Therefore, the mouse PE can be seen on both sides of the embryo (Fig. 9.2). Second, the mouse PE has generally shorter and thinner villi than the avian PE. Third, the mouse PE arises from a mesothelial cell layer covering a protruding structure called the septum transversum (ST) (Fig. 9.2). Finally, the border of the mouse PE with the embryo body is not as clear as the avian A

Mouse embryo

HT

SV

PE

E9.75 B

HT ST PE

Figure 9.2 Whole mount b-galactosidase staining of a PE-specific reporter in transgenic mice. (A, B) Histochemical staining for LacZ activity in an E9.5 mouse embryo expressing a b-galactosidase reporter gene from the Tbx18 locus. Tbx18 is highly expressed in the mesothelial cells of the PE (arrow, A and B). Note in (A) that the PE appears over the junction of the sinus venosus (SV) with the inflow tract of the heart, and that it spans the entire width of the embryo at E9.5. HT, heart tube; PE, proepicardium; ST, septum transversum; SV, sinus venosus.

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PE (Fig. 9.2), and more attention is required in its microdissection. Three gentle cuts have to be made to free the mouse PE from the embryo. The first cut is between the PE and sinus venosus, the second cut is between the PE and the hind-limb bud and the third cut is horizontal to the ST. The same procedure for picking up the avian PE in a pipette tip and transferring it to culture medium applies to the dissected mouse PE. For explant culture, the dissected avian or mouse PE is placed in prewarmed M199 culture medium supplemented with 30 mM D-glucose, 5 mM glutamine, 1.25 mM putrescine, 2 antibiotic-antimycotic solution (Gibco-BRL, Gaithersburg, MD), and 10% fetal bovine serum in 24-well tissue culture trays (Becton-Dickinson, Lincoln Park, NJ), and incubated at 37
C in a 95% air/5% CO2 tissue culture incubator as described (Lu et al., 2001).

3. Isolation of Total RNA from Individual PEs for Gene Expression Studies 3.1. Isolation of total RNA from single PEs Obtain single avian or mouse PEs using the microdissection procedures described above, rinse with PBS, and drop PE into 200 ml of solution D (4 M guanidinium isothiocyanate, 25 mM sodium citrate, 0.5% (w/v) N-laurosarcosine (Sarkosyl), 0.1M 2-mercaptoethanol) (Chomczynski and Sacchi, 2006). At this step, the PE lysate can be stored at 4
C in solution D for RNA isolation at a later date. To proceed with RNA isolation, to 200 ml of RNA sample in solution D, add sequentially 20 ml of sodium acetate (2 M, pH 4.0), 200 ml of water-saturated phenol, and 40 ml of chloroform: isoamyl alcohol (49:1), mix well and place on ice for 15 min, and then centrifuge at 4
C for 20 min at 14,400 rpm in a bench-top centrifuge (Eppendorf ). Carefully transfer the aqueous phase (200 ml) into a 1.5ml microfuge tube, add 200 ml isopropanol and 20 mg of oyster glycogen (Boehringer, Inc.) as a carrier, mix well, and leave at –20
C overnight to precipitate. On the second day, collect the RNA pellet at 14,400 rpm for 30 min at 4
C in a bench-top centrifuge. The RNA-glycogen pellet is then washed with 95% ethanol once, air dried (not too long), and resuspended in 20 ml of DEPC-treated H2O. Residual genomic DNA carried down with the pellet is removed by the addition of 0.5 ml of RNase inhibitor (20 U/ml, Applied Biosystems), and 1.0 ml of RNase-free DNase (1 U/ml, Promega) to 20 ml of RNA sample, followed by incubation for 30 min at 37
C. After DNase digestion is complete, RNA is extracted by addition of 11 ml of phenol and 11 ml of chloroform, followed by vortex mixing and centrifugation at 4
C for 5 min at 14,400 rpm. The aqueous phase is transferred to a fresh 1.5-ml microfuge tube; an equal volume of chloroform:isoamyl

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alcohol (24:1) is added to the sample and vortexed well, and then centrifuged at 4
C for 5 min at 14,400 rpm. Transfer the aqueous phase to a fresh 1.5-ml microfuge tube, add 2.5 volumes of 100% ethanol and 20 mg of oyster glycogen to the RNA sample tube, mix well, and leave at –20
C overnight to precipitate. On the third day, spin down the purified RNA pellet at 14,400 rpm at 4
C for 30 min, wash the pellet once with 95% ethanol, carefully remove residual ethanol with a pipette tip, air dry the RNA pellet, and resuspend each RNA pellet from a single PE in 20 ml of TE buffer made with DEPC-treated H2O. For long-term storage, keep the RNA sample at –80
C until use.

3.2. Analysis total RNA from individual PEs by RT-PCR In our experience, 20 ml of total RNA obtained from a single PE (1500 cells) ( Jenkins et al., 2005) are sufficient for amplification of up to 10 individual gene products. The concentration of total RNA isolated from a single PE is generally too low to be reliably measured using a UV spectrophotometer. Therefore, a standard source of total RNA with a known RNA concentration (usually whole embryo) is serially diluted in 1:10 in TE buffer, and 2 ml of serially diluted samples are reverse transcribed using oligo-d(T) primers, and then amplified for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene expression for 20, 25, and 30 cycles of PCR following the manufacturer’s protocol exactly (Applied Biosystems). At the same time, 2 ml of single PE RNA sample is also reverse transcribed and amplified for GAPDH gene expression for 20, 25, and 30 cycles of PCR, and the intensity of gene product over the linear amplification range is matched with that of the known RNA standard. Further adjustments of the dilution series of RNA standard may be needed to obtain the best estimate of RNA concentration in PE samples. The RNA sample should be stored at –80
C until use.

4. Explant Culture of the Epicardium Prior to obtaining hearts for explant culture, 12-well or 24-well culture plates are coated with 0.1% gelatin at room temperature overnight, and fresh culture medium is prepared (DMEM containing 10% fetal bovine serum and 1 AB/AM). Embryonic mouse hearts are collected at E12.5 by aseptic technique, taking particular care not to damage the outer epicardial layer when using forceps or other dissecting instruments. Place in ice-cold PBS, trim off the outflow tract vessels and both atria, place the trimmed heart in the center of a gelatin-coated well (24-well tray) containing 200 ml of culture medium described above, and gently place in cell culture

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incubator a 37
C without bumping so as not to disturb attachment of the heart to the gelatin substrate. After 24 h, heart explants are examined, wells containing epicardial cell outgrowth are marked, and the tray gently returned to the incubator for another 24 h. After 48 h, remove the heart with forceps from the wells that exhibit epicardial outgrowth, return the plate to the incubator and repeat after another 24 h. Do not allow cultures to reach the point when fibroblasts appear in the cell layer surrounding the heart. In some cases, no epicardial cell outgrowth is visible around the explanted heart until the heart is removed from the well, and small patches of epicardial cells are evident underneath where the heart contacted the gelatin substrate. If no epicardial cells are present in the well after removing the heart explant, it probably means that the heart did not make good contact with the gelatin substrate. The procedure for obtaining epicardium from explanted avian embryo hearts at HH stage 25 is the same as described above, except that culture medium is M199 containing 10% FBS and AB/AM. Remove the culture medium from epicardial cell–containing wells, rinse wells with PBS, and place the 12- or 24-well tray on ice. Add 200 ml of solution D per well, and then scrape the cell layers with a pipette tip while pipetting up and down to dissolve the cultured epicardial cell monolayer. Collect the epicardial cell lysate into a 1.5-ml microfuge tube where it can be stored at 4
C for RNA isolation at a later time. The subsequent procedure is identical to that described in Section 3.1.

5. Analysis of PE and Epicardium In Vivo by Scanning Electron Microscopy Avian embryos (stage HH17.5) or mouse embryos (E9.5) are obtained, washed in PBS, and freed from any extraembryonic membranes. Because the looped heart tube obstructs the view of the PE, embryos can be stretched in the craniocaudal axis, thereby displacing the heart more cranially and away from the underlying PE. Following dissection and stretching, embryos are immediately fixed with a mixture of 2.5% glutaraldehyde, 2% paraformaldehyde, and 15 mM sodium phosphate pH 7.4 for 24 h. Embryos are then dehydrated through an ascending series of alcohol washes (10 min each wash): (1) 30% ethanol (2), (2) 50% ethanol (1), (3) 75% ethanol (1), and (4) 100% ethanol (3). When completely dehydrated in 100% ethanol, embryos are critical-point dried in CO2 using a Balzers Union CPD 020 (BAL-TEC AG) or equivalent. Subsequent to critical-point drying, embryos are adhered to 13-mm aluminum stubs with conductive carbon adhesive pads and sputter-coated with a 60/40 Au/Pd alloy to an approximate thickness of 10 to 15 nm using

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Figure 9.3 Imaging of developing coronary vasculature. (A) Scanning electron micrograph of the epicardium in an HH25 (E5) quail heart. Note that the epicardial cells are characterized by dense microvilli ruffling at their lateral edges. (B to F) Confocal image stacks of whole-mount quail embryos stained for endothelial cell (EC) or smooth muscle cell (SMC) markers. (B) Low-power image of E7 (HH26) heart shows the early network of vessels (arrow) and clusters of QH1-positive angioblasts (arrowhead and inset). (C) Double immunostaining for ECs (QH1, red) and SMCs (SMaA, yellow) at E11 (HH37) shows a large septal artery (arrow) running along the interventricular septum. Inset: Small artery being covered by SMCs (green), while nearby vein (red) remains uncovered by SMCs (arrowhead). (D) SMCs (SMaActin, green) can be seen migrating (arrow) over growing arteries (QH1, red) at E11 (HH37). Inset shows SMCs covering the arteriole network in the myocardium. (E) Endothelial cells (green, ZO1, arrow) align parallel to the direction of blood flow, while SMCs (red, caldesmon, arrowheads) orient in a circumferential fashion. Inset shows SMCs organizing on a large septal artery costained for SM22 (green) and SMaActin (red) at E13 (HH39). (F) The right coronary artery (arrow, green, SM22) and its main branches distribute blood flow to the myocardium. Large veins (arrow, red, QH1) lack SMCs and return blood flow to the right atrium at E134 (HH39).

a Hummer X sputter coater (Anatech Ltd, Alexandria, VA) or equivalent. Sputter-coated PE and hearts are examined and imaged using a Zeiss Supra 25 FESEM (Thornwood, NY), or equivalent, set to an accelerating voltage of 5 kV (Fig. 9.1). Avian or murine embryos are obtained at the desired developmental stage, and hearts are carefully exposed by removing membranes or tissues that obscure it, taking care not to allow forceps to close around the heart, thus damaging the epicardial layer. Fixation, dehydration, critical-point drying, sputter coating, and viewing in the scanning electron microscope, use the methods described above (Section 5) (Fig. 9.3).

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6. Methods to Study Coronary Vessel Development In Vivo 6.1. Whole-mount immunostaining of avian embryos This protocol produces excellent results in avian embryos from E3 to E13, and in mouse embryos up to E12.5. On the first day, whole avian embryos or isolated hearts are fixed in 4% paraformaldehyde (PFA) for 15 min. Specimens are simultaneously permeabilized with PBT (PBS plus 0.2% Triton X-100) and blocked with 0.1% BSA and 2% normal goat serum for 1h at room temperature. Saponin (0.1%) can be substituted for 0.2% Triton X-100 for a less harsh antigen retrieval option, but then 0.1% saponin must be maintained during every subsequent step. Primary antibodies are added to a blocking buffer (0.1% BSA and 2% normal goat serum) and incubated overnight at 4
C. To identify endothelial cells in quail embryos, the QH1 monoclonal antibody (Developmental Studies Hybridoma Bank, University of Iowa) is used at a dilution of 1:50 from the supernatant. On the second day, embryos are rinsed in PBS, and incubated in a secondary block using PBT and normal goat serum at room temperature for 1 h. Matching secondary antibodies are added in 0.1% BSA and incubated for 1 hour at room temperature (1:400 goat anti-mouse IgG1 (g1)-AlexaFluor 594, Invitrogen, A21125). Nuclei are counterstained using either 1 mM DRAQ5 (Biostatus Limited, Leicestershire, UK), 5 nM Sytox Green (Invitrogen) or 5 nM TOPRO-3 (Invitrogen), rinsed with PBS, and cleared in a solution of 1:1 glycerol:PBS for at least 3 h at 4
C before imaging (Fig. 9.3).

6.2. Whole-mount PECAM1 immunostaining of mouse embryos This protocol is optimized for use in older specimens (E12.5-E18.5) when the heart tissue has become thicker and more pigmented. Isolated hearts are washed in ice-cold PBS and fixed with 4% PFA for 30 min, then permeabilized in methanol/DMSO (4:1) for 30 min at 4
C. Specimens are bleached in methanol/DMSO/30% H2O2 (4:1:1) for 1hr at room temperature. Smaller specimens can be bleached adequately for shorter durations. Specimens are subsequently permeabilized with PBT.3 (PBS plus 0.3% Triton X-100) and incubated for 1 h at room temperature. The primary antibody PECAM-1 (CD31, clone MEC13.3, BD Pharmingen, cat550247) is diluted to 1:100 with PBS, 0.1% BSA, and 2% blocking reagent (Roche, cat-1096176) and incubated overnight at 4
C. Specimens are washed with PBT.3 at 4
C for 1 h, followed by PBT.5 (PBS plus 0.5% Triton X-100) at 4
C for 1 h. After washing, a matching secondary antibody

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(donkey anti-rat IgG-Alexa 594, 1:400, Invitrogen, A21209) is added to PBS plus 0.1% BSA, and incubated for 1 h on a rocking table at room temperature. Nuclei are counterstained with DRAQ5, Sytox Green, or TOPRO-3 as described previously, and rinsed with PBS and cleared in a solution of 1:1 glycerol:PBS for 30-60 min at room temperature before imaging (Fig. 9.3).

6.3. Alternate antigen-retrieval methods The above-described protocols use Triton X-100, a nonionic surfactant, for antigen retrieval. However, some antigens are retrieved poorly, or not at all, by Triton X-100. Therefore, modified antigen retrieval methods are required. Many antibodies against components of the actin cytoskeleton produce better results if fixed with ice-cold methanol for 15 min followed by a quick 30-s permeablization step with acetone/methanol (1:3). Antibodies against SMaA (Sigma, A2547), SMgA (Seven Hills Bioreagents, cat#LMAB-b4), and calponin (Sigma, C6047) work very well using an acetone/methanol antigen–retrieval method. Except for omitting Triton X-100, every subsequent step of the above protocol remains unchanged. In order to identify which antigen retrieval method yields the best results, it is recommended that both methods described above are tested.

6.4. Confocal microscopy Fluorescent antibody staining results are best visualized using a confocal microscope, such as a Zeiss LSM5 Pascal confocal microscope (Carl Zeiss MicroImaging Inc., Thornwood, NY), an Olympus FV500 confocal microscope (Olympus America Inc., Center Valley, PA) or equivalent. Confocal microscopy gives superior spatial resolution and control over images in the Z-plane of the sample. Whole-mount stained specimens are gently laid inside of glass-bottom culture dishes (MatTek Corporation, Asland, MA) and scanned. A limitation of confocal microscopy is the shallow penetration depth of the laser light (100 mm). It is therefore important to place a glass cover-slide on top of the specimen and slowly press down to flatten the tissue sample. Images are acquired as single confocal images or confocal image stacks, which are presented as average Z-plane projections. Light sources and filter sets should be carefully matched to the fluorescent reporter group to be detected, usually conjugated to the secondary antibody.

6.5. Image processing To achieve accurate morphological representation, each confocal image scan is saved as either 8-bit TIFF files or 12-bit LSM files (depending on the confocal microscope used), and then converted into 8-bit Tiff files using

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ImageJ software (NIH, Bethesda, MD) and the LSM toolbox 4.0b plug-in. Each TIFF file is then opened in Adobe Photoshop 7.0.1 (Adobe Systems, San Jose, CA) and converted to RGB, which produces a properly pseudocolored image (original green channel images appear green, etc.). Images are combined using Photoshop layers and blending tools to produce a clear morphological image (Fig. 9.3). Alternative, and sometimes newer, methods do exist that can produce similar results.

6.6. Whole-mount immunostaining for smooth muscle–marker proteins The development of coronary smooth muscle throughout the vascular network is best studied by whole-mount immunostaining with SMC marker– specific antibodies, or by use of various transgenic mouse lines with lacZ or EGFP reporters driven by vascular SMC–specific promoter/enhancer cis regulatory elements (see below). For the whole-mount immunostaining, freshly dissected hearts are fixed and stained with a smooth muscle–marker antibody of interest. Commonly used antibodies for this purpose include SMaActin, SM22a, calponin, SMgA, caldesmon, and SM-myosin heavy chain (Owens et al., 2004). At early stages of heart development, the myocardium will express some of these smooth-muscle selective markers. Therefore, parallel immunostaining with MF20 (Developmental Studies Hybridoma Bank, University of Iowa), can be used to identify cardiac myocytes, and QH1 (avian) or PECAM1 (mouse) will identify endothelial cells. When antibodies to two or more cell types are combined during confocal imaging, patterns of smooth muscle formation in the developing coronary vascular network can be revealed in exquisite detail. When using low-power objectives (5 and 10) with the confocal microscope, it is important that confocal image stacks are presented as average Z-plane projections (Fig. 9.3).

6.7. Whole mount b-galactosidase staining To visualize coronary smooth muscle in transgenic mice, a number of different transgenic lines have been developed in which vascular SMC– specific cis regulatory elements drive expression of a bacterial b-galactosidase reporter gene in vivo (Kim et al., 1997; Li et al., 1996; Madsen et al., 1998; Mack and Owens, 1999). To visualize lacZ activity in coronary smooth muscle, whole embryos or isolated hearts are fixed in fresh 2% paraformaldehyde/0.2% glutaraldehyde (pH7.4) for 1 h on ice, washed three times with rinse buffer (100 mM sodium phosphate, 2 mM MgCl2, and 0.1% Triton X-100 at pH 7.9) at room temperature, and stained with a solution of rinse buffer containing 5 mM potassium ferricyanide, 5 mM potassium ferrocyanide, and 1 mg/ml X-gal substrate) at 37
C overnight. Postfixation is performed in 4% paraformaldehyde at 4
C overnight (Fig. 9.4).

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Figure 9.4 Whole-mount PECAM staining. Confocal image stacks of whole-mount mouse hearts stained for endothelial cells (PECAM). (A) E12.5 left ventricle free wall shows primary coronary vascular plexus prior to the onset of blood flow through the coronary vasculature (PECAM, red). (B) E18.5 left ventricle costained for ECs (PECAM, red lines, thick arrow), nuclei (TOPRO-3, red spheres), and red blood cells (green autofluorescence, thin arrow). (C) A dorsal view (10) of E18.5 heart shows branching pattern of a left main coronary artery (arrow) that penetrates into the myocardium and quickly ramifies into an arteriole bed (*)(PECAM, green; ZO1, red). (D) A higher-power image (20) of the left main coronary artery and branch vessels stained for PECAM (red) at E18.5.

6.8. Preparation of coronary vascular casts Vascular casting is an excellent method to visualize the architecture of the entire coronary vasculature down to the precapillary level. Once made, the cast is a permanent representation of the vascular network. We prepare murine embryo coronary vascular casts as described (Adamson et al., 2002) with the following minor modifications. In brief, uteri containing embryos at E18.5 are removed and immediately immersed in ice-cold PBS until further dissection. Individual embryos are perfused through the umbilical artery with heparinized lidocaine (1% lidocaine from VetTek, Inc., 1 U/ml

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A

B

800 mm Figure 9.5 Structure of coronary artery vasculature. (A) Whole mount b-galactosidae staining of coronary smooth muscle. Heart was obtained at P0 (shortly after birth) from a transgenic mouse pup expressing b-galactosidase under the control of muscle-specific regulatory elements of the SM22a promoter.The main coronary arteries are well formed at this time, and contain several layers of smooth muscle cells in the tunica media. Note also the strong expression of lacZ activity in the walls of the aorta and pulmonary trunk (top, dark blue). (B) Vascular corrosion cast of the coronary artery network at E18.5. Coronary casts were prepared as described in the text, mounted on a small piece of modeling clay made to fit onto an aluminum stub, and examined by scanning electron microscopy. Note the branching architecture of the main distributing coronary arteries, and the ability of vascular casting to capture small arterial vessels in the heart.

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heparin in 0.9% sodium chloride, Sigma) followed by 4% paraformaldehyde (pH 7.4) via a 32-gauge needle. A small incision is made in the umbilical vein to serve as a vent and to monitor completeness of perfusion. The methyl methylacrylate casting compound is prepared as follows: 1 ml monomer base, 0.3 ml catalyst, 0.02 ml promoter, a minimum quantity of pigment dye (Bateson’s #17 kit, Polysciences, Inc.) to monitor completeness of perfusion of the casting compound as empirically determined for each application, and 0.48 ml of Jet Acrylic liquid (Land Dental Mfg. Co., Ltd). Liquid casting compound is injected into embryos through the umbilical artery via a 30-gauge needle until dye-colored casting compound becomes visible in limb vessels, indicating successful perfusion with the casting compound. All perfusions on E18.5 embryos are carried out manually using a 1-ml syringe due to the small size of vessels involved and the need for careful control of perfusion pressure. Vascular casts are allowed to cure overnight at 4
C, followed by digestion of the surrounding tissues to completion with daily changes of 7 M potassium hydroxide. Digested coronary casts are mounted on a base of nonhardening modeling clay (Van Aken, Intl.) made to fit onto 13-mm aluminum SEM stubs (see above). For this purpose, the aortic cast is broken free of the systemic vascular cast, and the broken edge of the aortic cast is gently inserted into the modeling clay so that the coronary cast is elevated off the clay surface with the apex of the ventricles oriented upward. The clay base is then mounted on aluminum stubs with double-sided tape, sputter coated as described above, and visualized by SEM (Fig. 9.5). Casts of adult hearts are made by similar methods, using retroperfusion from the femoral artery.

ACKNOWLEDGMENTS The authors acknowledge Tom Landerholm, Jun Lu, Robert J. Tomanek, and Robert J. Schwartz Vicky Madden and Robert Bagnell for helpful discussions. Support for this work is from the National Institutes of Health (HL-19242, HL-07816), the American Heart Association, and the Carolina Cardiovascular Biology Center.

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Methods for Evaluating Uteroplacental Angiogenesis and Their Application Using Animal Models Pawel P. Borowicz,* Shireen Hafez,† Dale A. Redmer,* and Lawrence P. Reynolds* Contents 1. Introduction 2. Importance of the Uteroplacental Circulation to Normal Fetal Growth and Development 3. Methods to Evaluate and Quantify Placental Angiogenesis 3.1. Early pregnancy 3.2. Late pregnancy 4. What we have Learned from Studies of Uteroplacental Angiogenesis Using Animal Models 4.1. Normal pregnancy 4.2. Compromised pregnancies 5. Conclusions Acknowledgments References

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Abstract In this chapter, we present some of the compelling evidence confirming the importance of placental angiogenesis to fetal growth and development in normal and compromised pregnancies. We then describe the methodology that has been used to evaluate placental angiogenesis throughout pregnancy, including both the practical methods used to obtain reliable samples of the placental microcirculation as well as computerized methods used to analyze and reconstruct it. We then briefly describe the changes in placental angiogenesis and

* {

Center for Nutrition and Pregnancy, and Department of Animal Sciences, North Dakota State University, Fargo, North Dakota Department of Anatomy and Embryology, College of Veterinary Medicine, Alexandria University, Edfina, Elbehera, Egypt

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03010-3

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function in the models of normal and compromised pregnancy that we have used, and conclude with what we have learned from these studies and what we believe are the larger questions remaining.

1. Introduction The importance of the placenta to fetal development has been appreciated since ancient times. As Needham (1934) so clearly pointed out, ‘‘[E] ven at the most remote times children were being born, and, though the practitioners of ancient folk-medicine might confine their ideas for the most part to simple obstetrics, they yet could hardly avoid some slight speculation on the growth and formations of the embryo.’’ This speculation included the placenta, which was ‘‘an easily observed biological phenomenon, [and] was regarded as of great importance, since it was thought to be the especial seat of the external soul.’’ Perhaps based in part on the study of bird embryos (the Egyptians and probably also the Chinese developed a system for the artificial incubation of bird eggs as early as 3000 BCE) (Needham, 1934), the ancient investigators recognized the correct function of the placenta and umbilical cord. For example, Aristotle (ca. 340 BCE) in his great embryological treatise De Generatione Animalium (On the Generation of Animals) stated that ‘‘The [umbilical] vessels join on the uterus like the roots of plants and through them the embryo receives its nourishment.’’ The ancient investigators also recognized that early development of the chick and mammals was very similar. Along with the development of practical methods to incubate fertilized chicken eggs, this revelation led to the use, in vertebrates, of the developing chick as the most important model organism for embryological research from that time forward, and established the importance of animal models in obstetric research. Thus, animal models have been central to the study of the placenta and placental circulation since the earliest times, and much of our knowledge of placental vascular development continues to be derived from comparative studies in animals (Kaufmann et al., 2004; Reynolds et al., 2005b). Fortunately, the power of the comparative approach in solving complex biological problems is widely recognized. Nevertheless, almost 50 years ago Donald Barron (1959) opined, ‘‘Obstetricians . . . deprive themselves to a surprising degree of sources of information which could be available to them from animal experiments. . . [A]nimal experiments are carried out to obtain vistas and to get ideas of the mechanisms of biological operation.’’ More recently, the critical role of the continued growth of placental circulation throughout gestation has been confirmed in many experimental studies, including those of both normal and compromised pregnancies

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(‘‘compromised’’ referring to pregnancies in which fetal or placental growth, or both, are affected), including those associated with maternal or fetal genotype, increased numbers of fetuses, maternal nutrient deprivation or excess, environmental heat stress, high altitude, and others (Reynolds et al., 2005a,b, 2006). In this chapter, we first present some of the compelling evidence confirming the importance of the placental circulation to fetal growth and development in normal and compromised pregnancies. Once we have reviewed this evidence, we describe the methodology that has been used to evaluate placental angiogenesis throughout pregnancy, including both the practical methods used to obtain reliable samples of the placental microcirculation as well as the computerized methods used to analyze and reconstruct it. We then briefly describe the changes in placental angiogenesis and function in models of both normal as well as compromised pregnancy that we have used, and conclude with what we have learned from these studies and what we believe are the larger questions remaining. Throughout the chapter, we emphasize data in ruminants, for which there are numerous well-established models of compromised pregnancy, but we also discuss the relevance to other species, including humans, where applicable. Ruminant models are relatively unique, in that, macroscopically, the ruminant placenta comprises 60 to 100 discrete structures, or units, known as placentomes (Mossman, 1987; Ramsey, 1982). Each of these placentomes consists of maternal caruncular and fetal cotyledonary portions, which interdigitate and thus are in close apposition, which of course facilitates transplacental exchange of nutrients, respiratory gases, and wastes (Fig. 10.1) (Ramsey, 1982). Microscopically, the sheep placenta belongs to a classification, first used by Grosser in 1927, known as epitheliochorial, which signifies that the fetal chorion is in direct contact with the maternal uterine (endometrial or uterine mucosal) epithelium (Mossman, 1987; Ramsey, 1982). Most investigators further classify the sheep placenta as a subset of epitheliochorial, termed syndesmochorial or synepithelial, to indicate that the uterine epithelium is fused with chorionic binucleate cells to form a feto–maternal syncytium ( Wooding and Flint, 1994). Chorion refers to the outer tissue layer (ectodermal epithelium, also termed the trophectoderm or trophoblast, plus the underlying mesoderm) of the fetal chorioallantois, which is the outer fetal membrane surrounding the fetal compartment (Mossman, 1987; Ramsey, 1982). In mammals with epitheliochorial placentation (moles, manatees, whales, horses, pigs, cattle, sheep, and a few prosimians), the chorioallantois is minimally invasive, and thus the uterine epithelium remains intact during pregnancy (Mossman, 1987; Ramsey, 1982). Because of this, the ruminant placenta is an ideal model for studying placental development because the maternal and fetal portions of the placenta remain closely associated but completely intact throughout gestation, and thus one can evaluate each tissue separately (Ramsey, 1982; Reynolds and Redmer, 1995; Reynolds

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Fetal (cotyledonary) portion

Uterine wall Maternal (caruncular) portion

Figure 10.1 Schematic representation of the sheep placentome. The maternal, or caruncular, portion is represented by the white, stippled areas, and the fetal, or cotyledonary, portion is represented by the grayish areas. The vascular supply for each portion of the placentome is represented by the blackish (maternal, caruncular) or whitish (fetal, cotyledonary) vessels. (Adapted from Reynolds, L. P., Borowicz, P. P.,Vonnahme, K. A., Johnson, M. L., Grazul-Bilska, A. T., Redmer, D. A., and Caton, J. S. (2005a). Placental angiogenesis in sheep models of compromised pregnancy. J. Physiol. 565,43^58.)

et al. 2005a,b). In addition, as described in the next section, although the epitheliochorial placenta of ruminants and other ungulates is quite different anatomically from the hemochorial placenta of most mammals, functionally they appear to operate quite similarly.

2. Importance of the Uteroplacental Circulation to Normal Fetal Growth and Development The placenta’s primary role is to provide for physiological exchange between the fetal and maternal systems (Meschia, 1983; Ramsey, 1982; Reynolds and Redmer, 1995). In this context, the importance of the placental circulation to successful pregnancy is exemplified by the close relationships among fetal weight, placental size, and uterine and umbilical blood flows during normal pregnancies in many mammalian species (Reynolds et al., 2005a,b). Uterine and umbilical blood flows, which primarily represent the circulation to the maternal and fetal portions of the placenta, respectively (Mossmann, 1987; Ramsey, 1982), increase exponentially throughout gestation, essentially keeping pace with fetal growth (Magness, 1998; Reynolds and Redmer, 1995). For example, in sheep the absolute rate of uterine blood flow

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increases approximately threefold (0.4 to 1.2 l/min) throughout the last half of pregnancy (day 71 to 131, or 49 to 90% of gestation, respectively) (Meschia, 1983). Over a similar interval of gestation, uterine blood flow in cows increases 4.5-fold (2.9 to 13.2 l/min) (Reynolds et al., 1986), and that of humans increases 2.5-fold (0.33 to 0.83 l/min) (Konje et al., 2003). The continual increase in the rate of uterine blood flow also seems to be the case for other mammalian species (Meschia, 1983; Metcalfe et al., 1988; Reynolds and Redmer, 1995). Similarly, in sheep and cows umbilical (fetal placental) blood flow increases dramatically throughout pregnancy, such that it keeps pace with fetal growth throughout the last half of gestation (Reynolds et al., 2005b). Although not measured directly, in humans umbilical blood flow velocity increases and resistance decreases throughout the last half of gestation (Gadelha da Costa et al., 2005). Not only do the absolute rates of uteroplacental blood flow increase throughout pregnancy, but importantly, the proportions of the total uterine and umbilical blood flows received by the caruncular and cotyledonary tissues, respectively, increase as gestation progresses (Makowski et al., 1968a,b; Meschia, 1983; Rosenfeld et al., 1974). Other critical placental functions such as the transport of oxygen and water also keep pace with fetal growth (Meschia, 1983; Reynolds and Redmer, 1995). As reported for umbilical blood flow, oxygen uptake and water transport remain constant when expressed per unit of fetal weight (Meschia, 1983; Reynolds et al., 1986; Reynolds and Ferrell, 1987). Similarly, fetal uptake of glucose essentially keeps pace with the rate of fetal growth (Molina et al., 1991; Reynolds et al., 1986). Based on the Fick principle, placental transport capacity, also termed placental uptake is calculated as

Uptake ¼ blood flow  ½A  V; where [A – V] represents the arterio–venous concentration difference (Reynolds and Redmer, 1995). Thus, transplacental exchange could increase by increasing the rate of extraction (the A – V concentration difference) or by increasing the rate of blood flow, or both. Based on numerous studies, it seems that increased blood flow is the primary mechanism of increased transplacental exchange throughout gestation (Meschia, 1983; Metcalfe et al., 1988; Reynolds et al., 1986; Reynolds and Redmer, 1995). For example, in cattle, oxygen extraction by the gravid uterus increases only 0.4-fold, whereas uterine blood flow increases approximately 4.5-fold from mid to late gestation. Thus, increased uterine blood flow accounts for most of the five- to six-fold increase in total gravid uterine oxygen uptake. The 16-fold increase in oxygen uptake of the bovine fetus from mid to late gestation also can be accounted for primarily by the increased rate of umbilical blood flow (Reynolds et al., 1986). Similarly, in sheep, gravid uterine oxygen extraction increases approximately 0.4-fold

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from mid to late gestation, whereas uterine blood flow increases approximately threefold (Meschia, 1983). Moreover, the large increases in gravid uterine and fetal uptakes of glucose, lactate, and amino acid nitrogen from mid to late gestation in cattle seem to depend primarily on large increases in uterine and umbilical blood flows because the arterio–venous concentration differences for these nutrients remain relatively constant (Reynolds et al., 1986; Reynolds and Redmer, 1995). Thus, adequate blood flow to the placenta is critical for normal fetal growth. However, increased abundance of specific transporters and an increase in the maternal to fetal concentration gradient also seem to be important components of increased transplacental exchange, at least for substances that are diffusion limited, such as glucose (Bell et al., 1999). Nevertheless, gravid uterine and umbilical glucose uptakes, which provide for about 60% of fetal metabolic needs (Bell et al., 1999; Reynolds et al., 1986), are reduced approximately in proportion to the reduction in placental mass and blood flows in pregnancies compromised nutritionally or by environmental heat stress (Reynolds et al., 1985, 2006). Based on the concept that chronic increases in blood flow to any growing tissue depend on vascular growth, or angiogenesis, Meschia (1983) reasoned that ‘‘the large increase of blood flow to the uterus during pregnancy . . . results primarily from the formation and growth of the placental vascular bed.’’ In fact, numerous studies have indicated that angiogenesis is indeed a major component of the increase in placental blood flow throughout gestation, and establishment of functional fetal and placental vascular beds is one of the earliest events during embryonic/ placental development (Borowicz et al., 2007; Charnock-Jones et al., 2004; Kaufmann et al., 2004; Magness, 1998; Mayhew et al., 2004; Reynolds and Redmer, 1992, 1995; Reynolds et al., 2005a,b). In fact, not only does the sustained increase in gravid uterine and umbilical blood flows depend on development of the placental vascular beds, but placental growth itself depends on placental angiogenesis because tissue growth of any magnitude normally cannot occur in the absence of vascular growth (Bassingthwaighte and Goresky, 1984; Hudlicka, 1984). This dependence on vascular development results from the high metabolic demands associated with tissue growth and the limited ability of respiratory gases, nutrients, and metabolic wastes to diffuse through the extracellular compartment ( Adair et al., 1990; Bassingthwaighte and Goresky, 1984). Thus, growth and development of the vascular beds are critical components of tissue growth and function, including that of the uteroplacenta, and as mentioned, the importance of vascular development to placental function has long been recognized. However, research on placental vascular growth has comprised primarily descriptive histological studies, whereas only a handful of quantitative studies of placental angiogenesis have been reported (Reynolds et al., 2005b).

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3. Methods to Evaluate and Quantify Placental Angiogenesis Most evaluation of placental angiogenesis has used two methodologies: (1) classical histological methods involving embedding, sectioning, and staining, followed by stereological analysis (more recently using computerized image analysis) of the microcirculation, and sometimes involving perfusion fixation; and (2) vascular casting procedures utilizing perfusion with an epoxy resin, such as Mercox (Ladd Industries, Williston, VT), followed by digestion of the tissue and electron microscopic evaluation of the remaining microvascular cast.

3.1. Early pregnancy To evaluate placental, or uteroplacental, angiogenesis during early pregnancy, we have used simple, classical histological methods, with immersion fixation followed by paraffin embedding and sectioning of the fixed tissues. We have been able to use these methods because the tissues are still relatively ‘‘normal,’’ and thus, the vessels comprising the microcirculation (arterioles, capillary beds, and venules) are more easily identified than those during mid to late pregnancy, which we discuss later. These methodologies have been described in detail by Reynolds and Redmer (1992). Although we have used several more specific methods to identify the microvessels in other tissues, such as immunohistochemistry for Factor VIII or histochemistry using specific lectins (Redmer et al., 2001), these methods thus far have not worked well for the fetal placenta. Thus, we have used primarily periodic acid–Schiff’s reagent, which is a more general staining method for basement membranes, to identify the placental microcirculation (Borowicz et al., 2007; Hudlicka, 1984; Reynolds and Redmer, 1988; Reynolds and Redmer, 1992).

3.2. Late pregnancy In contrast with those during early pregnancy, the uteroplacental microcirculatory beds later in pregnancy are highly interdigitated and the microvessels, especially those of the fetal cotyledons (or villi), are much more difficult to identify, both because many of the capillaries seem to be so immature that they do not readily stain using classical microvascular markers (Factor VIII, lectins, etc.), and because they seem to collapse easily, probably because of the high water content of the placental tissues. Thus, to solve this problem, we have been forced to develop perfusion fixation techniques similar to those we have previously described ( Jablonka-Shariff et al., 1996). These placental perfusion

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fixation methods that we have developed have been described very recently by Borowicz et al. (2007) and Vonnahme et al. (2007). Briefly, for each animal, several of the placentomes near the fetus were fixed with Carnoy’s solution by perfusion of the main arterial vessel(s) supplying the caruncle or the cotyledon (i.e., by perfusion of a branch of the uterine or umbilical artery, respectively). After the initial perfusion fixation, the placentomes were perfused with Mercox, which was allowed to harden for 1 h, postfixed by immersion, embedded in paraffin, sectioned at 6 mm, and stained, again, as with tissues from early pregnancy, with hematoxylin and periodic acid–Shiff’s (H and PAS) reagent as described previously (Reynolds and Redmer, 1988, 1992). Photomicrographs were taken at 200 magnification using a Nikon DXM 1200 digital camera (Fryer Company, Chicago). An additional set of pictures was taken using phase contrast to evaluate the quality of perfusion. As shown in Figs. 10.2 and 10.3, this perfusion fixation methodology clearly resulted in good perfusion of the placental capillary beds, resulting in readily visible capillaries. Two criteria indicate that the size of these capillaries was not affected by the perfusion: first, in all of the histological sections, the capillaries appeared to be intact (see Fig. 10.2); and second, the size of the capillaries, as reflected by their average diameter, was within the range reported for a variety of tissues (5 to 11 mm) (Borowicz et al., 2007; Vonnahme et al., 2007; Wiedeman, 1984). Representative micrographs of stained histological sections from tissues perfused via the caruncular or cotyledonary arteries are presented in Fig. 10.3. Vascularity was then determined by image analysis (Image-Pro Plus, version 5.0, Media Cybernetics, Houston, TX). For each ewe, 20 areas (64,326 mm2 each) per placentome were analyzed, with the following parameters determined for each section and for each placental tissue type (i.e., for both caruncular and cotyledonary tissues): tissue area, shrinkage area (effect of fixation that was subtracted from the tissue area), crosssectional capillary area density (CAD) (total capillary area as a proportion of tissue area), capillary number density (CND) (total number of capillaries per unit of tissue area), and capillary surface density (CSD) (total capillary circumference per unit of tissue area). Although this latter parameter, CSD, actually represents the circumference of the capillary cross-sections, it is nevertheless proportional to their surface area (Borowicz et al., 2007; Reynolds et al., 2005b). To provide a measure of average capillary size, we also calculated the average cross-sectional area per capillary (APC) for caruncular and cotyledonary by dividing the CAD by the CND. As mentioned previously, the stereological analysis has recently been supported by using placental perfusion fixation procedures along with vascular casting and electron microscopy to extend our observations concerning the architecture of the placental microvasculature.

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A

B

Figure 10.2 (A) Brightfield and (B) phase-contrast photomicrographs of the same histological section of Mercox-perfused sheep cotyledon (villus) from day 140 of gestation. In (B), note how the luminescent plastic resin (Mercox), which fills the capillaries, corresponds with the capillary vessels seen under brightfield in (A). Histochemical staining with H and PAS, 400 magnification. (Adapted from Borowicz, P. P., Arnold, D. R., Johnson, M. L., Grazul-Bilska, A. T., Redmer, D. A., and Reynolds, L. P. (2007). Placental growth throughout the last two-thirds of pregnancy in sheep: Vascular development and angiogenic factor expression. Biol. Reprod. 76, 259^267. Similar data are presented for cow placenta by Vonnahme, K. A., Zhu, M. J., Borowicz, P. P., Geary, T.W., Hess, B.W., Reynolds, L. P., Caton, J. S., Means,W. J., and Ford, S. P. (2007). Effect of early gestational undernutrition on angiogenic factor expression and vascularity in the bovine placentome J. Anim. Sci. 85, 2464^2472.)

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Caruncular

Cotyledonary

Figure 10.3 Photomicrographs of Mercox-perfused caruncular (left) and cotyledonary (right) histological sections stained histochemically with H and PAS, from day 140 of gestation. Note how the fixed-perfused capillaries stay open in both the caruncular (arrowheads) and cotyledonary (arrows) sections. 400 magnification. (Adapted from Borowicz, P. P., Arnold, D. R., Johnson, M. L., Grazul-Bilska, A. T., Redmer, D. A., and Reynolds, L. P. (2007). Placental growth throughout the last two-thirds of pregnancy in sheep: Vascular development and angiogenic factor expression. Biol. Reprod. 76, 259^267. Similar data are presented for cow placenta byVonnahme, K. A., Zhu, M. J., Borowicz, P. P., Geary,T.W., Hess, B.W., Reynolds, L. P., Caton, J. S., Means,W. J., and Ford, S. P. (2007). Effect of early gestational undernutrition on angiogenic factor expression and vascularity in the bovine placentome. J. Anim. Sci. 85, 2464^2472.)

4. What we have Learned from Studies of Uteroplacental Angiogenesis Using Animal Models 4.1. Normal pregnancy As shown in Fig. 10.4, total capillary volume of the placenta begins to increase dramatically after day 18 of pregnancy. This growth of the placental microvascular beds continues throughout pregnancy (Fig. 10.5). The quantitative methods we described briefly have allowed us to develop an empirical model of placental microvascular development (Fig. 10.6). In this empirical model, the caruncular (maternal placental) capillary beds grow primarily via an increase in capillary diameter, with only small increases in

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Capillary volume (ml x 100)

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18 24 Day after mating

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Figure 10.4 Total capillary vascular volume from day12 through 30 of early pregnancy in sheep. (Adapted from data in Reynolds, L. P., and Redmer, D. A. (1992). Growth and microvascular development of the uterus during early pregnancy in ewes. Biol. Reprod. 47, 698^708.)

capillary number or surface densities, resulting in a modest 3.3-fold increase in CAD. In contrast, the cotyledonary (fetal villous) capillary beds grow primarily by branching, resulting in a large, 12.3-fold increase in CND, accompanied by a decrease in capillary size. The relatively large, sixfold increase in capillary area and surface densities of the fetal villi (Fig. 10.5) can be explained by this branching pattern of growth. Our empirical model of angiogenesis in the caruncles and cotyledons is corroborated by studies of the physiological and anatomical constraints on morphology of vascular beds (Reynolds et al., 2005c). For example, according to ‘‘Murray’s Law,’’ the diameter (D) of a parent vessel is equal to the sum of the cube roots of the diameters of its branches; that is, (Dparent)1/3 ¼ (D branch 1)1/3 þ (Dbranch 2)1/3 þ . . . þ (Dbranch n)1/3 (Hutchins et al., 1976; Murray, 1926; West et al., 1997). Assuming twofold branching (i.e., two branches per parent vessel, or a bifurcated system) and using Murray’s law, we determined that the number of generations of capillary branches required to go from the smallest caruncular and cotyledonary arterioles (mean D ¼ 80 mm) to the average diameter of caruncular and cotyledonary capillaries at day 140 of gestation was 10 for the maternal caruncles and 15 for the fetal cotyledons, which would result in 25- or 32-fold more branches in the cotyledons compared with the caruncles (Borowicz et al., 2007; Reynolds et al., 2005c; Vonnahme et al., 2007). Thus, our empirical model supports the conclusion that the fetal placental capillary beds are much more highly branched than those of the maternal placenta. This difference in the pattern of growth between the fetal and maternal placental capillary beds is associated with dramatic differences in the microscopic anatomy and, most likely, in physiological function. As shown in

COT CAD = 0.838e0.0209 day of gestation R2 = 0.81; p < 0.0001 CAR CAD = 5.336e0.012 day of gestation R2 = 0.57; p < 0.0002

COT CND = 0.270e0.0292 day of gestation R2 = 0.90; p < 0.001 CAR CND = 4.484e0.0037 day of gestation R2 = 0.15; p < 0.02

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Day of gestation COT CSD = 1760.64e0.02004 day of gestation R2 = 0.91; p < 0.001 COT CSD = 608.154 + 4.958 day of gestation R2 = 0.13; p < 0.01

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COT APC = 2.45e(−178.586) day of gestation + 15087 R2 = 0.62; p < 0.0001 CAR APC = 9646.59 x 249.35 day of gestation R2 = 0.62; p < 0.0001

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Fetal (cotyledonary) vascularity: ↑↑ Capillary area density (6.2 X) ↑↑ Capillary no. density (12.3 X) ↑↑Capillary surface density (6.0 X)

Maternal (caruncular) vascularity: ↑ capillary area density (3.3 X) ↑ capillary no. density (1.5 X) ↑ capillary surface density (1.7 X)

Day 50

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Figure 10.6 Empirical model of angiogenesis in the maternal caruncular and fetal cotyledonary portions of the sheep placenta throughout the last two-thirds of gestation. In this model, the caruncular capillary beds grow primarily by increased capillary size, with only small increases in capillary number or surface densities, resulting in a modest increase in capillary area density. In contrast, the cotyledonary capillary beds grow primarily by branching, resulting in a large increase in capillary number density accompanied by a decrease in capillary size. The relatively large increase in capillary area and surface densities of the fetal cotyledons can be explained by this branching pattern of growth. As discussed in the text, this empirical model of angiogenesis is corroborated by studies of physiological and anatomical constraints on morphology of vascular beds. (Adapted from Borowicz, P. P., Arnold, D. R., Johnson, M. L., Grazul-Bilska, A.T., Redmer, D. A., and Reynolds, L. P. (2007). Placental growth throughout the last two-thirds of pregnancy in sheep: Vascular development and angiogenic factor expression. Biol. Reprod. 76, 259^267.)

Fig. 10.7, from very early in gestation the maternal placental capillary bed begins to form a ‘‘capillary plexus,’’ initially at the surface of the uterine lumen where the caruncle contacts the fetal chorioallantoic (compare nonpregnant caruncle with a caruncle on day 30 and 50 of gestation) (Fig. 10.7). By the beginning of the third trimester of pregnancy (about day 90 to 100 of gestation in sheep), the maternal caruncular tissue is composed primarily of these large ‘‘capillaries,’’ many exhibiting diameters Figure 10.5 Regressions of vascularity measures for caruncular (CAR, maternal) and cotyledonary (COT, fetal villus) placental tissues throughout the last two-thirds of gestation in sheep. CAD, capillary area density (%) (total capillary area as a proportion of tissue area); CND, capillary number density (total number of capillaries per tissue area); CSD, capillary surface density in micrometers (total capillary circumference per tissue area); APC, area per capillary in square micrometers (average cross-sectional area per capillary). (Adapted from Borowicz, P. P., Arnold, D. R., Johnson, M. L., Grazul-Bilska, A. T., Redmer, D. A., and Reynolds, L. P. (2007). Placental growth throughout the last two-thirds of pregnancy in sheep:Vascular development and angiogenic factor expression. Biol. Reprod. 76, 259^267.)

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Caruncle–schematic showing actual size (left) and enlargement of area in box (right)

Caruncle– actual histological section

Nonpregnant Day 30

Day 50

Day 30 and Day 50

Day 30 and 50

Day 130

Figure 10.7 Developmental changes in the architecture of the maternal placental (caruncular) microvasculature throughout gestation in sheep.The left portion of each panel shows a schematic of a caruncle at actual size (left) and then an enlarged schematic of the area enclosed by the box (right).The right portion of each panel shows a micrograph of an actual histological section; note that all micrographs are at the same magnification.

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of 20 mm (Fig. 10.7) (Borowicz et al., 2007; Reynolds et al., 2005c). In contrast, the diameter of fetal capillaries decreases throughout gestation, resulting in an average capillary diameter of 4.5 mm by the end of gestation (Fig. 10.7) (Borowicz et al., 2007; Reynolds et al., 2005c). In addition, our recent electron miscroscopic evaluation of vascular casts has confirmed this dramatic difference in microvascular architecture between the fetal and maternal placental tissues (Fig. 10.8). The microvascular architecture of the maternal placental compartment; that is, being made up almost exclusively of very large capillaries, dictates a low-velocity, ‘‘irrigation-flow’’ or slowly percolating type of system, which is designed primarily as a delivery (and, conversely, a waste-removal) system. Whether low velocity in the maternal placental capillary beds also reflects increased blood transit time is not known, and would depend on the microvascular architecture, including not only capillary size but also capillary lengths and the degree of branching. In contrast, the microvascular architecture of the fetal placenta, which is highly branched and composed primarily of abundant small capillaries, is designed as a high-velocity, rapidtransport system (Borowicz et al., 2007; Reynolds et al., 2005c). This suggestion agrees with recent functional observations. For example, resistance across the cotyledonary microvascular bed in an in vitro perfusion system is twofold greater than that of the caruncular microvascular bed at physiological flow rates throughout the last half of gestation (Vonnahme et al., 2004a,b). In addition, umbilical blood flow increases at two to three times the relative rate (proportional or percentage increase per day) as that of uterine blood flow during the last two-thirds of gestation (Reynolds and Redmer, 1995; Reynolds et al., 2005a,b, 2006). Moreover, because of increased branching and thus larger numbers of capillaries per unit of tissue Figure 10.7 (Continued). Panel (A) represents the caruncular microvascular architecture in a nonpregnant animal. Note that even in the nonpregnant state, the caruncle is highly vascular and many capillaries can be seen in both longitudinal and cross-section. Panel (B) shows the microvascular architecture of the caruncle at days 30 and 50 of gestation. Day 30 represents early formation of the placentome, and day 50 represents the end of the first trimester of pregnancy. Note the formation of a subepithelial capillary plexus on day 30 and maintenance of a similar microvascular architecture on day 50. Fetal membranes, which overly the caruncle but are as yet devoid of recognizable fetal cotyledons, or villi, are shown in the upper right of the day-30 micrograph and the upper left of the day-50 micrograph. Panel (C) represents a caruncle on day 130 of gestation (i.e., about 2 weeks before parturition). Note the formation of an extensive microvascular tree, represented in cross-section by large capillaries, many up to 20 mm in diameter, surrounded by a well-developed basement membrane and little other tissue. In fact, the maternal caruncular areas are easily identified by their heavy basement membranes, which are stained intensely pink. These maternal caruncular areas are surrounded by fetal cotyledonary tissue (lighter pinkish staining). (Adapted from Reynolds, L. P., Biondini, M. E., Borowicz, P. P., Grazul-Bilska, A.T.,Vonnahme, K. A., Caton, J. S., and Redmer, D. A. (2005c). Functional significance of developmental changes in placental microvascular architecture:The sheep as a model. Endothelium 12,11^19.)

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Figure 10.8 Electron photomicrographs of microvascular casts obtained after perfusion of the (A) maternal caruncular artery or (B) fetal cotyledonary artery of the sheep placentome (day 90 of gestation) with Mercox, followed by tissue digestion; both

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(Fig. 10.6), the surface area available for exchange is greater in fetal cotyledonary villi compared with maternal caruncles during the last trimester (Reynolds et al., 2005a,b). Additionally, the thickness of the barrier between the fetal and maternal capillaries also may be reduced throughout gestation (Faber and Thornburg, 1983; Longo, 1987). Taken together, these observations help to explain why the proportion of the nutrients and oxygen taken up by the gravid uterus that is transported to the fetus increases two- to four-fold from mid to late gestation, essentially keeping pace with the rate of fetal growth (Borowicz et al., 2007; Reynolds and Redmer, 1995; Reynolds et al., 2005c). Thus, although the placental microvascular architecture of sheep is ideally suited for both nutrient delivery, on the maternal side, and nutrient uptake and transport, on the fetal side, this arrangement does not appear to be unique to sheep. For example, we have listed several of the striking similarities in placental function between the sheep and the cow, which is another placentomal, or cotyledonary, mammal. The capillary plexus that forms on the maternal portion of the placenta early in pregnancy also has been observed in pigs (Assheton 1906; King et al., 1982), although to our knowledge this has not been examined later in pregnancy. In primates, including humans, the chorioallantois is so invasive that a portion of the maternal endometrium is eroded, and the fetal villi are bathed in maternal blood (Ramsey, 1982). The arrangement of the primate placenta thus represents the ultimate in a low-velocity, irrigation-flow or percolating system. It is interesting to note that this ‘‘hemochorial’’ type of placenta (signifying that the chorion, or outer layer of the chorioallantois, is being bathed in maternal blood) is widespread among mammals, being present not only in the vast majority of primates but also in rodents, insectivores, and bats, which together comprise 95% of the more than 4000 mammal species (Ramsey, 1982; Nowak, 1991).

4.2. Compromised pregnancies Because adequate blood flow to the placenta is critical for normal fetal growth, it is not surprising that conditions associated with reduced rates of fetal and placental growth (e.g., maternal or fetal genotype, increased numbers of fetuses, maternal nutrient deprivation or excess, environmental heat stress, high altitude) are associated with reduced rates of placental blood Figure 10.8 (Continued) micrographs were taken at 50 magnification. Note the difference in the microvasculature architecture of the maternal caruncle compared with that of the fetal cotyledon (fetal villus) as well as the dramatic difference in capillary size. These observations confirm those we have made using light microscopy (Fig. 10.3) and image analysis (Fig. 10.5) as well as the validity our empirical model of placental angiogenesis (Fig. 10.6). Hafez, Borowicz, Reynolds, and Redmer, previously unpublished.

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Table 10.1 Changes in fetal and placental weight, uterine and umbilical blood flows, and placental vascularity in various models of compromised pregnancy in sheep Day of gestation

Fetal weight

Placental weight

Uterine blood flow

Umbilical blood flow

Overfed adolescenta

130–134

#20–28%

#45%

#36%

#37%

Underfed adolescence

130

#17%

NSE

—

—

Underfed adultc

130–144

#12%

—

#17–32%

NSE

Adolescent vs. adultd Genotypee

135

#11%

#29%

—

—

—

130

#43%

#47%

—

—

"36% f

Heat-stressed adultg Multiple pregnancyh

133–135

#42%

#51%

#26%

#60%

—

Scheaffer et al., 2004; Borowicz et al., 2004 Regnault et al., 2003

140

#30%

#37%

#23%

—

#30% (total capillary volume, COT)

Grazul-Bilska et al., Christianson and Prior, 1978

Model

Vascularity

References

#31% (total capillary volume) #20% (capillary area density, CAR)b #14% (capillary area density, CAR)

Wallace et al., 2002; Redmer et al., 2004 Luther et al., 2005

Chandler et al., 1985; Leury et al., 1990; Newnham et al., 1991; Kelly et al., 1991; Arnold et al., 2000 Borowicz et al., 2004

a

High dietary Sei

135

NSE

#24%

—

—

Hypoxic (hypobaric) stress

140

NSE

—

#35% j

—

"20% (capillary number density, COT) " (capillary area density, CAR and COT)

Ward et al., 2004

Krebs et al., 1997

Uterine blood flow measured on day 130, and fetal and placental weights on day 134 of gestation. Although capillary area density (capillary area as a percent of tissue area) was reduced by 20% in maternal CAR, capillary number density (capillary number per unit of tissue area) was increased by 23 and 22% in CAR and COT, respectively. c When nutrient restriction was severe (30 to 40% of full-fed controls) and during late pregnancy (day 120 to 144), uterine blood flow was reduced by 20 to 33%. When nutrient restriction occurred during mid-pregnancy, uterine blood flow was reduced by 17% or unaffected. In addition, capillary area density only tended (p<0.09) to be reduced at day 130. d Adolescents were peripubertal (approximately 7 months of age) and adults were approximately 1 year and 7 months of age. Data are summarized for both Romanov (small-framed and low-birth-weight) and Columbia (large-framed and high-birth-weight) breeds. e Comparison of adult pregnancies in Romanov vs. Columbia breeds. f Although individual fetal weights were reduced by 43%, total fetal weight was similar or greater in Romanov vs. Columbia ewes because of the larger number of fetuses in Romanovs compared with Columbias (3 to 4 vs. 1 to 2). g Adult ewes were heat-stressed from day 80 to 120 of gestation. h Data compared were expressed as per fetus for single- versus triplet-bearing ewes (fetal and placental weights, and placental vascularity), and for single- versus twin-bearing ewes for uterine blood flow. All data are for adult ewes. i High (but subtoxic) dietary Se was fed from day 50 of gestation until necropsy. j Uterine blood flow data are for humans at 36 weeks (90%) of gestation, as no data are available for sheep. CAR, caruncle; COT, cotyledon; NSE, no significant effect. Adapted from Reynolds, L. P., Caton, J. S., Redmer, D. A., Grazul-Bilska, A. T., Vonnahme, K. A., Borowicz, P. P., Luther, J. S., Wallace, J. M., Wu, G., and Spencer, T. E. (2006). Topical review: Evidence for altered placental blood flow and vascularity in compromised pregnancies. J. Physiol. 572, 51–58. b

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flow and reduced fetal oxygen and nutrient uptakes in numerous mammalian species, including humans (reviewed in Mayhew et al., 2004; Poston, 1997; Reynolds et al., 2005a,b, 2006; Reynolds and Redmer, 1995). In fact, increased uterine vascular resistance and reduced uterine blood flow can be used as predictors of high-risk pregnancies and are associated with fetal growth retardation (Trudinger et al., 1985; North et al., 1994). Thus, the impact of factors that influence placental vascular development and function on fetal growth and development is quite striking (Reynolds and Redmer, 1995; Reynolds et al., 2005a). Moreover, observations in humans and livestock indicate that compromised fetal growth affects not only newborns but may also compromise health and productivity throughout life (Barker and Clark, 1997; Breier et al., 2001). As summarized in Table 10.1, in sheep studied during late gestation, uterine or umbilical blood flows, or both, are reduced in every model of compromised pregnancy in which they have been evaluated. These models of compromised pregnancy include overfed adolescents, underfed adolescent and adult dams, as well as environmental heat stress, hypoxic stress, and multiple fetuses. These observations agree with those in women, in which placental perfusion is reduced in pregnancies with growth-restricted fetuses (Huppertz and Peeters, 2005; Moore et al., 2004; Poston, 1997; Redmer et al., 2004; ). Although placental vascular development also is decreased in several of the models of compromised pregnancy, it is increased in others (Table 10.1). Interestingly, in two of the models in which placental vascularity is increased (high dietary Se, or hypoxic stress) (Table 10.1), there was no effect on fetal size, suggesting an adaptive placental response that preserves the fetal nutrient supply. In the other model exhibiting increased placental vascularity (Romanov vs. Columbia genotype) (Table 10.1), the animals were subject to long-term genetic selection, resulting in increased litter size. This latter case resembles that of Chinese Meishan vs. European pigs, in which the Chinese pigs exhibit increased litter size and weight associated with increased placental vascularity and VEGF expression (Biensen et al., 1998; Vonnahme and Ford, 2004; Wilson et al., 1998). Altered placental vascular development and expression of angiogenic factors in several of the sheep models of compromised pregnancy is similar to that reported in compromised pregnancies in humans, and Mayhew and colleagues (2004) suggested that most of these changes could be ‘‘driven’’ by the relative fetal hypoxia. Nevertheless, alterations in fetal growth seem to be associated with altered placental vascular development, although the functional consequences of these alterations remain to be determined (Mayhew et al., 2004; Huppertz and Peeters, 2005; Reynolds et al., 2005a,b). Altered placental growth and vascular development have been associated with altered expression of the genes for the major angiogenic factors,

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including vascular endothelial growth factor (VEGF), as well endothelial nitric oxide synthase (eNOS, or NOS3), which produces nitric oxide (NO) and thus is an important regulator of both angiogenesis and vasodilation (Borowicz et al., 2007; Redmer et al., 2005; Reynolds et al., 2005a; Reynolds and Redmer, 2001). Placental explants from preeclamptic human pregnancies exhibit increased production and release of soluble VEGF receptor-1, which binds to and inhibits the activity of VEGF ligands (Ahmad and Ahmed, 2005). Thus, placental angiogenic and vasoactive factors might serve as therapeutic targets in compromised pregnancies in humans (Ahmad and Ahmed, 2005; Godfrey, 2002). In fact, we have recently suggested that the phosphodiesterase 5 (PDE5A)–specific inhibitors, which include sildenafil, tadalafil, and vardenafil (marketed under the trade names Viagra, Cialis, and Levitra, respectively) might serve as therapeutic agents to ‘‘rescue’’ uteroplacental blood flow and vascular development in compromised pregnancies (Reynolds et al., 2006). These pharmacological agents enhance the vasodilatory action of NO by inhibiting the breakdown of cGMP, the second messenger for NO, thus causing sustained relaxation of vascular smooth muscle (Michel, 2006). Moreover, NO, produced by endothelial cells, and VEGF, produced primarily by vascular smooth muscle and capillary pericytes, may interact by stimulating each other’s expression (Ahmed and Perkins, 2000; Redmer et al., 2001; Reynolds and Redmer, 2001). Thus, impaired placental syntheses of NO may provide a unified explanation for fetal growth retardation in both underfed and overfed sheep models of fetal growth restriction (Wu et al., 2004).

5. Conclusions The methodologies described in this chapter have given us a much better understanding of placental angiogenesis. For example, we have been able to develop models of the placental microvascular growth, and have begun to appreciate the dramatic differences in architecture between the maternal and fetal placental microvasculature, which explains some of the previously described physiological differences in the function of these capillary beds. We also have identified and quantified changes in placental microvascular architecture in compromised pregnancies, which are associated with changes in placental function and growth and consequent reductions fetal growth. Lastly, we have begun to understand the factors that may regulate placental angiogenesis in normal and compromised pregnancies. The challenges that lie ahead include developing better, more robust models of placental microvascular architecture and function, developing a better understanding of the factor regulating these processes (e.g.,

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which factor[s] regulate development of the dramatically different architecture of the maternal vs. the fetal placental microvascular beds), and identifying and validating therapeutic targets that can potentially rescue placental and fetal development in compromised pregnancies. All of these will contribute substantially to the health and well-being of mothers and their offspring in both humans and animals.

ACKNOWLEDGMENTS We would like to thank the many students and colleagues who have contributed to our work throughout the last two decades, but especially Anna Grazul-Bilska, Joel Caton, Jacqueline Wallace, and Stephen Ford. We also wish to acknowledge the support from the U.S. National Institutes of Health, U.S. National Science Foundation, and the National Research Initiative, U.S. Department of Agriculture.

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North, R. A., Ferrier, C., Long, D., Townend, K., and Kincaid-Smith, P. (1994). Uterine artery Doppler flow velocity waveforms in the second trimester for the prediction of preeclampsia and fetal growth retardation. Obstet. Gynecol. 83, 378–386. Nowak, R. M. (1991). ‘‘Walker’s Mammals of the World,’’ 5th ed. Johns Hopkins University Press, Baltimore. Poston, L. (1997). The control of blood flow to the placenta. Exp. Physiol. 82, 377–387. Ramsey, E. M. (1982). ‘‘The Placenta: Human and Animal.’’ Praeger, New York. Redmer, D. A., Doraiswamy, V., Bortnem, B. J., Fisher, K., Jablonka-Shariff, A., GrazulBilska, A. T., and Reynolds, L. P. (2001). Evidence for a role of capillary pericytes in vascular growth of the developing ovine corpus luteum. Biol. Reprod. 65, 879–889. Redmer, D. A., Wallace, J. M., and Reynolds, L. P. (2004b). Effects of nutrient intake during pregnancy on fetal and placental growth and vascular development. Domestic Anim. Endocrinol. 27, 199–217. Redmer, D. A., Aitken, R. P., Milne, J. S., Reynolds, L. P., and Wallace, J. M. (2005). Influence of maternal nutrition on messenger RNA expression of placental angiogenic factors and their receptors at mid-gestation in adolescent sheep. Biol. Reprod. 72, 1004–1009. Reynolds, L. P., and Ferrell, C. L. (1987). Transplacental clearance and blood flows of bovine gravid uterus at several stages of gestation. Am. J. Physiol. 253, R735–R739. Reynolds, L. P., and Redmer, D. A. (1988). Secretion of angiogenic activity by placental tissues of cows at several stages of gestation. J. Reprod. Fertil. 83, 497–502. Reynolds, L. P., and Redmer, D. A. (1992). Growth and microvascular development of the uterus during early pregnancy in ewes. Biol. Reprod. 47, 698–708. Reynolds, L. P., and Redmer, D. A. (1995). Utero-placental vascular development and placental function. J. Anim. Sci. 73, 1839–1851. Reynolds, L. P., and Redmer, D. A. (2001). Minireview: Angiogenesis in the placenta. Biol. Reprod. 64, 1033–1040. Reynolds, L. P., Ferrell, C. L., and Robertson, D. A. (1986). Metabolism of the gravid uterus, foetus and utero-placenta at several stages of gestation in cows. J. Agri. Sci. 106, 437–444. Reynolds, L. P., Borowicz, P. P., Vonnahme, K. A., Johnson, M. L., Grazul-Bilska, A. T., Redmer, D. A., and Caton, J. S. (2005a). Placental angiogenesis in sheep models of compromised pregnancy. J. Physiol. 565, 43–58. Reynolds, L. P., Borowicz, P. P., Vonnahme, K. A., Johnson, M. L., Grazul-Bilska, A. T., Wallace, J. M., Caton, J. S., and Redmer, D. A. (2005b). Animal models of placental angiogenesis. Placenta 26, 689–708. Reynolds, L. P., Biondini, M. E., Borowicz, P. P., Grazul-Bilska, A. T., Vonnahme, K. A., Caton, J. S., and Redmer, D. A. (2005c). Functional significance of developmental changes in placental microvascular architecture: The sheep as a model. Endothelium 12, 11–19. Reynolds, L. P., Caton, J. S., Redmer, D. A., Grazul-Bilska, A. T., Vonnahme, K. A., Borowicz, P. P., Luther, J. S., Wallace, J. M., Wu, G., and Spencer, T. E. (2006). Topical review: Evidence for altered placental blood flow and vascularity in compromised pregnancies. J. Physiol. 572, 51–58. Rosenfeld, C. R., Morriss, F. H., Makowski, E. L., Meschia, G., and Battaglia, F. C. (1974). Circulatory changes in the reproductive tissues of ewes during pregnancy. Gynecol. Invest. 5, 252–268. Trudinger, B. J., Giles, W. B., and Cook, C. M. (1985). Uteroplacental blood flow velocitytime waveforms in normal and complicated pregnancy. Br. J. Obstet. Gynecol. 92, 39–45. Vonnahme, K. A., and Ford, S. P. (2004). Differential expression of the vascular endothelial growth factor–receptor system in the gravid uterus of yorkshire and Meishan pigs. Biol. Reprod. 71, 163–169.

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Vonnahme, K. A., Zhu, M. J., Borowicz, P. P., Geary, T. W., Hess, B. W., Reynolds, L. P., Caton, J. S., Means, W. J., and Ford, S. P. (2007). Effect of early gestational undernutrition on angiogenic factor expression and vascularity in the bovine placentome. J. Anim. Sci. 85, 2464–2472. Wilson, M. E., Biensen, N. J., Youngs, C. R., and Ford, S. P. (1998). Development of Meishan and Yorkshire littermate conceptuses in either a Meishan or Yorkshire uterine environment to day 90 of gestation and to term. Biol. Reprod. 58, 905–910. Wooding, F. B. P., and Flint, A. P. F. (1994). Placentation. In ‘‘Marshall’s Physiology of Reproduction,’’ (G. E. Lamming, ed.), 4th ed., Vol. 3, pp. 233–460. Chapman, and Hall, London. Wu, G., Bazer, F. W., Cudd, T. A., Meininger, C. J., and Spencer, T. E. (2004). Maternal nutrition and fetal development. J. Nutr. 134, 2169–2172. Wiedeman, M. P. (1984). Architecture [of the microcirculation]. In ‘‘Handbook of Physiology,’’ (E. Renkin and C. C. Michel, eds.), vol. IV, pp. 11–40. American Physiological Society, Bethesda, MD.

sdfsdf

C H A P T E R

E L E V E N

Intravital Microscopic Investigation of Leukocyte Interactions with the Blood Vessel Wall Klaus Ley,* Javier Mestas,* Maria K. Pospieszalska,* Prithu Sundd,* Alexander Groisman,† and Alexander Zarbock*,‡ Contents 1. Introduction 2. Transillumination Intravital Microscopy 3. Measuring Leukocyte Rolling, Adhesion, and Transmigration 3.1. Surgical considerations 3.2. Fluorescence intravital microscopy 3.3. Lymphocyte rolling and adhesion in Peyer’s patches 3.4. Intravital microscopy of large vessels 3.5. Mixed chimeric mice 4. Spinning-Disk Confocal and Two-Photon Microscopy 4.1. Autoperfused flow chamber 4.2. Microfluidic flow chambers 5. Conclusion References

256 256 259 262 264 266 267 267 268 270 272 274 274

Abstract Intravital microscopy is a method to study the microcirculation in living tissues. Transillumination, oblique reflected light illumination, continuous and stroboscopic epifluorescence microscopy can be used to visualized specific cells and molecules. Intravital microscopy is further enhanced by the advent of laser scanning.spinning disk confocal and multi-photon microscopy. Recent advances include blood-perfused flow chambers and microfluidic devises for the study of blood cell interactions with molecularly defined substrates. This chapter focuses on the application of these techniques to study leukocyte interactions with the vascular wall and molecular surfaces.

* { {

La Jolla Institute for Allergy and Immunology, La Jolla, California Department of Physics, University of California San Diego, La Jolla, California Department of Anesthesiology and Intensive Care Medicine, University of Mu¨nster, Mu¨nster, Germany

Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03011-5

#

2008 Elsevier Inc. All rights reserved.

255

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1. Introduction The observation of leukocyte adhesion and rolling under flow is possible by the technique of intravital microscopy and its variations. In this chapter, we present detailed accounts of the preparation of tissues and techniques suitable for investigating leukocyte interactions with mouse blood vessel walls and artificial substrates in vivo and ex vivo through the use of intravital microscopic techniques.

2. Transillumination Intravital Microscopy Leukocyte rolling, adhesion, and transmigration (Dutrochet, 1824; Wagner, 1839) were first described using transillumination intravital microscopy, which remains unsurpassed for ease of operation and image quality, and is almost on par with fluorescence microscopy in terms of resolution. This technique is the oldest and still the most widely used. However, transillumination intravital microscopy is severely restricted in its ability to penetrate deeper tissues. Effectively, it is limited to tissues less than 100 mm thick like the mouse and rat cremaster muscle (Baez, 1973); various other thin muscles (Engelson et al., 1985; Lindbom et al., 1982); the hamster cheek pouch (Duling, 1973); the rat, cat, or rabbit mesentery (Zweifach, 1973); and the bat wing (Wiedeman, 1973). Mouse models are the most popular, because knockout and transgenic mice are widely available, and excellent blocking antibodies for chemokines and adhesion molecules involved in leukocyte–endothelial interactions are available. Oblique reflected light illumination can be used (Mempel et al., 2003) to allow the visualization of transmigrated leukocytes in the interstitial tissue. To successfully investigate a biological question with transillumination intravital microscopy, temporal and spatial resolution must be considered. A wide field of view (about 300 mm
400 mm) combined with reasonable resolution is achieved with a 20
salt water immersion objective, such as Zeiss 20/0.5, where the first number indicates the magnification and the second the numerical aperture (NA). Better resolution is possible with a 40/ 0.75 or, especially for studies of single adherent and transmigrating leukocytes, 63/0.9. Salt water immersion is preferred because of the superior NA, which is inversely related to the smallest distance (d) separating two objects according to Abbe’s law (d ¼ 0.5 l/NA, where l is the wavelength of the light used). Immersion objectives also make it easy to keep the tissue moist and warm without producing motion artifacts from dripping superfusion solution.



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As mentioned above, the resolution improves with shorter wavelength, reaching just under 200 nm with blue light (l  400 nm) and high NA. (It is possible to determine the position of small fluorescent objects with nanometer precision. Resolution as used here means discernible objects.) However, hemoglobin has an absorption maximum in the blue range (Pries et al., 1986), which makes the blood look black when blue light is used. At longer wavelengths, hemoglobin becomes almost transparent so that leukocytes are less obstructed by red blood cells (Ley and Gaehtgens, 1991). Also, many video cameras have unfavorably low quantum efficiency in the blue range. The field of view of the microscope is 100 /magnification, because 100 is the diameter of the microscope tubes. In practice, the field of view and the resolution not only depend on the objective, but also on the video camera used. The most expensive cameras have 100 sensor chips, which produces the lowest magnification and highest field of view. More economical cameras have 2/300 , 1/200 , or even 1/400 chips, which adds ‘‘empty’’ magnification without adding resolution and limits field of view. A reducing video coupler (range 0.3
to 0.6
) may be placed between the camera and the microscope. It is important to balance resolution with field of view. Nyqvist’s theorem requires 2.3 pixels per resolvel, which means that about 12 pixels per micrometer are desirable to make full use of the objective’s power. Objectives with smaller NA require fewer pixels per micrometer and produce inferior images. However, when detecting fluorescent signals, it may be more important to boost sensitivity by maximizing the NA and the CCD chip size, allowing the image to be undersampled. In this case, maximum possible resolution is sacrificed for better light sensitivity. In our laboratory, we use both digital (Sensicam QE, Cooke Corporation, Romulus, MI) and analog (model VE-1000CD, Dage-MTI) cameras. While the advantage of digital cameras is superior resolution, most modern high-resolution digital cameras still have difficulty capturing 30 frames per second, the standard CCTV rate. This is compounded by the inability of even the fastest computers with multiple hard disks to save full-frame highresolution images (e.g., 1000
1200 pixels or more) at sufficient speed, making it necessary to capture images in random access memory (RAM) and write them to hard disk off-line. Effectively, this limits recording time at 30 fps to a few seconds. Therefore, analog cameras still have their place in intravital microscopy. Although the resolution is quite low, especially in the vertical direction (575 lines), analog cameras make it possible to record the entire experiment and later digitize the sequences to be analyzed using an inexpensive frame grabber. Alternatively, a low- or intermediate-resolution CCD camera (e.g., 512
512 or 640
480 pixels) can be streamed directly to hard disk. When studying leukocyte interactions with the vessel wall, wall shear stress and wall shear rate must be estimated, because both rolling and

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adhesion are strongly dependent on these parameters. Wall shear stress is the drag force per unit of endothelial surface area that is exerted by the flow of viscous blood. Wall shear stress is the product of wall shear rate (the radial velocity gradient) and local viscosity (Chien et al., 1984). When only one shear stress or shear rate is reported without a qualifier, this usually refers to the wall shear stress or wall shear rate. However, we have no true knowledge of the actual wall shear rate in microvessels because of the presence of an endothelial surface layer (Vink and Duling, 1996) that does not allow significant plasma flow near the endothelial plasma membrane (Damiano et al., 1996; Smith et al., 2003). In venules with diameters between 15 mm and 50 mm, which are most relevant for leukocyte adhesion in inflammation, the effective shear rate at the interface between the endothelial surface layer and the free lumen is about five times higher (Long et al., 2004) than the (traditional) Newtonian estimate reported in most papers. Newtonian shear rate would be calculated as 8 vmean/diameter, where vmean is the average blood flow velocity, which is about 60% of the commonly measured centerline velocity (Baker and Wayland, 1974). The Newtonian estimate does not apply to blood flow in microvessels, because blood is not a Newtonian fluid, and the surface layer is ignored. A better estimate of wall shear rate is 40 vmean/diameter. Blood-flow centerline velocity can be measured using a dual photodiode sensor system like that distributed by Circusoft Instrumentation (Hockessin, DE). More accurate determinations of wall shear rates can be achieved by microparticle velocimetry (Smith et al., 2003), which requires stroboscopic epifluorescence microscopy (see below). The thickness and composition of the endothelial surface layer (ESL) probably varies between vascular beds, vessel sizes, and physiologic conditions, but most reports place its thickness between 0.3 and 0.6 mm (Pries and Kuebler, 2006; Vink and Duling, 1996). This layer contains proteoglycans, glycosaminoglycans, and hyaluronic acid. In addition to its effects on shear rate at the interface between free-flowing blood and the ESL, the mechanical properties of the ESL are such that there is almost no fluid flow through this layer along the vessel surface (Damiano et al., 1996). This means that the wall shear rate on the surface of the endothelium of healthy blood vessels is nearly equal to zero (Smith et al., 2003). However, the endothelial cell can still sense the presence of shear stress, because elements of the ESL transmit forces and torques (Weinbaum et al., 2003). In physical terms, shear stress is continuous and the viscosity inside the ESL is near infinite. Although adhesion molecules are buried deep in the ESL, neutrophils and other white blood cells pass through and exit from small capillaries and compress the endothelial surface layer sufficiently to initiate adhesive interactions and start rolling (Ley, 2005). Leukocyte rolling and adhesion are commonly limited to postcapillary venules, where wall shear stress is lower than in the corresponding arterioles. However, the shear rate and shear stress ranges of arterioles and venules overlap, so that shear stress differences

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are not sufficient to explain the difference in leukocyte rolling and adhesion between arterioles and venules (Ley and Gaehtgens, 1991). Relevant adhesion molecules like E-selectin ( Jung and Ley, 1997), P-selectin (Kim and Sarelius, 2004), ICAM-1, and probably other adhesion molecules are much more highly expressed in venules than arterioles, although P-selectin expression can be induced in arterioles by TNF-a (Kunkel et al., 1997; Thorlacius et al., 1997).

3. Measuring Leukocyte Rolling, Adhesion, and Transmigration Tracking rolling leukocytes recorded by transillumination microscopy can be challenging, because the vessels are not exactly flat across the imaging plane, and therefore rolling cells move in and out of focus. In addition, cells obscure each other, and the images are cluttered with many other structures such as overlying muscle fibers. A robust algorithm (which is freely available from the author) has been developed for tracking rolling leukocytes (Acton et al., 2002). However, some programming skills are required to implement this algorithm, because it is not part of a polished commercial package. Most investigators observe one or a small number of venules for several minutes and record the number of leukocytes rolling by (rolling leukocyte flux) (Sperandio et al., 2006). With stroboscopic epifluorescence microscopy (see below), all leukocytes including those in the bloodstream can be visualized to derive the leukocyte rolling-flux fraction (Sperandio et al., 2006), which is the ratio of rolling leukocyte flux divided by total leukocyte flux (Ley and Gaehtgens, 1991). If total leukocyte flux is not measured, it can be estimated from blood flow, calculated as mean velocity times cross-sectional area of the vessel, and systemic leukocyte concentration in a systemic blood sample (Ley et al., 1995). Leukocytes roll at characteristic velocities, determined by the adhesion molecules expressed, type of leukocyte, and wall shear stress. An average velocity can be determined by measuring the distance traveled during a fixed time interval, for example, 10 seconds. This time interval must not be changed for a set of experiments, because the velocity distribution obtained strongly depends on the interval over which rolling velocity is measured (Zhao et al., 1995) (Fig. 11.1). More precise measurements can be obtained by frame-to-frame tracking, which allows analysis of pause times during which the leukocyte does not move. The instantaneous velocity values obtained are critically dependent on the time resolution of the recording device, because the actual molecular events are much faster than the frame rate of commonly used video cameras (Fig. 11.2). Many labs represent the number of rolling cells per vessel length or endothelial surface area, which is influenced by both rolling flux and rolling velocity. For example, if the cells

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Probabilty density (s/μm)

2.5 2.0 1.5 1.0 0.5 0

Time-window Δt = 20 s

1.5 1.0 0.5 0

16 s

1.5 1.0 0.5 0

12 s

1.5 1.0 0.5 0

8s

1.5 1.0 0.5 0

4s

1.5 1.0 0.5 0

1s

0.0

0.5

1.0

1.5

2.0

2.5

3.0

Rolling velocity (μm/s)

Figure 11.1 Dependency of velocity distribution of rolling neutrophils on an E-selectin-coated surface on the time window Dt.The dashed line indicates the mean of rolling velocities, which is independent of time window Dt. (With permission, from Zhao, Y., Chien, S., and Skalak, R. (1995). A stochastic model of leukocyte rolling. Biophys. J. 69,1309^1320.)

roll half as fast at the same flux, twice as many cells are seen in the vessel segment at any given time. Under inflammatory conditions or in secondary lymphoid organs, leukocytes or lymphocytes arrest, i.e., stop rolling and become adherent. Arrest can be triggered rapidly by immobilized chemokines or slowly by selectin engagement (Ley et al., 2007). The number of arrested cells can be counted and related to rolling flux, providing a percentage of rolling cells that arrest per unit of time in the observed vessel segment. Care must be taken to observe the actual arrest event, because adhesion may not be stable (see below). Many labs express adhesion by simply counting the number of adherent cells per vessel segment or per unit surface area. Such data are hard to interpret, because they represent a dynamic equilibrium between three processes: arrest (adds new adherent cells), and detachment and transendothelial migration (both remove adherent cells from the vessel). In principle, these processes could be modeled by a set of differential equations, but no such model has been developed so far. An alternative view of the leukocyte adhesion process uses a Lagrangian approach, in which each rolling leukocyte is followed through the vascular

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A

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120 100 80 60 40 20 0 100 80 60 40 20 0 60 40 20 0

High resolution

H 50

0

B

0.958

0.965

Extended vertical axis I

20

C

10 0 4

D

20 0 20 E 0 20 F 0 20 G

J

2 0

K

2 0 2

0

L

0 0

1

2 3 4 5 6 7 Time (s)

8

9 10

0

1

2 3 4 5 6 7 Time (s)

8

9 10

Figure 11.2 Dependence of instantaneous leukocyte velocities on the time resolution of the measuring device. (A) Instantaneous translational velocity for a neutrophil rolling in a flow chamber on 150 molecules/mm2 of P-selectin at a wall shear rate of 50/s.The data were obtained by computer simulation using published values for microvilli, adhesion molecule numbers, and bond and cell properties at a time resolution better than 10 ms. (B to G) Translational velocity for the same cell filtered from the displacement data corresponding to A at a time resolution of 750 frames/s (fps) (B), 250 fps (C), 30 fps (D), 2 fps (E), 1 fps (F), and 0.5 fps (G). The average velocity for each time interval is marked at the center of the interval. (H) High resolution of a 0.007-s segment of data in A. (I to L) Same data as in D to G with extended vertical axis.The instantaneous translational velocity at time t ¼ 5 s has values of 1.875, 1.875, 1.836, 0, 0, 0, and 0 mm/s for G, F, E, D, C, B, and A, respectively. The translational velocity at time t ¼ 5.62 s has values of 1.631, 1.623, 1.764, 17.633, 73.206, 84.301, and 99.253 mm/s for G, F, E, D, C, B, and A, respectively. The values at a given point in time approach the true value for this point as time resolution increases.

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tree until it becomes arrested or detaches (Kunkel et al., 2000). This type of approach is rarely used, because it is technically difficult and labor intensive, but it provides valuable insights into how the leukocyte rolling and arrest process actually works. The process of gradual deceleration of myeloid cells was discovered through this approach (Kunkel et al., 2000), prompting research into its molecular underpinnings (Zarbock et al., 2007). Transmigration is often reported as the number of transmigrated cells seen in the tissue, either by oblique reflected light illumination (Mempel et al., 2003) or by counting the cells after the tissue is harvested and stained ( Jung and Ley, 1999). The former approach provides instantaneous results, whereas the latter allows some differentiation of leukocyte types. To interpret transmigration data, adhesion must also be measured, because decreased adhesion can result in decreased transmigration without a specific defect in the transmigration machinery. Each step in the leukocyte recruitment cascade can be described by an efficiency parameter to account for this interdependence ( Jung et al., 1998).

3.1. Surgical considerations Except for the bat wing, all tissues suitable for transillumination intravital microscopy are accessible only after surgery. Surgery requires anesthesia and causes trauma, which increases leukocyte–endothelial interactions (Fiebig et al., 1991; Ley, 1994; Thorlacius et al., 1994). The surgical trauma can be minimized, but not eliminated. It is useful to work with clean (cleaned with 70% ethanol) instruments. Iris scissors are suitable to make the small incisions in the tissue to be studied. For acute experiments, it is not necessary to autoclave the instruments, but buildup of blood or other debris on surfaces and in pockets must be carefully avoided. These biofilms are rich breeding grounds for bacteria and other microorganisms producing bioactive mediators. For the same reason, all glass- and plastic-ware should be clean and endotoxin-free. For convenience, most labs use injectable anesthetics such as ketamine hydrochloride (125 mg/kg, Sanofi Winthrop Pharmaceuticals) and xylazine (12.5 mg/kg, Tranqui Vet, Phoenix Scientific). Atropine sulfate (0.025 mg/ kg, Fujisawa) can be added to reduce cardiac reflexes and tracheal secretion, which are a problem in Sprague-Dawley rats, but usually not in C57BL/6 mice. Venous access is mandatory if using injectable anesthetics, because supplemental doses will be necessary starting at about 1 h after initial anesthesia. Supplemental intraperitoneal injections are not acceptable, because it takes too long for the anesthetic to reach the brain, which creates a tendency to overdose. Intravenous injection has immediate effects that can be monitored by a momentary drop in blood pressure and heart rate, both of which recover quickly. Even with good technique, spontaneously breathing mice

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have reduced oxygenation with typical pO2 values around 70 mmHg instead of 100 when awake, and are hypercapnic with typical pCO2 values around 70 mmHg instead of 40. Mean arterial blood pressure is around 80 mmHg. In our lab, animals with mean arterial blood pressure below 60 mmHg are not used. The anesthetized mouse is kept warm by a water-circulation (Fisher Scientific, Pittsburgh, PA) heating mat. During the whole surgery, the tissue studied must be kept moist and warm. Keeping the tissue under physiologic conditions is of great importance for reproducible results. The superfusion solution must be iso-osmotic (about 280 to 300 mosm/kg), of tissue pH (7.3 to 7.4) and the correct temperature, which is 37  C for most tissues and 35  C for the cremaster. In muscle preparations, the pO2 should be low (about 30 mmHg or lower), because higher pO2 causes vasoconstriction (Lindbom et al., 1982; Duling, 1972). All four conditions can be met by using a bicarbonate-buffered superfusion solution (132 mM NaCl, 18 mM NaHCO3, 4.7 mM KCl, 2 mM CaCl2.2H2O, 1.2 mM MgCl2) percolated with 5% CO2 in N2 and warmed to 37  C by a water-jacketed beaker (Kunkel et al., 1996). The NaHCO3 should be added after starting the gas flow, because otherwise the solution will be too basic and calcium carbonate will precipitate. The tissue can be overlaid with or wrapped in plastic wrap, which prevents washing out tissue metabolites and local cytokines. To prepare the cremaster muscle, only the skin in the ventral area of the right scrotum is grabbed with surgical tweezers, lifted, and removed with scissors. It is important to touch only the skin with the tweezers, because otherwise the delicate underlying tissue is damaged. After the first cut, the incision is extended in a horizontal direction toward the left testicle (incision should be approximately 1 cm long). The connective tissue around the cremaster muscle is then carefully separated, avoiding touching the cremaster. After removing the connective tissue, the tip of the cremaster sack is seized with small atraumatic forceps and the remaining connective tissue is removed. The cremaster is then fixed on the raised area of a custom-built plexiglass stage (Fig. 11.3) with a pin manufactured from a 27-gauge cannula, so that the cremaster lies flat on the cover slip. Using iris scissors, an incision is made from the point of fixation up the center of the cremaster sack. Both arteries and veins should be saved in order to maintain a good blood supply and avoid bleeding. Usually, the cremaster is fed by three main arteries. For good results, it is imperative to leave the middle arterial tree untouched, which then falls in the center of the finished preparation. Four to five pins are pierced through the edges of the tissue into a silicone ring around the viewing window in order to hold the cremaster onto the stage. The last step in the preparation is to pin the epididymis and testis to the side. We do not sever the connection between cremaster and epididymis, because the artery–vein pair connecting the epididymis to the cremaster is often a major feed–drainage pair.

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Vertical divider wall Silicone ring

Cover slip

Figure 11.3 Photograph of the stage used for the cremaster muscle preparation. The mouse is placed with its hindpaws against a vertical divider wall that separates the viewing pedestal from the rest of the stage.The viewing window is sealed with a10-mm diameter round coverslip and surrounded by a well filled with cured silicone rubber, which serves to anchor the needles.This stage is milled from solid plexiglass to prevent leakage.

3.2. Fluorescence intravital microscopy Due to the structure and location of most tissues, leukocyte rolling cannot be investigated by transillumination intravital microscopy. In these cases, fluorescence microscopy has become a valuable technical tool to visualize leukocyte recruitment. This technique allows to study leukocyte recruitment in Peyer’s patches (Miura et al., 1987), peripheral lymph nodes (von Andrian, 1996), brain (Liu and Kubes, 2003), liver (Liu and Kubes, 2003), pancreas (Cavallari et al., 1967), lung (Tabuchi et al., 2008), and many more (Halin et al., 2005; Sumen et al., 2004). The fundamental problems with fluorescence intravital microscopy are light toxicity and photobleaching of all fluorescent dyes, both of which are caused by the generation of free radicals that produce cell and tissue injury. It is not uncommon to view a FITC-stained preparation with an Hg or Xenon lamp under epifluorescence (excitation in the blue range, emission in the yellow-green range) and see the fluorescent label completely disappear within a few seconds. Bleaching can be so quick that the exposure time for a still microphotograph must be adjusted while the photo is taken. In such circumstances, the results are uninterpretable, because severe tissue injury must be inferred from the observed photobleaching. Two techniques have been successful in limiting light toxicity. In stroboscopic epifluorescence microscopy (Ley and Gaehtgens, 1991), the tissue is

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illuminated for only a few microseconds and allowed to recover in the dark for about 30 ms before the next stroboscopic flash is applied. This technique almost completely eliminates photobleaching, yet achieves intensities and image contrast similar to continuous excitation. There are very few strobe units on the market that are fast enough for video rates. A commonly used unit is made by Rapp Engineering in Hamburg, Germany. Stroboscopic epifluorescence microscopy is fast (at least 30 frames/s) and allows for multiple exposures within the same frame (for example, 1 ms apart) such that velocities of fluorescent objects or cells in blood vessels can be determined from a single frame (Smith et al., 2003). This is much better than the streak method, which requires prolonged exposure to fluorescence illumination and is limited by photobleaching as well as relatively large errors in measuring streak length. Shuttered cameras can also produce short exposure times, but since the tissue is still exposed to continuous light, bleaching and phototoxicity remain problematic. Shuttered light sources are a better choice, although matching the light intensity and density (lumens per unit area) of a stroboscopic flash unit can be challenging. The disadvantage of stroboscopic epifluorescence microcopy is that its penetration depth is no better than conventional epifluorescence. The other technique is multiphoton microscopy, which limits phototoxicity to a very small volume in which the actual excitation takes place (Halin et al., 2005). Multiphoton microscopy provides excellent penetration depth and is ideal for following slow movements such as lymphocytes and dendritic cells migrating in lymph nodes. Theoretically, its resolution is a little lower than that of conventional bright field or confocal microscopy, because the wavelength of the exciting light is so long, but in practice the resolution is comparable to confocal microscopy, because the most important consideration is the wavelength of the image-forming light, which is essentially the same in both modalities. Multiphoton microscopy is unsurpassed at rejecting out-of-focus light and is a rapidly growing intravital microscopic modality. Tracking fluorescent cells, whether rolling or migrating, is much easier than in transillumination microscopy. Even centroid trackers that completely fail in the transillumination case provide decent results with fluorescent cells or even molecules (Cheezum et al., 2001). A usable tracker is available from Ed Marcus laboratories (http://www.marcuslabs.com/). The next consideration is the labeling technique used to detect the leukocytes. Rhodamine 6G (Galkina et al., 2007) and acridine orange (Ley and Gaehtgens, 1991) are widely used, but they cannot differentiate between leukocyte subsets such as neutrophils and lymphocytes except in rare cases (Tangelder et al., 1995). Several transgenic and knockin mice are available (Table 11.1) that express green fluorescent protein (GFP) in only one leukocyte subset under control of a subset-specific promoter (Faust et al., 2000; Jung et al., 2000; Manjunath et al., 1999; Singbartl et al., 2001; Unutmaz et al., 2000). Leukocyte subsets can also be isolated in vitro by various techniques, including density gradient (Percoll, Ficoll), magnetic

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Table 11.1 Examples of mice for intravital microscopy with GFP leukocyte subsets Gene name

Protein

Cells expressing GFP

References

Cxcr6

NKT cells, some T cells

Unutmaz et al., 2000

Cd4

CD4

Lzm

Lysozyme M

Csf1

MCSF receptor (CD115)

Monocytes, precursors, some NK cells T cells: bright CD8 cells, dim CD4 cells T cells: bright CD4 and CD8 cells Neutrophils, monocytes Monocytes

Jung et al., 2000

Cd2

Chemokine receptor CXCR6 Chemokine receptor CX3CR1 CD2

Cx3cr1

Singbartl et al., 2000 Manjunath et al., 1999 Faust et al., 2000 Sasmono et al., 2003

bead selection, and flow cytometric cell sorting. Following isolation, leukocyte subsets can be stained with a fluorescent dye and then injected during an in vivo experiment. While ex vivo labeling works well for lymphocytes, myeloid cells are unintentionally activated during isolation (Glasser and Fiederlein, 1990; Forsyth and Levinsky, 1990; Kuijpers et al., 1991). For example, in vitro isolated and stained neutrophils do not show normal rolling behavior when injected back into mice.

3.3. Lymphocyte rolling and adhesion in Peyer’s patches Peyer’s patches are secondary lymph organs in the mucosa and submucosa of the small intestine. The highest density of Peyer’s patches is found in the terminal ileum. Interaction of lymphocytes with high endothelial cells in Peyer’s patches can be visualized by epifluorescence microscopy. In order to investigate lymphocyte rolling in vivo, lymphocytes can be stained by injecting rhodamine (0.15 ml of a 2-mg/ml rhodamine 6G stock solution). The rhodamine injection stains all leukocytes, so the rolling velocity, rolling flux, and adhesion can be investigated, but discrimination between the different lymphocyte subsets is not possible. Different subsets of lymphocytes (e.g., T-lymphocytes) can be detected after isolation from lymphoid tissue and ex vivo staining. Peripheral and mesenteric lymph nodes are a good source of lymphocytes. The desired cell population can be separated by magnetic beads (positive or negative selection; these kits are available

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from Miltenyi Biotec, among others) or FACS sorter. The isolated cells are then stained with a cell tracker (10 mM 5-(and-6)-(((4-chloromethyl)benzoyl)amino)tetramethylrhodamine, CMTMR or 5 mM 5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester [CFSE], both from Molecular Probes). It is a good idea to switch trackers between experiments in order to exclude any influence of the cell tracker on cell function. After the isolation and labeling process, cells are injected intra-arterially intravenously and their rolling velocity, rolling flux, and adhesion are determined. After injecting the labeled cells, a wave of cells appears in the microcirculation of the Peyer’s patch, which subsides over time due to redistribution and adhesion of the cells. Therefore, it is important that the recording is always analyzed at the same time after injection. Investigation of the Peyer’s patch microcirculation starts by anesthetizing a mouse as described. The mouse is placed on a heating pad and a 1.5-cm aseptic incision is made along the linea alba in order to avoid bleeding. After opening the peritoneal cavity, the mouse is turned on its side, and the intestines are pushed out of the peritoneal cavity by gently pressing on the back. The intestines are superfused with temperaturecontrolled (37  C) superfusion buffer (see above). By using moist cotton swabs, the intestine is placed on the coverslip of the circularly raised area in a way that places the Peyer’s patch on top of the gut. In order to immobilize the small intestine and allow for visualizing of the Peyer’s patch microcirculation, a plastic coverslip cut to approximately 2 cm2 is overlaid with two parallel strips of silicone vacuum grease. This coverslip is then laid over the preparation, with the small intestine lying parallel between the two rows of grease and the Peyer’s patch is flattened against the coverslip.

3.4. Intravital microscopy of large vessels While intravital microscopy is immensely popular for use in microcirculation, large vessels can also be observed. Preparations have been described for the mouse aorta (Eriksson et al., 2001) and carotid artery (Huo et al., 2003). In both cases, the motion artifact is very large and limits the usefulness of these preparations. Isolated perfused mouse arteries are more widely used (Ramos et al., 1999; von Hundelshausen et al., 2001). The major limitations of these preparations include the need for surgical removal and cannulation, which can alter the physiological state of the endothelium, and the need to isolate and label blood cells to be investigated.

3.5. Mixed chimeric mice To compare two sets of leukocytes, such as knockout and wildtype in the same microvessel, mixed chimeric mice can be generated by bone marrow transplantation from two donors. Endothelial and hemodynamic parameters

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are identical for both types of leukocytes observed in the microvessels of the recipient mice, and even small differences in rolling velocity can be measured reliably. For this technique, it is necessary to combine transillumination and fluorescence microscopy (see above) in order to see greenfluorescence protein (GFP)–positive and negative cells at the same time. Recipient mice are lethally irradiated by two doses of 600 rad each approximately 4 h apart. Donor mice are killed by either a lethal injection of anesthesia or CO2 inhalation followed by cervical dislocation. Bone marrow cells are harvested by flushing both femurs and tibias with medium (RPMI 1640) under sterile conditions. For neutrophil investigations, the knockout cells are typically nonfluorescent and the wildtype cells express GFP under the lysM promoter (Faust et al., 2000), but many other combinations can be used. Suspended bone marrow cells are washed and erythrocytes are lysed in 0.15 M NH4Cl lysing solution. Approximately 2 to 5 million unfractionated bone marrow cells in 300 ml medium are injected intravenously through the tail vein of each recipient mouse. Recipient mice are housed in a barrier facility (individually ventilated cages with highefficiency particulate air filter) under pathogen-free conditions before and after bone marrow transplantation. After bone marrow transplantation, the mice need autoclaved and acidified water with antibiotics (5 mM sulfamethoxazole, 0.86 mM trimethoprim) and autoclaved food in order to prevent infection during the neutropenic period following reconstitution. Experiments can be performed 6 weeks after bone marrow transplantation. In order to determine the reconstitution rate, CD45.1 and CD45.2, both measured by flow cytometry, can be used. Alternatively or in addition, male donor marrow can be transplanted into female recipients and leukocyte DNA from the reconstituted mice can be investigated for an area specific to the Y chromosome by real-time PCR (Zarbock et al., 2006).

4. Spinning-Disk Confocal and Two-Photon Microscopy A major difficulty in deep tissue imaging is the scattering of light in thick tissues, resulting in image blurring. The use of conventional laserscanning confocal microscopy has greatly improved the resolution and contrast achievable in in vivo imaging. Confocal microscopy relies on the absorption of single high-energy (blue or green) photons, sufficient to excite fluorophores and produce a fluorescent signal. The excitation beam is focused at a particular plane within the tissue and the emitted fluorescent light is collected and focused onto a small pinhole, thus blocking fluorescent light coming from out-of-focus regions, and detected behind the pinhole

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(Denk and Svoboda, 1997; Norman, 2005). This allows the detection of structures not distinguishable by conventional epifluorescence microscopy. Laser-scanning confocal systems make possible the generation of threedimensional (3D) images by scanning the laser beam over every point of the focal plane, collecting a stack of sections from different planes and reconstructing a 3D image from the stack of sections. Although the acquisition of 3D images is generally time consuming and currently does not permit the tracking of rolling and free-flowing leukocytes in 3D in vivo vasculature, these systems continue to improve (Wolleschensky et al., 2006). The need for fast image acquisition during many intravital microscopy applications makes spinning-disk confocal systems a suitable choice (Nakano, 2002). The introduction of a rotating disk with small pinholes arranged in a spiral pattern allows the focusing of the excitation light at multiple points in the same focal plane. Simultaneous detection of fluorescence emitted from these points with a charge-coupled device (CCD) camera increases the rate at which images can be captured, as compared with laser-scanning confocal systems that use a photomultiplier tube to detect the fluorescent light. The spinning disk confocal microscopy was recently used to generate intravital microscopic recordings of endogenous T-cell trafficking (Norman et al., 2008). In conventional confocal microscopy, exposure to the excitation beam always extends throughout the entire depth of the specimen, while only the photons emitted from the focal plane are detected at each moment. Therefore, 3D confocal scans cause substantial fluorophore photobleaching and photodamage in the specimen. The problem is exacerbated by the scattering and absorption of the excitation light in thick specimens that require higher intensity of the excitation beam. In contrast to conventional confocal microscopy, two- or multi-photon microscopy relies on the almost simultaneous absorption of at least two photons of lower energy light in order to bring the fluorophores to the excited state. The requirement for multiphoton absorption confers the advantages of imaging at greater tissue depths, thinner optical sectioning, and decreased photobleaching and photodamage (Denk and Svoboda, 1997; Helmchen and Denk, 2005). Because of the multiphoton absorption requirement, the excitation only occurs in a small 3D region near the focal point of the excitation beam, where photon flux is greatest. Therefore, only fluorophores near the focal point are excited, and the photobleaching and photodamage in 3D optical scans are greatly reduced (Piston, 1999). Moreover, multiphoton microscopy also obviates the need for a pinhole at the detector, and the emission photons scattered on their way through the specimen still contribute to the usable signal, thus maximizing the efficiency of collecting the fluorescent light and reducing the photobleaching and photodamage even further. This is particularly advantageous when imaging deep tissue samples for long periods of time, as needed in time-lapse intravital microscopic techniques. The longer wavelength of the

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excitation light as compared to conventional confocal microscopy (near infrared vs. visible) reduces the absorption and scattering of the excitation beam in the specimen, but can cause thermal problems. (Rubart, 2004; Piston, 1999).

4.1. Autoperfused flow chamber The in vivo environment is very complex, and unknown factors may influence leukocyte rolling and adhesion. The vessel wall is covered by endothelial cells, which express different adhesion molecules at variable site densities leading to changes of leukocyte rolling behavior in the same vessel. Furthermore, hemodynamic parameters like wall shear stress change with the vessel diameter and flow rate. In order to simplify the environment and study leukocyte rolling under defined and reproducible conditions, several flow chamber systems have been developed. These systems allow examining the function of individual molecules in an environment that imitates hemodynamic conditions in blood vessels. Parallel plate-flow chambers were first used in 1912 (Burrows, 1912). In 1987, they were adapted to study selectin- and integrin-mediated leukocyte rolling and adhesion (Lawrence et al., 1987), and since this time, the parallel-flow chamber was modified several times (Cao et al., 1997; Ruel et al., 1995; Usami et al., 1993; Yuan et al., 2001). Because conventional parallel plate flow chambers consume a lot of liquid, they are not suitable for perfusion with whole mouse blood and normally require isolation and resuspension of the cells to be studied. The isolation is problematic for neutrophils and other myeloid cells, because it is known to activate the cells and cause increased expression of Mac-1 and decreased surface expression of L-selectin (Glasser and Fiederlein, 1990; Forsyth and Levinsky, 1990; Kuijpers et al., 1991). Dramatic changes in gene expression were also observed when isolating blood monocytes (Cho et al., 2007). Initial designs of autoperfused flow chambers, which were made of a glass capillary connected to the mouse by plastic tubing, required returning the blood into the donor mouse (Smith et al., 2004; Hafezi-Moghadam et al., 2004), which can be problematic because of accumulation of activating factors induced by blood passage through the capillary and tubing. A more recent modification has a substantially reduced cross-sectional area of the glass capillary, 20
200 mm, thus obviating the need to return blood (Chesnutt et al., 2006). As an added benefit, visualization of unlabeled leukocytes is possible. Furthermore, the use of the small cross-section capillaries makes it easy to control the wall shear stress and reduces the blood volume loss to a minimum (typically less than 1 ml/min). The flow chamber is assembled by placing a rectangular capillary (20 mm
200 mm, Fiber Optic Center, New Bedford, MA) with a length of 30 mm between two plastic microscope coverslips. The coverslips are attached to a glass microscope slide by using clear nail polish. Following the assembly, the

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capillary is coated with one or more adhesion molecules (recombinant adhesion molecules from R&D Systems, Minneapolis, MN: P-selectin/Fc, 20 mg/ ml; E-selectin/Fc, 30 mg/ml; ICAM-1/Fc, 15 mg/ml) dissolved in PBS by a 2-h incubation of the solution in the capillary. During the incubation time, the capillary is inserted into a 5-cm–long heparinized polyethylene (PE) 50 tubing (ID 0.58 mm, OD 0.965 mm, Becton Dickinson). The connection is sealed with a two-part epoxy gel resin. After the coating, the surface of flow chambers is blocked by a 1-h incubation of 10% casein solution (Pierce Chemicals), the free end of the capillary is placed in a saline-filled PE 50 tubing with a length of 150 cm, and the connection is sealed with a two-part epoxy gel resin. Mice are anesthetized and temperature controlled. The carotid artery is cannulated with a heparinized PE 10 tubing (ID 0.28 mm, OD 0.61 mm) connected to PE 50 tubing attached to the capillary by a metallic connector (Small Parts Inc.). To observe leukocyte rolling, the flow chamber is placed on an intravital microscope with a saline immersion objective. The wall shear stress in the autoperfused flow chamber can be controlled by adjusting the downstream 150-cm–long, saline-filled PE 50 tubing fitted to an open syringe. Zero pressure is determined by raising the outflow syringe until flow stops. The average wall shear stress (tw) in the autoperfused flow chamber can be calculated as

tw ¼

DPwh ; 2Lðw þ hÞ

where DP is the pressure head across the flow chamber, w is width, h is height, and L is length of the flow chamber. This shear stress estimate is valid at the upper and lower wall in the middle two-thirds of the chamber width, whereas the shear stress near the side walls is much lower. The typical height, width, and length of the flow chamber are 20 mm, 200 mm, and 30 mm, respectively (Chesnutt et al., 2006; Smith et al., 2004). In order to investigate the rolling velocity of neutrophils, cells can be observed and recorded while rolling on E-selectin, P-selectin, their combinations with ICAM-1, or other substrates. For chemokine-induced leukocyte arrest studies, we co-immobilize a chemokine (CXCL1, Peprotech) with adhesion molecules (typically a selectin and ICAM-1) (Zarbock et al., 2007). The flow chamber is perfused with blood for 6 min, and the rolling and arrest are subsequently recorded for 1 min in one representative field of view. The performance of the autoperfused flow chamber is limited by platelet accumulation, which usually starts at about 10 min. Presumably, platelet accumulation is mediated by von Willebrand factor, a very sticky protein in plasma. A disadvantage of the autoperfused flow chamber is that the site densities of adhesion molecules cannot be measured by radioimmuno assay, because the assay’s sensitivity is insufficient to detect the

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number of molecules on the small surface of the flow chamber. As an upper bound, it can be estimated that all molecules contained in the capillary were immobilized on the surface (Chesnutt et al., 2006).

4.2. Microfluidic flow chambers The performance of flow chambers is further enhanced by using microfluidic technology that allows fabricating complex microchannel networks (Toner and Irimia, 2005). Microfluidic devices have been used to capture different sub-populations of lymphocytes to substrates with various coatings from continuous flow (Cheng et al., 2007; Murthy et al., 2004), to test the strength of adhesion of neutrophils to a substrate (Gutierrez and Groisman, 2007), as well as to study the dynamics of rolling and adhesion of platelets to different substrates (Gutierrez et al., 2008) and of neutrophils to endothelial monolayers (Schaff et al., 2007). Microfluidic devices have become particularly attractive for academic research with the introduction of soft lithography and rapid prototyping (Whitesides et al., 2001). With these techniques, customized microchannel devices can be fabricated at low cost, quickly upgraded and improved, and produced in amounts sufficient for long series of repeated experiments. In a microfluidic device machined using soft lithography, a microchannel network is engraved on the surface of a chip made of polydimethylsiloxane (PDMS), an optically clear, chemically inert, porous silicone rubber. The microchannels are sealed by a regular microscope coverglass, which reversibly adheres to the PDMS chip. The chips are cast from master molds, which are lithographically fabricated using simple equipment, inexpensive materials, and fast protocols. A microfluidic device is typically placed with its coverglass side onto the stage of an inverted microscope (Figure 11.4A). The surface of the coverglass at the bottom of the microchannels can be coated with adhesion molecules, thus becoming a substrate for rolling and adhesion of leukocytes from blood flowing in the microchannels. Because of their construction and low autofluorescence of PDMS (lower than glass), the microfluidic devices are fully compatible with brightfield and fluorescence microscopy, including imaging with high-resolution immersion objective lenses. Moreover, segments of PE tubing inserted into inlets and outlets of a PDMS chip are held in place by the flexible silicone rubber and the connections are reliably sealed, making the microfluidic devices easier to work with than the glass capillary perfusion chambers discussed above. An important new capacity provided by the microfluidic technology is the possibility of concurrently testing the interaction of leukocytes from a small blood sample with differently coated substrates at a variety of flow conditions. So, the microfluidic device in Fig. 11.4B has an array of eight parallel flow chambers, each with a 24
200 mm cross-section, between the common inlet and outlet (Gutierrez et al., 2008). The hydrodynamic

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A

PE tubing Inlet PDMS chip

Objective

Outlet Cover glass Micro-channel with blood

B

Resistance channels Flow chamber 8 Inlet

Feeder channel

Outlet Bypass Flow chamber 1 24 mm deep

2 mm

250 mm deep

C

Collector channel

Out2 Vent

Out3

Inlet Out1

2 mm

Aux

Flow chambers

Figure 11.4 Microfluidic flow chambers. (A) Cross-section through a microfluidic device made of an engraved PDMS chip sealed with a regular microscope coverglass. (B) Layout of microchannels in the microfluidic device with an array of eight flow chambers used to study dynamic adhesion of platelets to substrate at various shear stresses. (Adapted from Gutierrez, E., Petrich, B. G., Sanford, J. S., Ginsberg, M. H., Groisman, A., and Kasirer-Friede, A. (2008). Exploring the role of integrin aIIb3 in sheardependent platelet adhesion in microfluidic devices. J Thomb Haemost (submitted).) 250-mm and 24-mm^deep channels are shown in light and dark gray, respectively.When a differential pressure of 2.5 kPa is applied between the inlet and outlet, wall shear stress varies from 40 dyn/cm2 in chamber 1to 4000 dyn/cm2 in chamber 8. (C) Micrograph of a microfluidic device, where the interaction of leukocytes with three differently coated substrates is tested concurrently at the same shear stress. Blood fed to the inlet is directed through three identical flow chambers to three separate outlets (out1 to out3). Prior to the experiment with blood, the flow chambers are coated in situ by perfusion with various protein solutions that are fed to ports out1 to out3 and drawn off an auxiliary outlet (aux).

resistances of microchannels connected to the flow chambers are such that the surface shear stress, tw, varies by a factor of 2 between the chambers, covering a 100-fold range across the array. Thus, the interaction of cells with the substrate can be simultaneously tested at conditions ranging from low venous to high arterial flow, with blood consumption below 4 ml/min

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and less than 100 ml per experiment. In the microfluidic device in Fig. 11.4C, which we built to simultaneously test the interaction of neutrophils with three differently coated substrates, whole mouse blood injected into the inlet is directed to three identical flow chambers that are 20
450 mm in cross-section. Three outlets (out1 to out3) and an auxiliary port (aux) of the device enable independent in situ coating of the flow chambers with various proteins by perfusion of the chambers with solutions of the proteins (e.g., P-selectin, P-selectinþICAM-1, and casein). Another port of the device (vent) is used to measure the pressure at the inlet (by a compensation method [Groisman et al., 2003]), when blood is coming to the inlet from an anesthetized animal.

5. Conclusion Intravital microscopy has undergone a tremendous renaissance, with much of the current effort focused on leukocyte and lymphocyte interactions with the vessel wall and tissue structures. The advent of new techniques like multiphoton and stroboscopic epifluorescence intravital microscopy has enabled studies that were not accessible to microscopic techniques even a few years ago. Autoperfused flow chambers and microfluidic devices add a new dimension to the molecular study of leukocyte adhesive interactions in their native whole blood.

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Placental Remodeling of the Uterine Vasculature Nathan M. Hunkapiller and Susan J. Fisher Contents 1. Introduction 2. Isolation and Culture of Human Cytotrophoblasts 2.1. Cytotrophoblast isolation procedure 3. Isolation and Culture of First-Trimester, Human Placental Villous Explants 4. Identification of Cytotrophoblast-Modified Blood Vessels in Tissue Sections 5. Cytotrophoblast Migration and Induction of Endothelial Cell Apoptosis During Co-Culture 5.1. Co-culture of Jar choriocarcinoma cells with endothelial monolayers 5.2. Co-culture of first-trimester CTBs with differentiated endometrial endothelial cells in a three-dimensional model 6. In Vitro Models of CTB Endovascular Invasion Using Explanted Spiral Arterioles 6.1. CTB invasion of term myometrial spiral artery segments in vitro 6.2. Chorionic villous CTBs invade first-trimester decidual explants in vitro 7. In Vivo Models of Human Cytotrophoblast Vascular Remodeling 7.1. Placental xenograft of the murine mammary fat pad and kidney capsule 7.2. Choriocarcinoma xenograft of the murine skin References

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Abstract In eutherian mammals, the first functional organ is the placenta, a transient structure that is rapidly assembled in the extraembryonic compartment. By necessity the placenta develops in advance of the embryo, which it supports

Department of Cell and Tissue Biology, University of California-San Francisco, San Francisco, California Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03012-7

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in utero by performing many of the same functions that the lungs, gastrointestinal tract, and urinary system carry out after birth. Specialized epithelial cells that arise from the placenta, termed cytotrophoblasts (CTBs), are responsible for redirecting maternal blood to the developing conceptus, which occurs as a result of the cells’ aggressive invasion through the maternal endometrial stroma (interstitial invasion) and resident blood vessels (endovascular invasion). The latter process involves displacement of maternal endothelium and induction of apoptosis in the surrounding smooth muscle. Together, these events result in a reduction of blood vessel elasticity and increased blood flow. In the past, investigations of human CTB endovascular invasion have been limited to immunohistochemical examination of tissue sections. In this chapter, we will discuss the use of in vitro and in vivo techniques that have been recently adapted for the study of the complex events that occur during CTB endovascular invasion. As an introduction, we provide background on placental anatomy and the molecular basis of CTB behaviors. To follow, we present techniques used in the isolation and culture of primary CTBs and chorionic villous explants. Approaches for identifying trophoblast-modified blood vessels in placental tissue sections are also described. Next, we review methods used by other groups to study CTB/endothelial interactions in culture focusing on techniques that employ isolated cells and chorionic explants. Finally, we conclude with methods devised by our group and others to explore the complex heterotypic cell–cell interactions that occur as CTBs invade blood vessels in vivo in the nude mouse.

1. Introduction The placenta is commonly depicted as a simple, disc-shaped structure connecting the embryo/fetus to the uterus (Fig. 12.1A). Within the developed placenta is a cavity (intervillous space) that possesses a vast network of chorionic villi which float in circulating maternal blood. Here, nutrients, gases, and wastes are exchanged across the surface of these fetal villi between the maternal blood and the fetal capillary beds underlying the villi. These structures are lined by two layers of trophoblast. The outer layer is composed of a multinucleated syncytium (syncytiotrophoblast) while the inner layer is a polarized CTB epithelial layer. Uterine attachment requires CTBs to differentiate and acquire the ability to aggressively invade maternal tissues (Fisher et al., 1989; Librach et al., 1991). This process also initiates the remarkable cell–cell interactions that occur as CTBs remodel the uterine vasculature into chimeric vessels, composed of fetal and maternal cells, that supply blood to the placenta. Key morphological aspects of the invasion process are diagrammed in Fig. 12.1B. CTB progenitors form a polarized epithelium that is attached to the basement membrane surrounding the mesenchymal cores of chorionic villi. During differentiation along the

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A

B Syncytiotrophoblasts

Fetal side Maternal side

EC

Mesenchymal core of anchoring villus

EC

Progenitor CTBs Proximal cell column CTBs Distal cell column CTBs

Chorionic plate

Invasive CTBs

Umbilical cord

Fetal membranes CTB differentiation

B Basal plate Floating villi Intervillous space (gray area)

Smooth muscle wall

Maternal spiral arteriole

Endovascular CTBs

Maternal ECs

Figure 12.1 Anatomy of the maternal^fetal interface: Anchoring chorionic villi (B) attach the placenta to the uterus and give rise to invasive CTBs. (A) Placental chorionic villi stem from the chorionic plate and lie within the intervillous space. The point of attachment between anchoring villi and the underlying tissue is referred to as the basal plate (box B). (B) Enlargement of the area in box B. Undifferentiated CTB progenitors in the anchoring villi give rise to invasive CTBs that invade the uterine interstitium (interstitial invasion) and the maternal endothelium (endovascular invasion).

invasive pathway, CTBs leave this basement membrane to form columns of unpolarized cells that attach to and then penetrate the uterine wall. The ends of the columns terminate within the superficial endometrium, where they give rise to invasive (extravillous) CTBs. During interstitial invasion, a subset of these cells, either individually or in small clusters, commingles with resident decidual, myometrial, and immune cells. During endovascular invasion, masses of CTBs breach and plug the vessels (Ramsey et al., 1976). Subsequently, these fetal cells replace the resident maternal endothelium and portions of the smooth muscle wall. Normally, this process encompasses the portions of uterine arterioles that span the decidua and the inner third of the myometrium. In contrast, only the termini of uterine veins are breached (Zhou et al., 1997). Together these two components of CTB invasion anchor the placenta to the uterus and permit a steady increase in the supply of maternal blood that is delivered to the growing embryo/fetus. Endovascular invasion, which is a major determinant of pregnancy outcome, incorporates the placenta into the maternal circulation. Some of the key molecular aspects of human CTB differentiation and invasion are known. The cells’ expression of several classes of functionally relevant molecules is precisely modulated as they invade either in situ (the uterine wall) or in vitro (extracellular matrix [ECM]). For example, work from our group and other investigators has produced a detailed picture of

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how invading CTBs modulate the expression of molecules that facilitate both interstitial and endovascular invasion. With regard to adhesion molecules, the onset of CTB differentiation/invasion is characterized by reduced staining for receptors characteristic of polarized CTB epithelial stem cells— such as integrin a6b4 and E-cadherin—and the onset of expression of adhesion receptors characteristic of endothelium—such as VE-cadherin, Ig family members VCAM-1 and PECAM-1, and integrins aVb3 and a1b1 (Blankenship and Enders, 1997; Damsky et al., 1992; Vicovac et al., 1995; Zhou et al., 1997). A portion of this switching program is regulated by the wide array of VEGF family members that CTBs produce (Zhou et al., 2002, 2003). Recently we discovered that CTBs also express L-selectin, a leukocyte cell surface receptor that promotes adhesion under shear stress (Genbacev et al., 2003). Similarly, cadherin-11 is upregulated on extravillous CTBs and on decidualizing endometrial stroma (MacCalman et al., 1996, 1998). Whether this interaction promotes or restrains interstitial CTB invasion is not yet clear. All CTBs in this pathway, regardless of location, express Mel-CAM, a melanoma-associated Ig family receptor also expressed on endothelium (Shih and Kurman, 1996). Finally, CTBs within the maternal blood vessels turn on neural cell adhesion molecule (NCAM, CD56) (Blankenship and King, 1996; Y. Zhou et al., unpublished observations), an adhesion receptor that is also expressed by maternal natural killer cells that home to the pregnant uterus (Starkey et al., 1988). Changes in the CTB adhesion molecule repertoire take place in the context of the cells’ equally dramatic modulation of their proteinase and proteinase inhibitor expression. Some aspects of this phenotypic transformation are undoubtedly linked to CTB acquisition of an invasive phenotype. For example, we found that expression and activation of matrix metalloproteinase (MMP)-9 is required for invasion in vitro (Librach et al., 1991). We speculate that CTBs upregulate the expression of other proteinases/inhibitors to present a nonthrombogenic surface to maternal blood. The urokinase plasminogen activators uPA (Queenan et al., 1987) and PAI-1 (Feinberg et al., 1989), as well as the thrombin receptor (EvenRam et al., 1998), might function in this manner. Invading CTBs upregulate the expression of immune molecules, which likely enables them to escape maternal rejection. Mechanisms used for this purpose include their expression of HLA-G, a unique major histocompatibility class Ib antigen with limited polymorphism (Ellis et al., 1990; Kovats et al., 1990), and interleukin-10, a potent immunosuppressive cytokine (Roth et al., 1996). Finally, CTBs modulate expression of Eph/ephrins that direct their migration through the uterus. Very recently, we found that, as CTBs begin to differentiate toward an invasive phenotype, they downregulate expression of EphB4 and upregulate expression of ephrinB2 (Red-Horse et al., 2005). Our work suggests that due to repulsive interactions induced by ephrinB2/ EphB4 ligation, ephrinB2-expressing CTBs are directed away from their

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EphB4-expressing progenitors and toward ephrinB2-expressing arteries, where permissive interactions take place (Red-Horse et al., 2005). In the sections that follow, we discuss methods applicable to the study of human endovascular invasion. Initially we provide basic techniques required for the isolation and culture of primary CTBs (Section 2) and explants (Section 3), as well as means for identifying CTB-modified blood vessels in tissue sections (Section 4). Next, we review methods used by other groups to study CTB/endothelial interactions in both cell culture (Section 5) and explant models (Section 6). We conclude with in vivo methods devised by our group and others to explore the complex heterotypic cell–cell interactions that occur as human CTBs invade blood vessels in the immunodeficient mouse (Section 7).

2. Isolation and Culture of Human Cytotrophoblasts To study cell behavior in culture, we isolate CTBs from whole placental tissue collected after elective pregnancy termination or at term. CTBs reside within the villous tissue of the placenta, which is also composed of syncytiotrophoblasts, red blood cells, fibroblasts, endothelial cells, leukocytes, and other mesenchymal progenitors. The major steps in the cell isolation procedure include removal of the outer layer of syncytium, release of the underlying CTBs by sequential enzymatic digestion, and purification of the cells by Percoll density-gradient centrifugation. Any remaining leukocytes are removed by virtue of their interactions with CD45-coated beads, which are then separated with a magnetic device. Using these methods for CTB isolation, we typically achieve greater than 95% purity. The viability and purity of the cells are subject to a number of factors including tissue quality, gestational age, and tissue handling during the procedure. Our methods have never been published in detail, and as such, we will describe the procedure that we have developed over the past 20 years.

2.1. Cytotrophoblast isolation procedure 1. Working with human tissue requires approval by an institutional human subjects review board. 2. Obtain the placental tissue as soon as possible after elective termination/ delivery and place immediately in cytowash medium (see Table 12.1). If the tissue is exposed to water, the CTBs initiate apoptosis. 3. Keep the tissue on ice prior to dissection. 4. Perform all work in a biosafety hood using techniques appropriate for working with potentially biohazardous tissues.

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Table 12.1 Preparation of medium, enzyme solutions, and Percoll density gradient Serum-free medium

500 ml

Cytowash

500 ml

DME/H-21 medium (Gibco, cat. # 11965092) Nutridoma (Roche, 11011375001)

469.5 ml

DME/H-21 medium (Gibco, cat. # 11965-092) Fetal bovine serum (Hyclone, SH30071.03) Glutamine Plus (Atlanta Biologicals, B92010) Penicillin/streptomycin (Invitrogen, 15140-122) Gentamicin (Invitrogen, 15750-060)

477 ml

10 ml

Sodium pyruvate (Sigma, 5 ml S8636) HEPES buffer 5 ml (Invitrogen, 15630080) Glutamine Plus (Atlanta 5 ml Biologicals, B92010) Penicillin/streptomycin (Invitrogen, 15140122) Gentamicin (Invitrogen, 15750-060)

12.5 ml

5 ml 5 ml

0.5 ml

5 ml

0.5 ml

Collagenase solution

100 ml

Trypsin solution

100 ml

1
PBS Mg2þ Ca2þ free (Gibco, 14190-144) Collagenase (Sigma, C-2674) DNase (Sigma, DN25) Hyaluronidase (Sigma, H-3506) BSA (Sigma, A7906)

100 ml

100 ml

0.062 g

1
PBS Mg2þ Ca2þ free (Gibco, 14190-144) Trypsin (Sigma, T-8003)

0.0069 g

0.040 g 0.069 g

DNase (Sigma, DN25) EDTA (Sigma, ED2SC)

0.0400 g 0.0200 g

0.100 g

Percoll gradient reagents

Percoll (Amersham, 17-0891-01) 10
Hanks’ BSS without phenol red (Invitrogen, 14185) 1
Hanks’ BSS without phenol red (Invitrogen, 14175) 1
Hanks’ BSS with phenol red (Invitrogen, 14170)

a

Percoll stock solutions

90% Percolla

Hanks’ 1


70% 60% 50% 40% 30% 20%

35.0 ml 30.0 ml 25.0 ml 20.0 ml 15.0 ml 10.0 ml

10.0 ml (w/o phenol red) 15.0 ml (with phenol red) 20.0 ml (w/o phenol red) 25.0 ml (with phenol red) 30.0 ml (w/o phenol red) 35.0 ml (with phenol red)

90% Percoll (270 ml Percoll þ 30 ml 10
Hanks’ BSS without phenol red).

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5. Place the tissue in a Petri dish and add fresh cytowash. Frequent exchange of the wash medium helps remove contaminating pathogens and maternal blood. 6. Remove fibrinoid, blood clots, amnion, and basement membrane from the placental villi using vannas scissors and forceps. Dissect villi into approximately 2- to 4-mm pieces. 7. Filter the tissue through a mesh strainer (1-mm gaps) to remove small pieces of villi that would otherwise undergo digestion too quickly, which results in DNA breakdown, a process that entraps the cells in a viscous gel. 8. Wash the remaining tissue with 1 l or more of cytowash to remove any remaining maternal blood. 9. Pour the dissected tissue into a 50-ml conical tube and fill with cytowash. Centrifuge the tube at 1300 rpm for 5 min at 4  C. This centrifugation speed corresponds to 365 rcf in a Sorvall Legend RT centrifuge. 10. Aspirate the supernatant and weigh the tissue pellet within the tube. 11. Prewarm the first enzyme dissociation solution by adding a volume of collagenase (Table 12.1) that is equal to six times the weight of the tissue to a sterile flask and place in a 37  C water bath shaker for approximately 10 min or until the desired temperature is reached. 12. Add the tissue to the warmed collagenase solution and shake in the water bath at 175 rpm for approximately 7 min. The amount of time necessary for digestion will depend heavily on the gestational age and health of the tissue. This step removes the syncytiotrophoblast layer (Fig. 12.2A). 13. Transport the flask from the water bath to the biosafety hood and place it on ice at an angle to allow the tissue to settle at the bottom. 14. Aspirate and discard the supernatant. 15. During Step 11, prewarm the trypsin solution (Table 12.1) to 37  C. 16. Add a volume of trypsin solution equal to six times the weight of the tissue, return flask to the water bath, and shake at 175 rpm for approximately 10 min at 37  C. The exact length of this incubation will again depend greatly on tissue health and gestational age. This step releases the CTBs from the villous tissue (Fig. 12.2B). 17. Transport the flask from the water bath to the biosafety hood and place on ice at an angle that allows the tissue to settle at the bottom. Immediately after the remaining tissue settles, collect the supernatant and add an equal volume of cytowash. 18. Place two layers of sterile gauze over the mouth of a 50-ml conical tube that contains 5 ml of fetal bovine serum (Hyclone, SH30071.03). To halt enzymatic digestion, transfer 45 ml through the gauze. Note: This step will require multiple 50-ml tubes, with the exact number depending on the volume of dissociation buffer used. 19. Centrifuge the tubes at 1300 rpm for 8 min at 4  C.

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Collagenase I

A

Collagenase II

C Percoll

Trypsin

B

D

Figure 12.2 Monitoring enzymatic digestions and cell purification during CTB isolation: (A) The first collagenase digestion releases the syncytiotrophoblasts from the surface of placental villi, which appear as multinucleated flattened cells at 20
magnification.The presence of any CTBs in the supernatant from the first collagenase digestion indicates that this step is complete and must be stopped immediately (not shown). Large clumps of CTBs are released by the trypsin digestion (B), which are broken down into single cells during the second collagenase treatment (C). Also present at this stage are blood cells, fibroblasts, leukocytes, and other cell types. (D) CTBs are preferentially isolated by the Percoll density-gradient, centrifugation step.

20. Meanwhile, prepare the Percoll gradient (Table 12.1) that will be used later. Slowly layer 6 ml of each concentration in a 50-ml conical tube in the following order: 70, 60, 50, 40, 30, and 20%. Store at 4  C. Make one tube for each 7 g of starting tissue. 21. Aspirate and discard the supernatant. Resuspend and combine the cell pellets in a single 50-ml conical tube with 50 ml of cytowash. Centrifuge again at 1300 rpm for 8 min at 4  C. 22. Aspirate and discard the supernatant. Then, resuspend the cell pellet in prewarmed collagenase solution using 1 ml/g of starting tissue. 23. Place the cell suspension in a sterile flask and transfer to the water bath. Shake the flask at 150 rpm for 3 min at 37  C. This second collagenase digestion dissociates clumps of CTBs into single cells (Fig. 12.2C). 24. Add cytowash to the collagenase solution up to 50 ml. Pass the cell suspension through a 100-mm cell strainer into a 50-ml conical tube. Centrifuge at 1300 rpm for 8 min at 4  C. 25. Aspirate and discard the supernatant. For every 7 g of starting tissue, resuspend the cell pellet in 4 ml of serum-free medium (Table 12.1). Very slowly add 4 ml of the cell suspension to the top of the Percoll gradient. Note: Multiple Percoll tubes are usually required.

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26. Centrifuge the tubes at 2700 rpm for 25 min at 4  C. This centrifugation speed corresponds to 1573 rcf in a Sorvall Legend RT centrifuge. This step separates cells according to their individual densities. 27. Aspirate the top two bands of the Percoll gradient. Collect the CTBcontaining layers between 20 and 28 ml. Combine the 8 ml harvested from each tube into a fresh 50-ml conical tube and fill with cytowash. Centrifuge at 1300 rpm for 8 min. The resulting semipurified preparation will contain CTBs along with other cells of similar size (Fig. 12.2D). 28. Aspirate and discard the supernatant, resuspend the pellet in 50 ml of cytowash, and centrifuge at 1300 rpm for 8 min at 4  C. 29. Aspirate and discard the supernatant, resuspend the pellet in 10 ml of serum-free medium. 30. Count cells using a hemocytometer. 31. Add cytowash to a final volume of 50 ml and centrifuge at 1300 rpm for 8 min at 4  C. 32. Aspirate the supernatant and resuspend the pellet in serum-free medium at a concentration of 106 cells/ml. Ideally, the cells are approximately 80 to 90% pure at this stage. Greater than 95% purity can be achieved by negative selection using CD-45–coated Miltenyi beads (Miltenyi Biotec), according to the manufacturer’s directions. 33. Prior to plating cells, coat tissue culture wells with a 1:1 mixture of Matrigel (BD Biosciences, cat. # 354234) and serum-free medium. For each well of a six-well plate, use 50 ml of the Matrigel:serum-free medium mixture. Allow the Matrigel to polymerize for 15 min at 37  C. Note: We use Matrigel lots prescreened for their ability to support CTB adhesion. 34. Plate 3 to 4 million cells per well. 2.2.1. Time considerations for enzymatic digestions First collagenase digestion: The gestational age and health of the tissue greatly affects the digestion time period. For example, first trimester tissue between 6 and 10 weeks of gestation usually requires about 6 to 8 min of digestion, tissue aged 13 weeks needs about 12 min, and villi from a 20-week placenta need 18 min or more. A good rule of thumb is to digest 1 min for every week in age. With regard to the health of the tissue, fibrinoid deposits and/ or clotted blood will add to the digestion time. Collagenase digestion can be monitored in real time by placing a drop of the reaction under a phasecontrast microscope and checking whether the syncytium has lifted from the surface of the villi (Fig. 12.2A). Clumps of syncytium should be visible in the enzymatic supernatant. The first appearance of large single cells (CTBs) indicates that the collagenase reaction has reached completion and

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should be halted. Note that the exact times used in this and all the other digestion steps are recalculated for each new batch of enzyme, which we purchase in lots. Trypsin digestion: The time required also increases with gestational age. Most first-trimester tissues will be adequately digested in about 10 min. Second trimester villi require about 14 min of incubation. As in the collagenase digestion, the enzymatic reaction progress can be verified under the microscope. Clumps should be visible along with single large CTBs (Fig. 12.2B). The larger clumps of CTBs will be dissociated in the second collagenase digestion (Fig. 12.2C). The presence of smaller fibroblast cells from the villous core indicates that the tissue has been over digested and the reaction must be stopped immediately. Second collagenase digestion: This step is less subject to factors that change the required time. Digestion for 3 min is almost always sufficient and should never exceed 5 min.

3. Isolation and Culture of First-Trimester, Human Placental Villous Explants Different culture conditions can be used to study a wide variety of CTB behaviors, including differentiation and invasion (recently reviewed in Miller et al., 2005). We have used the techniques described herein to generate explants for analysis in vitro (Prakobphol et al., 2006) and in vivo (Red-Horse et al., 2006) using the nude mouse as a host. In general, explant culture requires careful selection of anchoring villous remnants with attached cell columns. Floating villi are not adequate in studies of invasion because the component CTBs are programmed to differentiate into syncytium rather than invasive cells. Under a phase microscope, cell columns extend well beyond the translucent villous core, appearing more opaque with an irregularly shaped border (see Fig. 12.3A). Cell columns are most A

CC B

FV

Figure 12.3 Chorionic villous explants: (A) Remnants of anchoring villi containing CTB cell columns can be distinguished from floating villi by their irregular outlines and opaque caps that extend from their termini (inset). (B) Explant cultured on Matrigel showing CTB outgrowths (inset). CCs, cell columns; FV, floating villi.

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frequently observed in association with placentas up to 7 weeks of gestation. Placentas are collected and transported as described in Section 2 and dissected under stereomicroscopic visualization into explants between 2 and 5 mm in diameter. Depending on the application, the villous explants may be cultured in any number of conditions. For illustrative purposes, the following is a description of one method for culturing anchoring villous explants that we have used in many different applications. Briefly, villi are transferred to Matrigel-coated, 12-mm inserts that are placed in 24-well plates and completely submerged in several drops of medium (Ham’s F-12/Dulbecco’s modified Eagle’s medium [1:1, vol/ vol]) supplemented with an antibiotic/antimycotic mixture (100 U/ml penicillin, 100 mg/ml streptomycin, 50 mg/ml gentamicin) and 10% fetal calf serum. Eight to 24 h later, when the cell columns have adhered to the matrix, additional medium is added. The cultures are incubated at 37  C under standard conditions (5% CO2/95% air) for 72 to 96 h. Medium is changed daily. The morphology of the villi and their CTB outgrowths should be monitored at regular intervals using an inverted-phase microscope (Fig. 12.3B). Explants with similar growth patterns should be paired as experimental and control samples.

4. Identification of Cytotrophoblast-Modified Blood Vessels in Tissue Sections Microscopic analysis of histological sections of the pregnant uterus that contain spiral arterioles is often the starting point for gathering information about molecules that are relevant to CTB endovascular invasion. While this approach offers only a snapshot of the invasion process, it is a necessary prerequisite in forming hypotheses about the molecules that are involved. To play a role, they must be expressed in a relevant location at the right time. By way of background, CTB invasion of maternal blood vessels occurs in several stages. The fetal CTBs breach the termini of both arteries and veins. Within veins, invasion rarely progresses beyond the point of entry. In contrast, invasion within the spiral arterioles ultimately involves almost the entire intra-uterine course of these vessels, and normally goes as deep as the first third of the myometrium (Brosens et al., 1972). During much of the first trimester, these fetal cells plug uterine arterioles, an arrangement that is thought to maintain a hypoxic environment that favors CTB proliferation (Burton et al., 1999). Starting at about 10 weeks of gestation, the cells begin to migrate in a retrograde fashion up the vessel lumina, which rapidly recannulate (Enders and Blankenship, 1997). Once the vessels are fully modified, which includes a 10- to

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50-fold increase in diameter, they can deliver liters of blood per minute to the intervillous space. In these types of studies, several criteria should be considered, including the gestational age of the tissue and the site of placental insertion. Regarding gestational age, it should be noted that elective pregnancy termination in the first trimester is most commonly achieved by vacuum aspiration, a process that rarely, if ever, yields intact CTB-modified maternal blood vessels. In our experience, such vessels are best obtained following hysterectomies performed during the early gestation period. However, tissue acquired from terminations that are performed during the second trimester period by curettage often contains intact maternal blood vessels, which are grossly visible on the surface of the basal plate (Fig. 12.1A). These vessels are also found in association with placental tissue acquired after caesarian section. In addition, there are regional differences within the placenta. At the periphery, invasion is usually superficial as compared to the central region. Although morphological analyses are very informative, the value of these studies can be vastly increased when they are performed in the context of immunolocalization experiments. In this case, samples of the basal plate that contain spiral arterioles are collected and quickly processed following elective pregnancy termination to prevent degradation of the tissue. The vessels and the immediately adjacent decidua, 2 to 3 mm in diameter (Fig. 12.4A), are dissected free of the remaining tissue and fixed in 3% paraformaldehyde for 30 min. Then, the tissue is washed three times in PBS (4  C), infiltrated with gradually increasing concentrations of sucrose (5 to 15%) followed by OCT compound and freezing in liquid nitrogen. Placental tissues are processed using immunolocalization protocols that we published previously (Zdravkovic et al., 2006). Nonspecific antibody reactivity CTB modified blood vessel

A

B

CTBs SA

20 X

CK7/DAPI

Figure 12.4 Spiral arterioles and a histological section of the uterus that contains a CTB-modified blood vessel: (A) Spiral arterioles dissected from the decidua at 12 weeks of gestation. (B) Frozen section of an 18-week basal plate biopsy containing a CTBmodified blood vessel. CTBs were distinguished from other cell types by staining for cytokeratin 7 (CK7) (red); nuclei were visualized by staining with DAPI (blue).

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is blocked by incubation in 3% bovine serum albumin in PBS for 1 h. Primary antibodies for identification of CTBs in the basal plate include a rat anti-human cytokeratin 8 and 18 produced in our laboratory and a mouse anti-human cytokeratin 7 (Dako). The antibodies used for the identification of endothelial cells in incompletely remodeled blood vessels include a rabbit anti-human von Willebrand factor (Dako). CTB remodeling of large bore vessels often leaves no endothelial cells behind, and therefore, antibodies that recognize smooth muscle actin may help to visualize blood vessel remnants, although it should be noted that the muscular walls of the arterioles are also disrupted by endovascular invasion. Cytokeratin-positive glandular epithelial cells are often present in earlier samples from the first and second trimester. In such cases, endovascular CTBs can be distinguished from glands by virtue of their HLA-G expression (McMaster et al., 1995). Glands may also be distinguished morphologically by their columnar epithelial organization relative to the more stellate, disorganized distribution of endovascular CTBs. Tissue sections are incubated with primary antibodies overnight at 4  C, washed three times in PBS and incubated in the appropriate species-specific FITC- or rhodamineconjugated secondary antibodies. Negative control sections are incubated in the absence of primary antibodies. After staining, antibody reactivity is photographed using a Leica DM 500B fluorescence microscope (Leica Microsystems, Wetzlar, Germany). An example of a CTB-remodeled uterine spiral arteriole is shown in Fig. 12.4B. In this case, the tissue section was stained with antibodies that specifically reacted with cytokeratin 7 and nuclei were visualized by staining with DAPI.

5. Cytotrophoblast Migration and Induction of Endothelial Cell Apoptosis During Co-Culture Given the limited number of approaches available to study interactions between CTBs and host endothelial cells in vivo, several groups have attempted to model such behavior with co-culture systems. While the unique experimental conditions differed, for example, in the use of freshly isolated cells versus choriocarcinoma cell lines and two-dimensional versus three-dimensional culture conditions, several commonalities have emerged in the results. First, CTBs migrate toward and adhere to the endothelial cells. Second, the state of the endothelium, such as activation or differentiation, influences CTB behavior. Third, CTBs can induce apoptosis of the cultured endothelial cells. The methods used in these studies will be described with results highlighting the CTB behaviors that were observed.

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5.1. Co-culture of Jar choriocarcinoma cells with endothelial monolayers Rather than using freshly isolated cells, which are costly to prepare and have a limited lifespan in culture, the authors used choriocarcinoma cells (Chen et al., 2005; Chen et al., 2007). As background, choriocarcinoma arises during or after pregnancy with the common feature that the abnormal cells are derived from CTB progenitors (reviewed in Benirschke and Kaufmann, 2000; Shih and Kurman, 2002). This neoplasm is characterized by solid sheets of CTBs and syncytial trophoblasts with the distinctive features of aggressive vascular invasion (Benirschke and Kaufmann, 2000), frequent metastases (Berkowitz and Goldstein, 1996), and human chorionic gonadotropin (hCG) production. While it is certain that malignant choriocarcinoma cells do not replicate all the behaviors exhibited by primary CTBs in culture, they do share many of their properties, and therefore, are widely used as models in the study of CTB biology. As to the methods of this study, human umbilical vein endothelial cells (HUVECs) and human microvascular endothelial cells (HMECs) were stained with green cell tracker dye (CMFDA) and seeded (5
104cells/ml) onto plastic cover slips in six-well plates. Once confluent, Jar cells (2.5
104 cells/ml) that had been labeled with a red cell tracker dye (SNARF) were added to the endothelial cultures. Interactions among the cells were characterized at multiple time points up to 48 h. The Jar cells displaced the endothelial cells from the surface of the dish, a phenomenon that occurred progressively over time. Further investigation revealed that the choriocarcinoma cells induced apoptosis in the endothelial cells, which they phagocytosed. Interestingly, activation of the endothelial cells by addition of TNF alpha, interferon gamma, or necrotic cells prevented their displacement by Jar cells as well as their apoptotic cell death. These results suggest that the state of the endothelium regulates CTB– endothelial interactions, a novel observation.

5.2. Co-culture of first-trimester CTBs with differentiated endometrial endothelial cells in a three-dimensional model Aldo and colleagues (2007) tracked CTB migration in a three-dimensional matrix environment that promoted endothelial differentiation and tube formation. Human endometrial endothelial cells (HEECs) were isolated from endometrium by virtue of their affinity for biotinylated Ulex europaeus lectin and grown in EMB-2 medium (Cambrex). The HEECs were immortalized by retroviral transfection of telomerase (Krikun et al., 2004). For endothelial tube formation, 300 ml of Matrigel were added to 24-well tissue culture plates and polymerized at 37  C for 30 min. HEECs were stained with a green fluorescent dye PKH67 (Sigma-Aldrich) and added to

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the Matrigel-coated wells at a concentration of 100,000 cells per well. Endothelial tube formation occurred over the next 4 to 8 h in culture. First-trimester CTBs, isolated as described in Section 2, were labeled with a red fluorescent dye, PKH26 (Sigma-Aldrich), and seeded onto the endothelial tubes, 80,000 cells per well. Over the next 24 to 96 h, cell interactions were observed by fluorescence microscopy and photographed. CTBs migrated toward the endothelial tubes within 2 h and were either positioned on top or had intercalated within the lumina after 8 h. Interestingly, intercalation within the tubes was blocked by LPS activation of the endothelial cells prior to CTB addition. Using similar methods as those described above, other investigators have shown by time-lapse microscopy that both primary CTBs and CTB cell lines can induce endothelial cell and smooth muscle cell apoptosis through Fas/FasL signaling (Ashton et al., 2004; Harris, et al., 2006). A role for tumor necrosis factor a–related apoptosis inducing ligand (TRAIL) signaling in this process has also been described (Keogh et al., 2007).

6. In Vitro Models of CTB Endovascular Invasion Using Explanted Spiral Arterioles While cell culture models of CTB–endothelial interactions are very useful, they fail to provide many elements that are present during endovascular invasion in vivo, such as the stromal and extracellular matrix factors present in decidual tissue, the complex vessel architecture, and the variety of cell types present in this complex milieu. Additionally, endothelial and CTB cell lines used in the studies described above may not possess the same characteristics as their living equivalents. Therefore, several investigators have attempted to model CTB invasion using explanted tissues. Three unique approaches will be presented. In the first, spiral arterioles were dissected from myometrial biopsies obtained at term and infused with CTBs (Ashton et al., 2004; Cartwright, et al., 2002; Crocker et al., 2005). In the second, explanted spiral arterioles were denuded of endothelium to expose the smooth muscle lining (Harris, et al., 2006; Keogh et al., 2007). In the last, chorionic villous explants were co-cultured with explanted decidual tissue under conditions of hypoxia (Dunk et al., 2004).

6.1. CTB invasion of term myometrial spiral artery segments in vitro Spiral arteries were dissected under sterile conditions from myometrial biopsies obtained following caesarian section (Ashton et al., 2004; Cartwright, et al., 2002; Crocker et al., 2005). In choosing this approach, it is important to keep in mind that CTB invasion does not normally

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progress beyond the inner third of the myometrial segments of spiral arterioles. These arterial explants, which included adjacent decidua, allowed study of both the interstitial and endovascular components of CTB invasion. Either a CTB cell line (SGHPL-4) or primary CTBs, isolated from first-trimester tissue (see Section 2), were fluorescently labeled with CellTracker Orange (Invitrogen, cat. # C2927). To assay interstitial invasion, the ends of the spiral arterioles were tied off to prevent CTB infiltration along a luminal path. Ligated arteries were embedded in fibrin gels. To prepare the gels, fibrinogen (Sigma) was suspended in PBS (2.5 mg/ml) with 200 U/ml aprotinin (Trasylol, Bayer, Germany) in a six-well plate. Polymerization of the gel was induced by the addition of thrombin (0.625 U/ml). A total of 5
104 fluorescently labeled CTBs were resuspended in large EC growth medium (TCS Biologicals) at a concentration of 105 cells/ml and added to the top of the fibrin gel containing the arterial explant. Cultures were incubated for 5 days, and the fresh unfixed explants were embedded in OCT and 5-mm and 10-mm sections were prepared. To assay endovascular invasion, spiral arterioles were placed in a pressurized perfusion chamber (Living Systems), cannulated, and the lumina were maintained by exerting low pressure. Control vessel were perfused with medium alone (10 ml) and experimental vessel were exposed to medium containing fluorescently labeled CTBs (5
104 in 10 ml). Then, vessels ends were tied off and the explants were embedded in fibrin gels, incubated for 5 days, and analyzed as described previously. Regarding results, in the original publication describing this model (Cartwright et al., 2002), the authors observed CTB endovascular invasion through both interstitial and lumenal routes, with CTBs intercalating within the vessel wall and lining the lumen, respectively. Invasion was accompanied by apoptosis and loss of the resident endothelial cells as revealed by TUNEL staining and the presence of cleaved PARP within the remaining endothelial cells (Ashton et al., 2004). In later work, immunohistochemical analyses and electron microscopy (Crocker et al., 2005) revealed that CTBs disrupted both the endothelial and smooth muscle compartments, although the overall integrity of the vessel was not compromised. Vessels perfused with CTBs often lacked endothelium and showed reduced staining for smooth muscle actin. CTBs adherent to the vessel wall occasionally exhibited cellular processes extending deep into the smooth muscle layer. Invasion, which depended on oxygen concentration, was reduced in hypoxia. The same group studied interactions between CTBs and spiral arterioles that were denuded of endothelium by passing a column of air through the cannulated vessels (Harris, et al., 2006; Keogh et al., 2007). Perfusion of either CTBs or CTB-conditioned medium induced apoptosis in the smooth muscle wall of these vessels. Additional experiments showed that this phenomenon depended on Fas/FasL signaling (Harris, et al., 2006) and TRAIL signaling (Keogh et al., 2007).

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6.2. Chorionic villous CTBs invade first-trimester decidual explants in vitro Dunk and colleagues (2004) devised a novel model to study endovascular invasion. Chorionic villi were co-cultured with decidua parietalis explants from the same pregnancy. Experiments were conducted under conditions of physiological hypoxia, which were required for CTB invasion. This is in contrast to the experiments described previously in which hypoxia inhibited CTB invasion of spiral arterioles. As to methods, tissue samples were collected following elective termination of pregnancies, 6 to 9 weeks of gestation. Chorionic villous explants were prepared as described in Section 3, and co-cultured with decidual tissue that was dissected into cubes of approximately 2 to 3 mm3. Millicell-CM culture inserts with 0.4-mm pores (Millipore) were coated with 200 ml of Matrigel (BD Biosciences) and incubated at 37  C until the Matrigel polymerized. Decidual explants were transferred to the culture insert, epithelial surface facing upward, and placental villi added. Culture medium was changed every 48 h and consisted of a 50/50 mix of DMEM/Ham’s F12 medium supplemented with 10% FBS, 100 U/ml penicillin, 100 mg/ml streptomycin, and 0.25 mg/ml ascorbic acid (pH 7.4). Cultures were incubated under conditions of physiological hypoxia (3% oxygen) with 5% CO2 and were maintained up to 6 days. Decidual cultures without villous explants served as controls. Following termination of the experiments, tissues were fixed in 4% paraformaldehyde and 5-mm sections were prepared. Immunostaining for trophoblast, endothelial, and smooth muscle markers revealed that CTBs invaded through the decidua (interstitial invasion) and colonized blood vessels (endovascular invasion), which lost most of their resident endothelial cells. Those remaining often appeared to be partially detached from the underlying basement membrane. In addition, CTB-modified vessels showed reduced staining for smooth muscle actin. In contrast, decidual explants cultured without placental villi contained vessels with intact endothelial and smooth muscle layers. Interestingly, when experiments were carried out in a hyperoxic environment (17% oxygen) CTB invasion of the decidual explants was not observed.

7. In Vivo Models of Human Cytotrophoblast Vascular Remodeling During human pregnancy, placental CTBs of fetal origin invade the uterine wall. This process has two components. In the first, CTBs invade the uterine parenchyma where they interact with the stromal compartment and a resident maternal immune population. In the second, a subpopulation

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of CTBs invades uterine blood vessels with subsequent colonization of the arterial side of the circulation. Although some details are known about the molecular bases of these unique processes, a great deal remains to be learned. One reason is the lack of in vivo models for studying these unique heterotypic cell–cell interactions. For example, the mechanisms whereby CTBs replace the maternal endothelial lining of uterine arterioles and intercalate within the surrounding smooth muscle layer are difficult to study in vitro and almost certainly do not recapitulate the complex series of events that occur in vivo. Thus, the mechanisms of CTB vascular remodeling are of considerable interest.

7.1. Placental xenograft of the murine mammary fat pad and kidney capsule In a recent study, we devised a novel in vivo model to study CTB invasion in which human placental chorionic villi are implanted into the mammary fat pads or under the kidney capsules of SCID mice (Red-Horse et al., 2006). These in vivo models were very successful, as CTBs exhibited robust interstitial and endovascular invasion in both regions. Within the highly vascularized kidney capsule, CTBs invaded deeply, moving along the surfaces of capillaries. Xenograft in the mammary fat pad, which contains large-bore vessels, was more suited for the analysis of CTB interactions with arterioles. These experiments yielded a variety of interesting information such as the observation that CTB invasion of arterioles in this region was associated with endothelial apoptosis. The success of these experiments was not surprising given that CTBs exhibit invasive behavior in a variety of extrauterine sites. As examples, ectopic pregnancies can occur in the fallopian tube, abdominal wall, cervix, and ovary (Molinaro and Barnhart, 2007). In addition, choriocarcinomas frequently metastasize to the lungs, brain, and liver (Cheung, 2003). Regarding the methods, protocols involving animals and human fetal tissue were approved by the UCSF Committee on Animal Research and the UCSF Committee on Human Research, respectively. Homozygous C.B-17 scid/scid mice (Taconic) were the recipients. Anchoring placental villi were dissected into 2- to 5-mm pieces (as described in Section 3). Prior to surgery, mice were anesthetized with isoflurane anesthesia or ketamine/zylazine for mammary or kidney capsule procedures, respectively. For transplantation to the mammary fat pad, placental villi were surgically placed in a small incision made below this region, which was subsequently sutured. For transplantation under the kidney capsule, a small incision was made through the skin covering the lumbar area and the organ was exteriorized. Placental villi were implanted under the capsular membrane using techniques that have been described previously (McCune et al., 1988; Stoddart et al., 2001). Following both mammary fat pad and kidney capsule surgeries, mice were maintained

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under pathogen-free conditions for 1 to 3 weeks, at which time the experiments were terminated and either whole kidneys containing implants or implants dissected from a portion of the fat pads were immediately placed in 3% paraformaldehyde. Tissues were fixed for 3 h at 4  C before being passed through a sucrose gradient, snap-frozen in liquid nitrogen, and sectioned. Histological analyses allowed quantitative estimates of invasion that were made by calculating the distance between the implanted villi and the outer perimeter of placental cells that had invaded murine tissues.

7.2. Choriocarcinoma xenograft of the murine skin Another group reported an alternative model for investigating CTB invasive behavior (Grummer et al., 1999). Rather than studying primary CTBs or explants isolated from placental tissue, this group used the tumorigenic properties of the Jeg-3 human choriocarcinoma cell line, which was injected subcutaneously in the murine flank. In this location, Jeg-3 cells established tumors that grew rapidly within 2 weeks. Regarding endovascular invasion, two primary observations were made. First, Jeg-3 tumors had lacunar blood spaces that appeared to be the result of tumor cells colonizing host vessels, and Von Willebrand factor–positive endothelial cells and hCG-positive Jeg-3 tumor cells lined the blood-filled lacunae. Second, Jeg-3 cells replaced the host endothelium at junctional zones between the tumor and the host tissues. Taken together, these results indicate that Jeg-3 cells remodel the host vasculature. Regarding methods, the experiments were carried out in athymic nude mice (Han:NMRI nu/nu) maintained under pathogen-free conditions. Prior to transfer into 10-week-old recipient mice, Jeg-3 cells were grown to 80 to 90% of confluence and detached with trypsin/EDTA. One million cells in a volume of 200 ml were injected into the subcutaneous space of the flanks of male mice. To reduce the numbers of animals required, each mouse received injections on both sides of its body. Tumor tissue was collected at the termination of experiments 21 days following injection. For histological studies, tumor and adjacent host tissues were subject to one of the following fixation procedures. Some tissues were immediately embedded in OCT medium and frozen in liquid nitrogen. Others were fixed in 3% paraformaldehyde and processed in paraffin. Alternatively, specimens were fixed in 2.5% glutaraldehyde and embedded in Epon/ Araldite for fine structural analysis.

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Ashton, S. V., Whitley, G. S., Dash, P. R., Wareing, M., Crocker, I. P., Baker, P. N., and Cartwright, J. E. (2005). Uterine spiral artery remodeling involves endothelial apoptosis induced by extravillous trophoblasts through Fas/FasL interactions. Arterioscler. Thromb. Vasc. Biol. 25, 102–108. Blankenship, T. N., and King, B. F. (1996). Macaque intra-arterial trophoblast and extravillous trophoblast of the cell columns and cytotrophoblastic shell express neural cell adhesion molecule (NCAM). Anat. Rec. 245, 525–531. Blankenship, T. N., and Enders, A. C. (1997). Expression of platelet-endothelial cell adhesion molecule-1 (PECAM) by macaque trophoblast cells during invasion of the spiral arteries. Anat. Rec. 247, 413–419. Brosens, I. A., Robertson, W. B., and Dixon, H. G. (1972). The role of the spiral arteries in the pathogenesis of preeclampsia. Obstet Gynecol Annu. 1, 177–191. Burton, G. J., Jauniaux, E., and Watson, A. L. (1999). Maternal arterial connections to the placental intervillous space during the first trimester of human pregnancy: The Boyd collection revisited. Am. J. Obstet. Gynecol. 181, 718–724. Cartwright, J. E., Kenny, L. C., Dash, P. R., Crocker, I. P., Aplin, J. D., Baker, P. N., and Whitley, G. S. (2002). Trophoblast invasion of spiral arteries: A novel in vitro model. Placenta 23, 232–235. Chen, Q., Stone, P. R., McCowan, L. M., and Chamley, L. W. (2005). Interaction of Jar choriocarcinoma cells with endothelial cell monolayers. Placenta 26, 617–625. Chen, Q., Stone, P. R., McCowan, L. M., and Chamley, L. W. (2007). Activated endothelial cells resist displacement by trophoblast in vitro. Placenta 28, 743–747. Cheung, A. N. (2003). Pathology of gestational trophoblastic diseases. Best Pract. Res. Clin. Obstet. Gynaecol. 17, 849–868. Crocker, I. P., Wareing, M., Ferris, G. R., Jones, C. J., Cartwright, J. E., Baker, P. N., and Aplin, J. D. (2005). The effect of vascular origin, oxygen, and tumour necrosis factor alpha on trophoblast invasion of maternal arteries in vitro. J. Pathol. 206, 476–485. Damsky, C. H., Fitzgerald, M. L., and Fisher, S. J. (1992). Distribution patterns of extracellular matrix components and adhesion receptors are intricately modulated during first trimester cytotrophoblast differentiation along the invasive pathway, in vivo. J. Clin. Invest. 89, 210–222. Dunk, C., Petkovic, L., Baczyk, D., Rossant, J., Winterhager, E., and Lye, S. (2003). A novel in vitro model of trophoblast-mediated decidual blood vessel remodeling. Lab. Invest. 83, 1821–1828. Ellis, S. A., Palmer, M. S., and McMichael, A. J. (1990). Human trophoblast and the choriocarcinoma cell line BeWo express a truncated HLA Class I molecule. J. Immunol. 144, 731–735. Enders, A. C., and Blankenship, T. N. (1997). Modification of endometrial arteries during invasion by cytotrophoblast cells in the pregnant macaque. Acta Anat. (Basel) 159, 169–193. Even-Ram, S., Uziely, B., Cohen, P., Grisaru-Granovsky, S., Maoz, M., Ginzburg, Y., Reich, R., Vlodavsky, I., and Bar-Shavit, R. (1998). Thrombin receptor overexpression in malignant and physiological invasion processes. Nat. Med. 4, 909–914. Fisher, S. J., Cui, T. Y., Zhang, L., Hartman, L., Grahl, K., Zhang, G. Y., Tarpey, J., and Damsky, C. H. (1989). Adhesive and degradative properties of human placental cytotrophoblast cells in vitro. J. Cell Biol. 109, 891–902. Genbacev, O. D., Prakobphol, A., Foulk, R. A., Krtolica, A. R., Ilic, D., Singer, M. S., Yang, Z. Q., Kiessling, L. L., Rosen, S. D., and Fisher, S. J. (2003). Trophoblast L-selectin–mediated adhesion at the maternal–fetal interface. Science 299, 405–408. Grummer, R., Donner, A., and Winterhager, E. (1999). Characteristic growth of human choriocarcinoma xenografts in nude mice. Placenta 20, 547–553.

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Harris, L. K., Keogh, R. J., Wareing, M., Baker, P. N., Cartwright, J. E., Aplin, J. D., and Whitley, G. S. (2006). Invasive trophoblasts stimulate vascular smooth muscle cell apoptosis by a fas ligand–dependent mechanism. Am. J. Pathol. 169, 1863–1874. Keogh, R. J., Harris, L. K., Freeman, A., Baker, P. N., Aplin, J. D., Whitley, G. S., and Cartwright, J. E. (2007). Fetal-derived trophoblast use the apoptotic cytokine tumor necrosis factor-alpha-related apoptosis-inducing ligand to induce smooth muscle cell death. Circ. Res. 100, 834–841. Kovats, S., Main, E. K., Librach, C., Stubblebine, M., Fisher, S. J., and DeMars, R. (1990). A class I antigen, HLA-G, expressed in human trophoblasts. Science 248, 220–223. Krikun, G., Mor, G., Alvero, A., Guller, S., Schatz, F., Sapi, E., Rahman, M., Caze, R., Qumsiyeh, M., and Lockwood, C. J. (2004). A novel immortalized human endometrial stromal cell line with normal progestational response. Endocrinology 145, 2291–2296. Librach, C. L., Werb, Z., Fitzgerald, M. L., Chiu, K., Corwin, N. M., Esteves, R. A., Grobelny, D., Galardy, R., Damsky, C. H., and Fisher, S. J. (1991). 92-kD type IV collagenase mediates invasion of human cytotrophoblasts. J. Cell Biol. 113, 437–449. MacCalman, C. D., Furth, E. E., Omigbodun, A., Bronner, M., Coutifaris, C., and Strauss, J. F., 3rd. (1996). Regulated expression of cadherin-11 in human epithelial cells: A role for cadherin-11 in trophoblast-endometrium interactions? Dev. Dyn. 206, 201–211. MacCalman, C. D., Getsios, S., and Chen, G. T. (1998). Type 2 cadherins in the human endometrium and placenta: Their putative roles in human implantation and placentation. Am. J. Reprod. Immunol. 39, 96–107. McCune, J. M., Namikawa, R., Kaneshima, H., Shultz, L. D., Lieberman, M., and Weissman, I. L. (1988). The SCID-hu mouse: Murine model for the analysis of human hematolymphoid differentiation and function. Science 241, 1632–1639. McMaster, M. T., Librach, C. L., Zhou, Y., Lim, K. H., Janatpour, M. J., DeMars, R., Kovats, S., Damsky, C., and Fisher, S. J. (1995). Human placental HLA-G expression is restricted to differentiated cytotrophoblasts. J. Immunol. 154, 3771–3778. Miller, R. K., Genbacev, O., Turner, M. A., Aplin, J. D., Caniggia, I., and Huppertz, B. (2005). Human placental explants in culture: Approaches and assessments. Placenta 26, 439–448. Molinaro, T. A., and Barnhart, K. T. (2007). Ectopic pregnancies in unusual locations. Semin. Reprod. Med. 25, 123–130. Perry, E. (1978). The Realities of Primary Nursing Care: Needed Changes in Education. National League for Nursing (NLN), New York, N.Y. pp. 23–28. Prakobphol, A., Genbacev, O., Gormley, M., Kapidzic, M., and Fisher, S. J. (2006). A role for the L-selectin adhesion system in mediating cytotrophoblast emigration from the placenta. Dev. Biol. 298, 107–117. Queenan, J. T., Jr., Kao, L. C., Arboleda, C. E., Ulloa-Aguirre, A., Golos, T. G., Cines, D. B., and Strauss, J. F., 3rd. (1987). Regulation of urokinase-type plasminogen activator production by cultured human cytotrophoblasts. J. Biol. Chem. 262, 10903–10906. Ramsey, E. M., Houston, M. L., and Harris, J. W. (1976). Interactions of the trophoblast and maternal tissues in three closely related primate species. Am. J. Obstet. Gynecol. 124, 647–652. Red-Horse, K., Kapidzic, M., Zhou, Y., Feng, K. T., Singh, H., and Fisher, S. J. (2005). EPHB4 regulates chemokine-evoked trophoblast responses: A mechanism for incorporating the human placenta into the maternal circulation. Development 132, 4097–4106. Red-Horse, K., Rivera, J., Schanz, A., Zhou, Y., Winn, V., Kapidzic, M., Maltepe, E., Okazaki, K., Kochman, R., Vo, K. C., Giudice, L., Erlebacher, A., et al. (2006). Cytotrophoblast induction of arterial apoptosis and lymphangiogenesis in an in vivo model of human placentation. J. Clin. Invest. 116, 2643–2652.

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Roth, I., Corry, D. B., Locksley, R. M., Abrams, J. S., Litton, M. J., and Fisher, S. J. (1996). Human placental cytotrophoblasts produce the immunosuppressive cytokine interleukin 10. J. Exp. Med. 184, 539–548. Shih, I. M., and Kurman, R. J. (1996). Expression of melanoma cell adhesion molecule in intermediate trophoblast. Lab. Invest. 75, 377–388. Shih Ie, M., and Kurman, R. J. (2002). Molecular basis of gestational trophoblastic diseases. Curr. Mol. Med. 2, 1–12. Starkey, P. M., Sargent, I. L., and Redman, C. W. (1988). Cell populations in human early pregnancy decidua: characterization and isolation of large granular lymphocytes by flow cytometry. Immunology 65, 129–134. Stoddart, C. A., Liegler, T. J., Mammano, F., Linquist-Stepps, V. D., Hayden, M. S., Deeks, S. G., Grant, R. M., Clavel, F., and McCune, J. M. (2001). Impaired replication of protease inhibitor-resistant HIV-1 in human thymus. Nat. Med. 7, 712–718. Vicovac, L., Jones, C. J., and Aplin, J. D. (1995). Trophoblast differentiation during formation of anchoring villi in a model of the early human placenta in vitro. Placenta 16, 41–56. Zhou, Y., Fisher, S. J., Janatpour, M., Genbacev, O., Dejana, E., Wheelock, M., and Damsky, C. H. (1997). Human cytotrophoblasts adopt a vascular phenotype as they differentiate. A strategy for successful endovascular invasion? J. Clin. Invest. 99, 2139–2151. Zhou, Y., McMaster, M., Woo, K., Janatpour, M., Perry, J., Karpanen, T., Alitalo, K., Damsky, C., and Fisher, S. J. (2002). Vascular endothelial growth factor ligands and receptors that regulate human cytotrophoblast survival are dysregulated in severe preeclampsia and hemolysis, elevated liver enzymes, and low platelets syndrome. Am. J. Pathol. 160, 1405–1423. Zhou, Y., Bellingard, V., Feng, K. T., McMaster, M., and Fisher, S. J. (2003). Human cytotrophoblasts promote endothelial survival and vascular remodeling through secretion of Ang2, PlGF, and VEGF-C. Dev. Biol. 263, 114–125.

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An In Vivo Experimental Model for Postnatal Vasculogenesis Juan M. Melero-Martin and Joyce Bischoff Contents 1. Introduction 2. Isolation of Blood-Derived Endothelial Progenitor Cells 2.1. Materials/reagents 2.2. Recipes 2.3. Procedures 3. Expansion and Characterization of Blood-derived Endothelial Progenitor Cells 3.1. Additional materials/reagents 3.2. Additional recipes 3.3. Procedures 4. Growth of Human Smooth Muscle Cells 4.1. Procedure 5. In vivo Vasculogenic Assay 5.1. Additional materials/reagents 5.2. Additional recipes 5.3. Procedures 6. Conclusion Acknowledgments References

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Abstract Rapid and complete vascularization of ischemic tissues and thick engineered tissues is likely to require vasculogenesis. Therefore, the search for clinically relevant sources of vasculogenic cells and the subsequent development of experimental models of vasculogenesis is of utmost importance. Here, we describe a methodology adapted from the Matrigel plug assay to deliver human blood–derived endothelial progenitor cells (EPCs) and mature smooth muscle cells (SMCs) subcutaneously into immunodeficient mice. One week after

Vascular Biology Program and Department of Surgery, Children’s Hospital, Boston, Harvard Medical School, Boston, Massachusetts Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03013-9

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2008 Elsevier Inc. All rights reserved.

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implantation, an extensive microvascular network composed of the human EPCs and SMCs is formed within the Matrigel. The presence of human EPC-lined lumens containing host erythrocytes can be seen throughout the implants indicating not only the formation (de novo) of a vascular network, but also the development of functional anastomoses with the host circulatory system. This is a very versatile assay that allows (1) dialing the final microvessel density by varying either the total number of cells in the original cell suspension or the ratio between EPCs and SMCs, (2) studying the effect of substituting another type of perivascular cell for mature SMCs or another type of endothelial cell, (3) tracking each of the implanted cell types by labeling (e.g., GFP tagging) prior to implantation, and (4) studying the effect of genetically modifying the cells prior to implantation. Additionally, this assay is relatively simple to perform and it does not require an incision or surgical procedure. This murine model of human vasculogenesis is ideally suited for studies on the cellular and molecular components of microvessel development, pathologic neovascular responses, and for the development and investigation of strategies to enhance neovascularization of engineered human tissues and organs.

1. Introduction Current strategies to generate vascular networks in vivo are based on our understanding of the mechanisms of blood vessel formation. During embryogenesis, blood vessels are first formed de novo by the patterned assembly of angioblasts in a process termed ‘‘vasculogenesis’’ (Cleaver and Melton, 2003; Flamme et al., 1997; Risau and Flamme, 1995). Thereafter, the formation of new blood vessels occurs primarily via the sprouting of endothelial cells (ECs) from pre-existing vasculature (angiogenesis) (Carmeliet, 2003; Conway et al., 2001; Risau, 1997). After birth and during adulthood, the formation of new blood vessels occurs mainly through angiogenesis in a tightly controlled process where EC proliferation and migration are regulated by secreted cytokines, surrounding cells, and extracellular matrix (Carmeliet, 2003; Risau, 1997). Postnatal vasculogenesis involves the recruitment of endothelial progenitor cells (EPCs) as well as an intricate collaboration with hematopoietic cells (De Palma et al., 2005; Grunewald et al., 2006; Yoder et al., 2007). While EPC-derived ECs line the vessel lumen, accessory cells support vessel formation through secretion of cytokines and alteration of the extracellular matrix (Carmeliet, 2003; Hiraoka et al., 1998; Stetler-Stevenson, 1999; Zentilin et al., 2006). Dysregulated angiogenesis and vasculogenesis have been implicated in the pathogenesis of numerous diseases including vascular retinopathies, rheumatoid arthritis, vascular tumors such as infantile hemangioma, and cancer (Folkman, 1995).

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Our understanding of angiogenesis as a mechanism of blood vessel formation has evolved, in part, from numerous experimental assays that recapitulate the multistep angiogenic processes (Carmeliet, 2003; Carmeliet and Jain, 2000; Folkman, 1995, 2007; Risau, 1997). These include in vitro assays for endothelial proliferation, migration and capillary morphogenesis and in vivo assays such as the corneal micropocket assay (Gimbrone et al., 1974), the chick chorioallantoic membrane (CAM) assay (Auerbach et al., 1974; Ausprunk et al., 1974, 1975; Folkman, 1974), and the Matrigel plug assay (Passaniti et al., 1992), among other models (for review, see Norrby, 2006). Angiogenesis in these in vivo models is driven by exogenously added factors or relies on ongoing developmental processes, as is the case in the CAM assay. In some sense, the corneal micropocket and the Matrigel plug assays model regenerative processes in that angiogenesis, an essential component of tissue regeneration, is promoted by the growth of new blood vessels toward an angiogenic signal or factor (Isner and Asahara, 1999; Isner et al., 1996; Lee et al., 2002; Li et al., 2005). The mechanisms governing vasculogenesis are studied primarily during embryonic development in several model organisms and in murine embryonic stem-cell–derived embryoid bodies in vitro. Experimental models for postnatal vasculogenesis are less numerous. However, there is extensive interest in postnatal vasculogenesis because of the potential application(s) in tissueengineering and regenerative therapies. One approach that stimulated our thinking was the work of Schechner and colleagues (2000), in which they showed the preassembly of human umbilical vein ECs (HUVECs) into nascent microvascular networks in vitro, and the ability of these preformed endothelial networks to remodel into functional vessels when implanted subcutaneously into immunodeficient mice. In this pioneering work, HUVECs were plated in three-dimensional (3D) collagen/fibronectin gels in vitro for 24 h during which time the HUVECs organized into cellular cords with nascent lumens. The endothelial networks embedded within the collagen/fibronectin gels were then implanted into immunodeficient mice and followed for up to 2 months. Blood vessels lined with human endothelial cells and filled with red blood cells were detected after 1 to 2 months, demonstrating the inherent ability of human endothelial cells to assemble into vascular networks that connect with the host (murine) blood vessels. The HUVECs provided an appropriate starting point for this model, but the cells required transduction of the antiapoptotic gene Bcl-2 to augment survival. Later studies by the same group showed that cord blood and adult blood–derived endothelial cells did not require Bcl-2 transduction to form human vessels in this model (Shepherd et al., 2006). Furthermore, Yoder and colleagues (2007) used this model as a functional assay to demonstrate the crucial distinction between EPCs, called ECFC in their study, and hematopoietic/monocytic cells that express endothelial markers and have been called early EPCs, angiogenic EPCs, or CFU-ECs. (EPCs formed vessels whereas the hematopoietic/

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monocytic cells did not.) In summary, although the Schechner model system was developed to study immune cell interactions with human endothelium, it has also provided a valuable tool for investigators to test endothelial cell populations for ‘‘vasculogenic’’ potential—that is, the ability to form, de novo, networks of new vessels that can connect with the pre-existing host vasculature (Koike et al., 2004; Schechner et al., 2000). The cellular requirements for building blood vessel networks for tissue engineering and tissue regeneration is an area of active investigation. We showed in 2004 that a combination of human EPC-derived endothelial cells and mature human smooth muscle cells (SMCs) assembled into microvessel-like structures in vitro when co-seeded on a biodegradable scaffold of polyglycolic acid/poly-L-lactic acid (PGA/PLLA). In the absence of SMCs, the ECs were viable and maintained endothelial characteristics, but did not undergo morphological rearrangements suggestive of lumen formation (Wu et al., 2004). This study indicated that although endothelial cells have an inherent ability to form vascular cords and lumens, SMCs are required in certain, perhaps less favorable, 3-D environments such as that presented by the PGA/PLLA mesh. It also suggested that a twocell system composed of ECs and a smooth muscle support cell might prove to be more robust and efficient for building vascular networks de novo. The assay we described here supports this concept. We showed that cord blood–derived EPCs combined with mature human smooth muscle as a single-cell suspension in Matrigel form functional human blood vessel networks within 7 days after implantation into nude mice (Melero-Martin et al., 2007). Cord blood EPCs alone or human SMCs alone do not form vessels and Matrigel without cells is inert. Vessel density can be increased or decreased, over a range from 10 to 100 vessels/mm2, by varying cell numbers. Human cells can be identified by human specific antibodies or by GFP labeling of the cells. Other sources of human ECs, adult peripheral blood–derived EPCs, HUVECs, and human dermal microvascular endothelial cells (HDMECs) also form vessels at 7 days when combined with human SMCs, albeit at lower microvessel densities at this 80:20 ratio. Hence, this approach offers a versatile, quantifiable, and relatively simple model system to study postnatal vasculogenesis in vivo using human cells. The model can be used to study the vasculogenic potential of other sources of human endothelial and perivascular cells, such as human embryonic stem cells differentiated into endothelial and smooth muscle lineages (GerechtNir et al., 2003; Kaufman et al., 2001; Levenberg et al., 2002; Levenberg et al., 2007; Wang et al., 2004). The model can be used to screen for antiand/or proangiogenic compounds. And finally, the model can be used to study the role(s) of specific genes in the formation and function of a vascular network composed of human endothelium. We refer to this experimental model as ‘‘postnatal vasculogenesis’’ because (1) vessels are formed from cells isolated from postnatal human

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blood and tissues, (2) vessels form in an adult animal, and (3) vessels do not arise from pre-existing vessels but instead from single cells suspended in Matrigel. However, angiogenesis plays an important role in this assay because connections to the murine vasculature are needed to achieve red blood cell–filled vessels, the functional read-out in this assay. Whether the ability of human vessels to form productive connections with host vessels should be considered an aspect of vasculogenesis and angiogenesis would likely generate a lively discussion.

2. Isolation of Blood-Derived Endothelial Progenitor Cells The discovery of EPCs in peripheral blood was exciting because it suggested a promising opportunity to noninvasively obtain large quantities of autologous ECs for either therapeutic vascularization or tissue engineering, both of which require some form of postnatal vasculogenesis. However, the process for obtaining blood-derived EPCs with the ability to form blood vessels in vivo has not been straightforward. Most of the original studies identified circulating EPCs as cells expressing CD34, CD133, and the VEGF receptor 2 (KDR) (Asahara et al., 1997; Peichev et al., 2000; Reyes et al., 2002; Shi et al., 1998). However, it is known now that these cellular markers are shared by hematopoietic cells that can be mobilized into circulation from the bone marrow to home sites of neovascularization (Rafii and Lyden, 2003; Yoder et al., 2007). Although the hematopoietic and endothelial cell types are fundamentally different, many studies have referred to blood- or bone-marrow–derived adherent cells that express progenitor and endothelial markers such as CD34, CD133 and VEGFR2þ cells as EPCs (Prater et al., 2007). The hematopoietic accessory cells have been referred to as ‘‘colony-forming units-ECs’’ (CFU-ECs) (Gehling et al., 2000), ‘‘circulating angiogenic EPCs’’ (Rehman et al., 2003), ‘‘early EPCs’’ (Gulati et al., 2003; Hur et al., 2004), and ‘‘colony-forming unitsHill (CFU-Hill)’’ (Hill et al., 2003). On the other hand, the cells with direct involvement as the cellular lining of the blood vessel lumen have been referred to as ‘‘late outgrowth ECs’’ (Lin et al., 2000), EPCs (Kaushal et al., 2001), ‘‘late EPCs’’ (Hur et al., 2004), and ‘‘endothelial colony-forming cells’’ (ECFCs) (Ingram et al., 2004). Thus, the term EPCs has been applied to blood and bone-marrow cells with hematopoietic and endothelial features. Despite the ambiguous terminology, the functional distinction between the two groups of cells is becoming clearer. Yoder and colleagues (2007) demonstrated, in an elegant study, that most of the cells that have been long referred to as EPCs are in fact descendants of hematopoietic stem cells (HSCs); the cells express functional activities of myeloid

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cells and have no ability to differentiate into functional ECs in perfused blood vessels in vivo. Cells with bona fide blood vessel–forming ability, which we refer to as EPCs, and Yoder and Ingram refer to as ECFCs (Ingram et al., 2004), comprise a very small population of the circulating cells. EPCs are found at a concentration of about two to five cells per milliliter in human umbilical cord blood, and at a concentration of about 0.05 to 0.2 cells/ml in adult peripheral blood (Ingram et al., 2004) and in the vessel wall (Ingram et al., 2005). Both the low frequency of EPCs in circulation and the lack of a unique set of distinctive cellular markers have made the isolation of EPCs by flow cytometry or other immunological techniques very challenging. As a result, the most successful methodology for isolating EPCs is based on methods similar to those originally reported for endothelial outgrowth from peripheral blood (Lin et al., 2000). In this method, adult peripheral blood mononuclear cells (MNCs) or human umbilical cord blood MNCs are collected and plated onto collagen-coated plates in endothelial-specific growth media (Ingram et al., 2004; Lin et al., 2000; Yoder et al., 2007). Nonadherent cells are discarded and EC-like colonies emerge from the adherent cell population 14 to 21 days after plating adult MNCs and 5 to 7 days after plating human umbilical cord blood MNCs. The colonies display a cobblestone appearance typical of ECs, can be plated as single cells and routinely expanded for over 70 population doublings (Ingram et al., 2004, 2005; Lin et al., 2000; Melero-Martin et al., 2007; Yoder et al., 2007). EPCs obtained by this methodology are phenotypically indistinguishable from cultured mature ECs in terms of cobblestone morphology and expression of adhesion molecules and receptors (Ingram et al., 2004, 2005; Lin et al., 2000; Melero-Martin et al., 2007; Yoder et al., 2007). However, in functional assays, EPCs exhibit enhanced migratory and proliferative activity compared to mature ECs derived from existing vasculature (Ingram et al., 2004; Khan et al., 2006; Melero-Martin et al., 2007). More importantly, our group and two other independent groups have shown that EPCs obtained by this methodology possess de novo vessel-forming ability in vivo (Au et al., 2008; Melero-Martin et al., 2007; Yoder et al., 2007), and therefore these EPCs constitute one of the cellular building blocks for our experimental in vivo model of vasculogenesis.

2.1. Materials/reagents Heparin solution (American Pharmaceutical Partners, cat. # 504011) 19-gauge butterfly needle (Kendall, cat. # 225174) Ficoll-Paque Plus (Amersham Pharmacia, cat. # 17-1440-02) 50-ml Accuspin tubes (Sigma-Aldrich, cat. # A2055) Ammonium chloride solution (StemCell Technologies, cat. # 07850) Endothelial basal medium, EBM-2 (Lonza, cat. # CC-3156)

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EGM-2 Singlequot supplements (Lonza, cat. # CC-4176) Glutamine-penicillin-streptomycin solution, 100 GPS (Mediatech, Inc., cat. # 30-009-CI) Fetal bovine serum, FBS (Hyclone, cat. # SV30014.03). Heat inactivated at 56  C for 30 min Bovine serum albumin, BSA (Sigma-Aldrich, cat. # A7906) Gelatin (Fisher, cat. # DF0143-17-9) CD31 Dynal beads solution (DYNAL, cat. # 111.55) Magnetic particle concentrator (DYNAL, cat. # 120.20) Trypsin-EDTA solution, 1 (Mediatech, Inc., cat. # 25-052-CI) Dulbecco’s phosphate buffered saline, PBS (Sigma-Aldrich, cat. # D5652) Glucose (Sigma-Aldrich, cat. # G6152) Sodium citrate (Sigma-Aldrich, cat. # S4641) Citric acid (Sigma-Aldrich, cat. # 251275) Sodium carbonate, Na2CO3 (Sigma-Aldrich, cat. # 223530) Human plasma fibronectin (FN) (Chemicon International, cat. # FC-010) Cloning rings, 150 ml (Sigma-Aldrich, cat. # C1059) Sterile double-distilled water, dH2O

2.2. Recipes PBS, 1 l 9.6 g of PBS (Sigma-Aldrich, cat. # D5652) 1 l of dH2O Autoclaved at 121  C for 30 min 6% ACD-A solution, 1 l 22.3 g of glucose 22 g of sodium citrate 8 g of citric acid 1 l of dH2O Isolation buffer (PBS/0.6% ACD-A/0.5% BSA), 500 ml 50 ml of 6% ACD-A solution 2.5 g of BSA 450 ml of PBS Filter sterilized with a 0.2-mm–pore size vacuum filter EBM-2/20% FBS, 500 ml (herein called EPC medium) 395 ml of EBM-2 100 ml of FBS (20% final) 5 ml of 100 GPS All the EGM-2 Singlequot supplements except for hydrocortisone (i.e., VEGF, hFGF-B, R-IGF-1, hEGF, Heparin, ascorbic acid, and GA1000) Filter sterilized with a 0.2-mm–pore-size vacuum filter, divided into 45-ml aliquots and freeze down (–20  C) until needed.

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Isolation medium, 100 ml 81.24 ml of EPC medium 3.76 ml of FBS (to maintain 20% final) 15 ml of autologous plasma (obtained as described below; 15% final) Filter sterilize with a 0.2-mm–pore-size vacuum filter 1% gelatin solution, 500 ml 5 g of gelatin 500 ml of PBS Autoclaved at 121  C for 30 min. Filter sterilized with a 0.2-mm–poresize vacuum filter FN-coating solution (0.1 M Na2CO3), 500 ml 5.3 g of Na2CO3 500 ml of dH2O Adjust pH to 9.4 with HCl Filter sterilized with a 0.2-mm–pore-size vacuum filter

2.3. Procedures 2.3.1. Isolation of cord blood–derived EPCs (cbEPCs) 1. Coat 100-mm tissue-culture plates with 1% gelatin solution (10 ml per plate) and incubate at 37  C for 30 to 60 min. Prior to use, remove the gelatin solution and wash the plates once with PBS. 2. Set up 50-ml conical tubes with 10 ml of isolation buffer for blood collection. 3. Add 1 ml heparin solution into the syringe prior to drawing cord blood. Draw blood (typically 20 to 40 ml) from the umbilical vein using a 19-gauge butterfly needle. Collect every 25 ml of blood directly into 50-ml conical tubes with 10-ml isolation buffer. Place conical tubes with blood samples on ice. 4. Add 15 ml of Ficoll-Paque Plus to each 50-ml Accuspin tube. Spin at 1200 rpm for 1 min to sediment the Ficoll-Paque below the frit. 5. Add 30 ml of blood/isolation buffer to the top of each Accuspin tube. Spin at 2700 rpm for 15 min at room temperature (RT) with the brake of the centrifuge off. 6. Gently collect autologous plasma supernatant above the mononuclear cell layer. (Do not disturb the cell layer.) Save this autologous plasma to make up isolation medium. To minimize disturbances, do not remove all the plasma above the cell layer. 7. Using an 18-gauge needle on a 10-ml syringe, collect the mononuclear cell layer and transfer it to a 50-ml conical tube on ice. Add 5 ml of isolation buffer to every 10 ml of cells collected. 8. Spin the mononuclear cells at 2700 rpm for 5 min. Remove supernatant and resuspend the cell pellet in 10 ml of isolation buffer. Transfer to a

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15-ml conical tube and spin at 1200 rpm for 10 min. Remove supernatant. Add 1 ml of isolation buffer to dislodge the cell pellet and resuspend the cells. Add 3 ml of ammonium chloride solution to lyse erythrocytes. Incubate on ice for 5 to 10 min. Add 5 ml of isolation buffer and centrifuge at 1300 rpm for 5 min. Remove supernatant. If the cell pellet is not devoid of erythrocytes (i.e., pellet still has a red color), incubate again with 1 ml of isolation buffer and 3 ml of ammonium chloride solution on ice for 5 min. Add 5 ml of isolation buffer and centrifuge at 1300 rpm for 5 min. Remove supernatant. Resuspend the cell pellet in 10 ml of isolation medium per 25 ml of cord blood started with. Take 10 ml to count the cells in a hemocytometer and work out the total number of mononuclear cells per ml of cord blood sampled (approximately 4 to 8  107 MNCs in 25 to 50 ml of cord blood). Plate the mononuclear cell solution in 1% gelatin-coated 100-mm tissue culture plates. Use 2 100-mm tissue culture plates per 25 ml of cord blood sampled (equivalent to 4.5 cm2/ml cord blood sampled). Add 10 ml of cells suspended in isolation medium to each 10-cm plate, and place them in a humidified incubator at 37  C and 5% CO2 for 48 h. Forty-eight hours after plating, aspirate out the unbound cell fraction (which includes all the unattached hematopoietic cells), and feed the bound-cell fraction with fresh EPC medium. Feed the plates every 2 to 3 days with EPC medium. Screen plates for the presence of EC-like colonies. Cord blood–derived endothelial colonies that display cobblestone morphology will emerge in culture after 1 week (Fig. 13.1). The size, frequency, and time of appearance of these colonies will vary as reported by Ingram et al. (2004). Allow colonies to expand such that the plate is covered by a confluent cellular monolayer. Detach the cells using trypsin-EDTA solution and proceed to purify them by selection of CD31-positive cells.

2.3.2. Isolation of adult blood derived–EPCs (abEPCs) 1. Coat six-well tissue culture plates with 1% gelatin solution (2 ml per well) and incubate at 37  C for 30 to 60 min. Prior to use, remove the gelatin solution and wash the plates once with PBS. 2. Set up 50-ml conical tubes with 10 ml of isolation buffer for blood collection. 3. Add 1 ml heparin sodium into the syringe prior to drawing blood. Draw blood from the vein (typically 50 to 100 ml) using a 19-gauge butterfly needle. Collect 25 ml of blood directly into 50-ml conical tubes with 10 ml of isolation buffer. Place conical tubes with blood samples on ice.

A cbEPCs colony at day 7

cbEPCs at confluence

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Figure 13.1 Phenotypic characterization of EPCs. (A) EPCs emerge in culture as typical EC colonies, after 5 to 7 days when isolated from cord blood, and after 14 to 21 days when isolated from adult peripheral blood (left panels). CD31-selected EPCs present typical cobblestone morphology at confluence (right panels). Flow cytometric analysis of cultured (B) cbEPCs and (C) abEPCs show uniform expression of EC marker CD31, and negative expression of mesenchymal marker CD90 and hematopoietic markers CD45.This type of analysis should be performed routinely to verify that the cell population to be used for in vivo vasculogenesis is not contaminated with either mesenchymal or hematopoietic cells at any stage of their expansion in vitro.

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4. Add 15 ml of Ficoll-Paque Plus to each 50-ml Accuspin tube. Spin at 1200 rpm for 1 min to sediment the Ficoll-Paque to below the frit. 5. Add 30 ml of blood/isolation buffer to the top of each Accuspin tube. Spin at 2700 rpm for 15 min at RT with the brake of the centrifuge off. 6. Gently collect autologous plasma above the mononuclear cell layer. (Do not disturb the cell layer.) Save this autologous plasma to make up isolation medium. To minimize disturbances, do not remove all the plasma above the cell layer. 7. Using an 18-gauge needle on a 10-ml syringe, collect the mononuclear cell layer and transfer it to a 50-ml conical tube on ice. Add 5 ml of isolation buffer to every 10 ml of cells collected. 8. Spin the mononuclear cells at 2700 rpm for 5 min. Remove supernatant and resuspend the cell pellet in 10 ml of isolation buffer. Transfer to a 15-ml conical tube and spin at 1200 rpm for 10 min. Remove supernatant. 9. Add 1 ml of isolation buffer to dislodge the cell pellet and 3 ml of ammonium chloride solution to lyse erythrocytes. Incubate on ice for 5 to 10 min. 10. Add 5 ml of isolation buffer and centrifuge at 1300 rpm for 5 min. Remove supernatant. If cell pellet is not completely free of erythrocytes (i.e., pellet still has a red color), incubate it again with 1 ml of isolation buffer and 3 ml of ammonium chloride solution on ice for 5 min. Add 5 ml of isolation buffer and centrifuge at 1300 rpm for 5 min. Remove supernatant. 11. Resuspend cells in 12 ml of isolation medium per 25 ml of blood started with. Take 10 ml to count the cells in a hemocytometer and work out the total number of mononuclear cells per ml of blood sampled (approximately 10 to 15  107 MNCs in 50 ml of blood). 12. Plate the mononuclear cell solution in 1% gelatin-coated six-well tissue culture plates (2 ml per well). Use one six-well plate for each 25 ml of peripheral blood sampled (equivalent to 2.5 cm2/ml blood sampled). Place the plates in a humidified incubator at 37  C and 5% CO2. 13. Forty-eight hours after plating, add 2 ml of fresh isolation medium to each well. Do not aspirate the unbound cell fraction yet; place the plates back in the humidified incubator at 37  C and 5% CO2 for another 48 h. 14. Four days after plating, aspirate the unbound cell fraction and feed the bound cell fraction with fresh EPC medium (2 ml per well). 15. Feed the plates every 2 to 3 days with EPC medium. Screen plates for the presence of EC-like colonies. Peripheral blood-derived endothelial colonies (identified by typical cobblestone morphology) (Fig. 13.1) will emerge in culture after 2 to 3 weeks since isolation. The size, frequency, and time of appearance of these colonies will vary as reported by Ingram et al. (2004). Keep feeding the plates and leave the endothelial-like

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colonies to grow in size until they reach a diameter of approximately half the diameter of a cloning ring (3 mm). Before proceeding to collect the endothelial-like colonies, coat six-well tissue culture plates with 2 ml of FN-coating solution and 10 ml of human plasma fibronectin (final fibronectin concentration of 1 mg/ cm2) per well. Incubate the plates at 37  C for at least 60 min. Prior to use, remove the FN-coating solution and wash the plates once with PBS. Add 2 ml of EPC medium to each well where colonies will be plated. Each colony will be collected and replated in an individual well. Mark the location of the colonies of interest on the bottom of the wells with a marker. Verify that the colonies are circled by observing with an inverted microscope. Aspirate the medium from wells where colonies are present and wash each well with PBS. Aspirate the PBS and using a sterile forceps, place a cloning ring over the location of a colony to be selected and press down gently. The grease at the bottom of the cloning ring will help it stick to the culture plate. Add 100 ml of trypsin-EDTA solution inside each cloning ring and incubate at 37  C until the cells become loosely attached or detached, as observed in the inverted microscope. Transfer the 100 ml of cells in trypsin-EDTA from the cloning ring to the new FN-coated well with 2 ml of EPC medium. Add 100 ml of EPC medium to the cloning ring and gently resuspend any remaining cells inside the cloning ring; transfer this wash to the new FN-coated well with 2 ml of EPC medium. Finally, place the plates in the humidified incubator at 37  C and 5% CO2. Feed the plates every 2 to 3 days with EPC medium. At confluence, detach the cells using trypsin-EDTA solution and replate the cells from each well into one 100-mm, FN-coated tissue culture plate. Culture the plates in the humidified incubator at 37  C and 5% CO2. Allow cells to reach confluence. Detach the cells using trypsin-EDTA solution and proceed to purify them by selection of CD31-positive cells.

2.3.3. Magnetic bead purification of cultured CD31-positive EPCs 1. Coat 100-mm tissue-culture plates with 5 ml of FN-coating solution and 60 ml of human plasma fibronectin per plate (final fibronectin concentration of 1 mg/cm2). Incubate the plates at 37  C for at least 60 min. Before use, remove the FN-coating solution and wash the plates once with PBS. 2. For each confluent 100-mm cultured plate, aspirate the culture medium and wash the cells with 10 ml of PBS. Remove PBS and add 2 ml of trypsin-EDTA solution to each 100-mm plate. Gently rock the plates to evenly distribute the trypsin-EDTA solution. Incubate for 1 to 2 min. Gently tap the plate to facilitate cell detachment and verify under an inverted microscope that cells are detaching and in suspension.

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3. When cells completely detach, add 8 ml of EPC medium and collect the cell solution into a 15-ml conical tube. Take 10 ml to count the cells in a hemocytometer and work out the total number of cells harvested. 4. Centrifuge the cells at 1200 rpm for 5 min, remove the supernatant and wash the cell pellet in 5 ml of isolation buffer. Centrifuge the cells at 1200 rpm for 5 min, remove the supernatant and resuspend the cell pellet in 500 ml of isolation buffer. Transfer the cell solution into a 1.5-ml sterile centrifuge tube. 5. Add 12.5 ml of anti-CD31––coated Dynal beads and incubate for 5 min at 4  C with intermittent mixing. CD31-positive cells will attach to the anti-CD31–coated magnetic beads. 6. Place the centrifuge tube in the magnetic cell concentrator and hold for 1 min. Anti-CD31–coated magnetic beads will move toward the magnet (accumulating on the wall of the tube) leaving the nonattached cells (CD31-negative cell fraction) free in suspension. Gently aspirate the CD31-negative cell fraction. 7. Remove the centrifuge tube from the magnetic cell concentrator. Add 0.5 ml of isolation buffer and mix gently by pipetting. Place the tube back in the magnetic cell concentrator, hold for 1 min, and discard the negative fraction. Repeat the wash a total of three times. 8. After the final wash, resuspend the CD31-positive cells in 10 ml of EPC medium and plate the resulting cell solution on a 100-mm, FN-coated tissue culture plate. These cells can be referred to as passage 1 (P1) EPCs. 2.3.4. Notes 1. During the initial steps of the isolation procedure when the MNC fraction is plated on 100-mm dishes or 6-mm multiwell dishes, cells are fed with EPC medium with 15% of autologous plasma. 2. Because the EPCs comprise a very small population of the mononucleated cells, about two to five cells per milliliter in human umbilical cord blood and 0.05 to 0.2 cells per milliliter in adult peripheral blood (Ingram et al., 2004), the initial period of cell attachment is 2 days for cord blood samples and 4 days for adult peripheral blood samples. Thereafter, the unattached cells should be discarded and the cultures fed with EPC medium without autologous plasma. 3. As reported previously (Ingram et al., 2004; Yoder et al., 2007), EPC colonies should be evident 14 to 21 days after plating adult peripheral blood MNCs. However, since the number of expected colonies from adult peripheral blood is very low, a careful and thorough screening of the culture plates is advised, beginning in the second week after the procedure. To facilitate the screening (and the later picking of colonies), MNCs from adult blood samples can be divided among individual wells of a six-well plate instead of using 100-mm culture plates.

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4. Plates from adult peripheral blood preparations will present higher numbers of attached hematopoietic cells (e.g., monocytes and macrophages) than those from cord blood samples (Yoder et al., 2007). The myeloid cells will adhere to and populate the culture plates such that colonies of abEPCs will be left with limited free space to expand. Therefore, investigators are advised to pick the abEPC colonies with cloning rings and transfer the cells to new culture wells where they can be expanded freely from the hematopoietic cells. 5. Due to the inherent heterogeneity of blood samples, one concern about this method is the potential contamination with hematopoietic cells. However, hematopoietic cells that do attach to the culture plates are then difficult to detach during the trypsinization step. If such cells do detach, they do not proliferate appreciably in subsequent passages. Moreover, the superior proliferative capacity of EPCs over hematopoietic cells in culture will rapidly outpace any hematopoietic cells present at passage 1. In addition, the presence of hematopoietic cells is only significant in preparations from adult peripheral blood, wherein the EPC colonies are selected using cloning rings, thereby reducing the chances of contaminating cells. 6. An additional concern is the potential contamination with mesenchymal cells present in the blood samples. Circulating mesenchymal cells have been reported in adult peripheral blood (Simper et al., 2002) and cord blood samples (Kim et al., 2004; Le Ricousse-Roussanne et al., 2004; Lee et al., 2004). Therefore, blood-derived mesenchymal cells constitute a real potential contaminant since they can adhere and proliferate in the culture plates with ease. However, mesenchymal cells do not express the surface cell marker CD31, and therefore they will be depleted from the culture during the magnetic bead purification of CD31-positive EPCs.

3. Expansion and Characterization of Bloodderived Endothelial Progenitor Cells Prior to their use in vivo, EPCs will require a period of culture expansion in vitro. Although conditions for expanding EPCs can be found elsewhere in the literature, we have shown that up to 1013 cbEPCs (25 ml of cord blood) and 108 EPCs (50 ml of adult peripheral blood) can be obtained after only 40 days in culture following the methods described here (MeleroMartin et al., 2007). Similar expansion potential has been also reported by other authors for both blood-derived EPCs (Ingram et al., 2004; Lin et al., 2000). These remarkable numbers of human EPCs are likely to exceed (in the case of cord blood), and be sufficient (in the case of adult blood), the quantity that would be needed for most autologous regenerative therapies.

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However, it is important to consider the effect that culture conditions and the extent of expansion may impose on cellular phenotype and function. We and others have reported that EPCs expanded in vitro are phenotypically indistinguishable from cultured ECs (Ingram et al., 2004, 2005; Lin et al., 2000; Melero-Martin et al., 2007; Yoder et al., 2007). For example, flow cytometric analysis of EPCs shows uniform expression of EC markers CD31, VEGF-R2, and vWF, and negative expression of mesenchymal marker CD90 and hematopoietic markers CD45 and CD14. This type of analysis should be performed routinely to verify that the cell population to be used for in vivo vasculogenesis is not contaminated with either mesenchymal or hematopoietic cells at any stage of their expansion in vitro (Melero-Martin et al., 2007). Additionally, immunofluorescence staining should show that EPCs express CD31 and VE-cadherin at the cell–cell borders and vWF in a punctuate pattern in the cytoplasm, clear indications of EC properties (Melero-Martin et al., 2007). Despite the consistent and uniform expression of endothelial markers, EPCs can undergo cellular and functional changes in culture. For instance, cord blood–derived EPCs have been reported to change their morphology, growth kinetics, migration, proliferative responses toward angiogenic factors, and in vivo vasculogenic ability as they are expanded in culture (Ingram et al., 2004; Khan et al., 2006; Melero-Martin et al., 2007). Therefore, it is important to keep track of the number of population doublings that EPCs undergo in vitro prior to their use in vivo. In this section we provide details for (1) routine culture of EPCs, (2) evaluation of the accumulative number of population doublings during expansion, and (3) simplified phenotype confirmation by flow cytometry. In summary, we would like to stress the importance of these quality control measures to ensure that the cells to be used are homogenous and have expected endothelial features.

3.1. Additional materials/reagents High-glucose Dulbecco’s Modified Eagle Medium, 1 DMEM (Gibco, cat. # 10564) MEM nonessential amino acid solution (NEAA), 100 (Sigma-Aldrich, cat. # M7145) 0.5 M EDTA solution, pH 8.0 (Gibco, cat. # 15575-038) Mouse IgG1-PE (BD Pharmingen, cat. # 555787) Mouse IgG1-FITC (BD Biosciences, cat. # 349041) CD31-PE (Ancell, cat. # 180-050) CD90-PE (BD Pharmingen, cat. # 555596) CD45-FITC (BD Biosciences, cat. # 347463) Sterile double-distilled water, dH2O

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3.2. Additional recipes DMEM/10% FBS medium, 500 ml 440 ml of DMEM 50 ml of FBS (10% final) 5 ml of 100 NEAA 5 ml of 100 GPS Filter sterilized with a 0.2-mm–pore-size vacuum filter FACS buffer (PBS/0.5% BSA/2mM EDTA), 100 ml 9.6 ml of PBS 0.5 g of BSA 0.4 ml of 0.5 M EDTA solution

3.3. Procedures 3.3.1. Expansion of blood-derived EPCs Feed P1 blood-derived EPCs every 2 to 3 days using EPC medium (10 ml of medium per each 100-mm tissue-culture plate). At confluence, subculture the cells as follows: 1. Aspirate out the culture medium and wash the cells with 10 ml of PBS. 2. Remove PBS and add 2 ml of trypsin-EDTA solution to each 100-mm plate. Gently rock the plates to evenly distribute the trypsin-EDTA solution. Incubate for 1 to 2 min. Gently tap the plate to see the detached cells in suspension under an inverted microscope. 3. When cells completely detach, add 8 ml of EPC medium and collect the cell solution into a 15-ml conical tube. Take 10 ml to count the cells in a hemocytometer and work out the total number of cells harvested. Calculate the number of population doublings (PD) as follows:


Ln XF X0 PD ¼ Lnð2Þ where X0 (cells) refers to the initial cell number seeded in this culture, XF (cells) refers to the final cell number observed at the time of harvesting, and PD refers to the number of doublings that the cell population underwent during this passage. 4. Plate the cells in FN-coated (1 mg/cm2) tissue-culture plates at a seeding density of 5000 cell/cm2 using EPC medium. Place the plates in a humidified incubator at 37  C and 5% CO2 and feed them every 2 to 3 days with EPC medium. These cells can be referred to as passage 2 (P2) EPCs.

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Repeat this procedure for subsequent passages. Keep track of the accumulative value of PD as the cell population is expanded. 3.3.2. Cell characterization by flow cytometry (CD31, CD90, and CD45) 1. Aspirate out the culture medium from one confluent 100-mm culture plate and wash the cells with 10 ml of PBS. 2. Remove PBS and add 2 ml of trypsin-EDTA solution to each 100-mm plate. Gently rock the plates to evenly distribute the trypsin-EDTA solution. Incubate for 1 to 2 min. Gently tap the plate to see the detached cells in suspension under an inverted microscope. 3. When cells completely detach, add 8 ml of DMEM/10% FBS medium and collect the cell solution into a 15-ml conical tube. Take 10 ml to count the cells in a hemocytometer and work out the total number of cells harvested. 4. Centrifuge the cells at 1200 rpm (<300 g) for 5 min. Remove supernatant and resuspend the cell pellet in 10 ml of FACS buffer and centrifuge again at 1200 rpm (<300 g) for 5 min. Remove supernatant. 5. Resuspend the cell pellet in 500 ml of FACS buffer (100 ml per label). Split the cell sample by transferring 100 ml of the cell suspension into five individual 1.5-ml centrifuge tubes. Add 1 ml of conjugated antibody (IgG-FITC, IgG-PE, CD31-PE, CD90-PE, and CD45-FITC) and incubate for 20 min at 4  C with frequent mixing (tapping). 6. Following incubation, add 1 ml of FACS buffer to the tubes and centrifuge at 3000 rpm (800 g) for 3 min. 7. Wash the pellet in 1 ml of FACS buffer and centrifuge again. Repeat the wash once again. 8. Resuspend the cell pellets in 1% paraformaldehyde (made in PBS) and transfer the cells to FACS tubes. Keep the samples at 4  C in the dark up to 1 week before analysis. Flow cytometric analyses can be performed using standard instruments (e.g., a Becton Dickinson FACScan flow cytometer) and the collected data analyzed by specialized software (e.g., FlowJo software from Tree Star Inc.). 3.3.3. Notes 1. When expanding cells in culture, it is common among investigators to report the passage number of the cultured cells. However, it is more accurate to keep track of the number of population doublings (PD) to reflect the degree of expansion exerted on them. The functional changes that cultured cells experience as a result of their expansion in vitro are likely to correlate with the number of PD, but not necessarily with the number of passages. Two different investigators can easily reach the same number of PD after two different passage numbers.

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2. Anti-VE-cadherin (CD144) can be used in place of anti-CD31 to verify endothelial phenotype. PE-conjugated, anti-human VE-cadherin is available from R&D Systems. 3. Expanded EPCs can be cryo-preserved using standard methods (e.g., using 90% FBS and 10% DMSO as freezing medium and liquid nitrogen for storage). We have tested EPCs cryo-preserved at different passages (up to passage 15), and their phenotype as well as their in vitro and in vivo functions were properly maintained (Melero-Martin et al., 2007).

4. Growth of Human Smooth Muscle Cells This assay requires SMCs as a source of perivascular cells. Human SMCs can be isolated from discarded vascular tissues (e.g., human saphenous vein-derived SMCs). Additionally, investigators can purchase commercially available human SMCs (e.g., Cell Application, Inc. ScienCell Res Lab; Cascade Biologics; Lonza Inc.).

4.1. Procedure Plate SMCs in noncoated tissue-culture plates at a seeding density of 10,000 cells/cm2. Place the plates in a humidified incubator at 37  C and 5% CO2 and feed them every 2 to 3 days with DMEM/10% FBS medium (10 ml of medium per 100-mm tissue-culture plate). At 80% confluence, subculture the cells as follows: 1. Aspirate out the culture medium and wash the cells with 10 ml of PBS. 2. Remove PBS and add 2 ml of trypsin-EDTA solution to each 100-mm plate. Gently rock the plates to evenly distribute the trypsin-EDTA solution. Incubate for 1 to 2 min. Gently tap the plate to see the detached cells in suspension under an inverted microscope. 3. When cells completely detach, add 8 ml of DMEM/10% FBS medium and collect the cell solution into a 15-ml conical tube. Take 10 ml to count the cells in a hemocytometer and work out the total number of cells harvested. Calculate the number of PDs with the previous equation. 4. Plate the cells in noncoated tissue-culture plates at a seeding density of 10,000 cell/cm2 using DMEM/10% FBS medium. Place the plates in a humidified incubator at 37  C and 5% CO2 and feed them every 2 to 3 days with DMEM/10% FBS medium. Repeat this procedure for subsequent passages. Keep track of the cumulative value of PD as the cell population is expanded.

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5. In vivo Vasculogenic Assay The Matrigel plug assay was introduced by Passaniti and co-workers in 1992 (Passaniti et al., 1992), and is regarded as a useful assay for in vivo screening of potential pro- and anti-angiogenic compounds (Auerbach et al., 2003). Matrigel, which is an extract of the murine EngleberthHolm-Swarm tumor, is composed of basement membrane proteins. Although it takes the form of a liquid at 4  C, Matrigel reconstitutes into a gel or plug at body temperature when injected subcutaneously into mice, where it is progressively surrounded by granulation tissue. This assay was originally conceived as a mechanism to study the growth of new vessels into the Matrigel in response to an angiogenic factor. For instance, Matrigel plugs elicit an intense vascular response when supplemented with acidic FGF (Passaniti et al., 1992), basic FGF, or VEGF (Kano et al., 2005). One aspect of this assay that has been often criticized is the fact that Matrigel has not been fully defined chemically. It contains collagen IV, laminin, nidogen/entacin, heparin sulfate proteoglycan, and growth factors such as epidermal growth factor, transforming growth factor beta, platelet-derived growth factor, insulin-like growth factor-1, nerve growth factor, and bFGF (Baatout, 1997; Vukicevic et al., 1992). This suggests that caution should be exercised in the interpretation of experiments on cellular activities related to Matrigel (Vukicevic et al., 1992). However, subcutaneous implantation of Matrigel alone (without any additional angiogenic factor) does not initiate an angiogenic response from the host; when plugs of Matrigel alone are implanted for 1 to 2 weeks into immunodeficient mice, they remain largely inert with no vascular structures inside the implants and only a few cells invading the plugs (Melero-Martin et al., 2007). On the contrary, implantation of Matrigel containing EPCs and SMCs supports the progressive formation of vascular networks, reproducing aspects of postnatal vasculogenesis such as cellular assembly, lumen formation and network remodeling. Additionally, this assay is relatively simple to perform; it does not require an incision or surgical procedure, which reduces the potential influence of wound healing. Finally, the assay can be carried out in either athymic nu/nu or NOD/SCID mice.

5.1. Additional materials/reagents Phenol-red–free BD Matrigel Matrix (BD Bioscience, cat. # 356237) Six-week-old male athymic nu/nu mouse (Massachusetts General Hospital, Boston) Histological Tissue-Tek unicassette (Sakura Finetek, cat. # 4117-02) Isoflurane liquid for inhalation (Buxter Healthcare Corporation, cat. # NDC 10019-360-40) 10% neutral buffered formalin (Sigma-Aldrich, cat. # HT501128)

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5.2. Additional recipes Matrigel aliquots, 1ml Matrigel should be divided into aliquots as recommended by the manufacturer. Briefly, thaw 10 ml of Matrigel on ice overnight and transfer the liquid Matrigel into ten 1.5-ml centrifuge tubes (1-ml aliquots). Freeze the Matrigel aliquots at –20  C until needed. The day of the experiment, thaw the required aliquots on ice 1 to 2 h prior to their use.

5.3. Procedures Before the experiment, make sure that sufficient EPCs and SMCs are available in culture; 1,500,000 EPCs and 375,000 SMCs will be required for each implant and mouse. 1. Aspirate the medium of each culture plate and wash the cells with 10 ml of PBS. Remove PBS and add 2 ml of trypsin-EDTA solution to each 100-mm plate. Gently rock the plates to evenly distribute the trypsinEDTA solution. Incubate for 1 to 2 min. Gently tap the plate to see the detached cells in suspension under an inverted microscope. 2. When cells completely detach, add 8 ml of DMEM/10% FBS and collect the cell solution into a 15-ml conical tube. Take 10 ml to count the cells in a hemocytometer and work out the total number of EPCs and SMCs harvested. 3. Transfer 7,500,000 EPCs (5  1,500,000 cells) and 1,875,000 SMCs (5  375,000 cells) together into a single 50-ml conical tube. This is the total amount of cells required for five individual mice. Centrifuge at 1200 rpm and remove the supernatant. The total number of cells/ implant can be varied from one-third to three times this amount to achieve lower or higher microvessel density (Melero-Martin et al., 2007). Furthermore, the ratio of EPCs to SMCs can also be varied to achieve different degrees of vessel formation. 4. Resuspend the cell pellet on 1 ml of ice-cold Matrigel. Mix the cells very gently to avoid bubbles within the Matrigel. Load the cell–Matrigel mixture into a 1-ml sterile syringe, and place a 26-gauge needle with its cap on the tip of the syringe. Keep the loaded syringe deep on ice until injection. 5. Prior to the injection, anesthetize the immunodeficient mice by placing them in a gas chamber delivering isoflurane. Allow the mice to inhale the isoflurane for approximately 2 min until they are asleep (monitor their heart beats by inspection). For each mouse, inject 200 ml of the cell– Matrigel mixture subcutaneously into the upper dorsal region using a 26-gauge needle. Matrigel forms a gel at 37  C so that the implant should form a small bump just under the skin (Fig. 13.2). After the injection,

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7.

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place the mice on a layer of gauze for comfort and warmth and observe them until they become ambulatory. Then, observe the mice daily for the first 3 days. One week after the injections, euthanize the mice by placing them in a gas chamber delivering compressed CO2 gas. Once euthanized, cut open the skin near the area of the injection and surgically removed the Matrigel plug (Fig. 13.2). Digital photographs of the retrieved Matrigel plugs with a scale are advised. Place the harvested Matrigel plugs into histological cassettes and deep them into 10% neutral buffered formalin overnight at RT. After fixation, wash the 10% neutral buffered formalin away with dH2O and place the histological cassettes at 4  C in PBS until histological evaluation. For histological evaluation, the implants are embedded in paraffin and sectioned (7-mm–thick sections) using standard histological procedures. Standard protocols for hematoxilin and eosin (H&E) can be found elsewhere. Quantify microvessel density by evaluation of 10 randomly selected fields (0.1 mm2 each) of H&E stained sections taken from the middle part of the implants. Microvessels can be identified as lumenal structures containing red blood cells and counted (Fig. 13.3). Report microvessel density as the average number of red blood cell-filled microvessels from the fields analyzed and expressed as vessels per square millimeter; 93  18 vessels/mm2 correspond to the average values plus/minus standard deviation obtained from four individual mice using cbEPCs at passage 3 plus SMCs (Melero-Martin et al., 2007).

5.3.1. Notes 1. The time between resuspension of the cells in Matrigel and injection into the mice should be kept to a minimum (30 to 60 min).

Figure 13.2 Appearance of cell/Matrigel plugs. Human cord blood EPCs and smooth muscle cells were suspended in Matrigel as described in the text. Right panel shows location and appearance of a cell/Matrigel suspension implanted subcutaneously into a nude mouse. Middle panel shows appearance of the cell/Matrigel plug 7 days after implantation. The left panel shows the appearance of the cell/Matrigel plug removed from the mouse after 7 days and prior to processing for histology.

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A

B

H&E

50 mm

CD31

50 mm

Figure 13.3 In vivo vasculogenic potential of EPCs. Matrigel implants containing human EPCs and SMCs were evaluated after 1 week. (A) H&E staining of implants revealed the presence of an extensive network of microvessels containing red blood cells. (B) Immunohistochemical staining at 1 week with anti-human CD31 antibody revealed that the lumenal structures were formed by the implanted EPCs.

2. Note that in this assay it is difficult to generate identical 3D plugs, even though the total Matrigel volume is kept constant (Auerbach et al., 2000). Therefore, injections should be performed gently and the cell– Matrigel mixtures slowly released. In this regard, further improvements can be achieved by using subcutaneous chambers that allow for constant 3D form and volume of the Matrigel plug; these chambers have been reported to make the original Matrigel plug assay more reproducible in mice and rats (Kragh et al., 2003; Ley et al., 2004). 3. To further characterize the microvascular structures detected, sections of the retrieved Matrigel plug should be immunohistochemically stained with a human-specific CD31 antibody using standard staining protocols. Lumenal structures will stain positive for human CD31 (Fig. 13.3), confirming that those lumens are formed by the implanted human EPCs and not by the host cells. This evaluation is important because it demonstrates that the formation of microvascular vessels within the implant is the result of a process of in vivo vasculogenesis carried out by the implanted cells, and is not due to blood vessel invasion and sprouting (i.e., an angiogenic response from nearby host vasculature). We recommend using the monoclonal mouse anti-human CD31 antibody from DakoCytomation (Clone JC70A, cat. # M0823) at a 1:20 dilution. The human specificity of this antibody was confirmed by the negative reaction obtained with a diversity of mouse tissue sections that were stained in parallel (Melero-Martin et al., 2007). SMCs can be localized by immunostaining with anti-alpha smooth muscle actin (clone 1A4, Sigma). However, this antibody reacts with both murine and human alpha–smooth muscle actin.

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6. Conclusion Rapid and complete vascularization of ischemic tissues and thick engineered tissues are likely to require vasculogenesis. Therefore, the search for clinically relevant sources of vasculogenic cells and the subsequent development of experimental models of vasculogenesis are of utmost importance. Here, we describe a methodology adapted from the Matrigel plug assay to deliver human blood–derived EPCs and mature SMCs subcutaneously into immunodeficient mice. One week after implantation, an extensive microvascular bed that forms anastomoses with the host vasculature will be created by the implanted cells inside the plugs. The presence of human EPC–lined lumens containing host erythrocytes can be seen throughout the implants indicating not only the formation (de novo) of a vascular network, but also the development of functional anastomoses with the host circulatory system. Varying the number of cells in the original cell suspension can be used to manipulate the microvessel density achieved at 7 days. Altering the ratio between EPCs and SMCs, substituting another type of perivascular cell for mature SMCs, and GFP tagging the EPCs or perivascular cells are potential modifications (Melero-Martin et al., 2008). This murine model of human vasculogenesis is ideally suited for studies aimed at cellular and molecular components of microvessel development and pathologic neovascular responses, and for the development and investigation of strategies to enhance neovascularization of engineered human tissues and organs.

ACKNOWLEDGMENTS This research was supported by funding from the U.S. Army Medical Research and Material Command (W81XWH-05-1-0115).

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Stetler-Stevenson, W. G. (1999). Matrix metalloproteinases in angiogenesis: A moving target for therapeutic intervention. J. Clin. Invest. 103, 1237–1241. Vukicevic, S., Kleinman, H. K., Luyten, F. P., Roberts, A. B., Roche, N. S., and Reddi, A. H. (1992). Identification of multiple active growth factors in basement membrane Matrigel suggests caution in interpretation of cellular activity related to extracellular matrix components. Exp. Cell Res. 202, 1–8. Wang, L., Li, L., Shojaei, F., Levac, K., Cerdan, C., Menendez, P., Martin, T., Rouleau, A., and Bhatia, M. (2004). Endothelial and hematopoietic cell fate of human embryonic stem cells originates from primitive endothelium with hemangioblastic properties. Immunity 21, 31–41. Wu, X., Rabkin-Aikawa, E., Guleserian, K. J., Perry, T. E., Masuda, Y., Sutherland, F. W., Schoen, F. J., Mayer, J. E., Jr., and Bischoff, J. (2004). Tissue-engineered microvessels on three-dimensional biodegradable scaffolds using human endothelial progenitor cells. Am. J. Physiol. Heart Circ. Physiol. 287, H480–H487. Yoder, M. C., Mead, L. E., Prater, D., Krier, T. R., Mroueh, K. N., Li, F., Krasich, R., Temm, C. J., Prchal, J. T., and Ingram, D. A. (2007). Redefining endothelial progenitor cells via clonal analysis and hematopoietic stem/progenitor cell principals. Blood 109, 1801–1809. Zentilin, L., Tafuro, S., Zacchigna, S., Arsic, N., Pattarini, L., Sinigaglia, M., and Giacca, M. (2006). Bone marrow mononuclear cells are recruited to the sites of VEGF-induced neovascularization but are not incorporated into the newly formed vessels. Blood 107, 3546–3554.

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Assessment of Arteriogenesis Michael Simons Contents 1. Introduction 2. Anatomical Assessment of Arteriogenesis 2.1. High-resolution micro-computed tomography 2.2. Micro-CT coronary angiography 3. Microangiography 3.1. X-ray microangiography: Animal preparation and image acquisition (mice) 3.2. Coronary fluorescent microangiography: Animal preparation and image acquisition (mice) 4. Functional Assessment of Arteriogenesis 5. Laser-Doppler Perfusion Imaging 5.1. Animal preparation and data acquisition 6. Electron Paramagnetic Resonance Tissue Oxymetry 6.1. Electron paramagnetic resonance data acquisition 7. Magnetic Resonance Imaging 7.1. Time-of-flight magnetic resonance imaging data acquisition Acknowledgments References

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Abstract Assessment of arterial circulation development and function in mice is a frequent experimental challenge. A number of techniques including micro-CT angiography, fluorescent angiography, laser-Doppler perfusion imaging electron pramagnetic resonance and nuclear magnetic resonance can be used to assess the anatomic extent and functional state of an arterial circulation in a given organ. The chapter discusses the application of these tools in adult mice.

Section of Cardiology, Angiogenesis Research Center, Dartmouth Medical School, Hanover, New Hampshire Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03014-0

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1. Introduction Arteriogenesis is a process of arterial vasculature formation that occurs both during embryonic development and in adult tissues. During embryonic development arteriogenesis involves coating of newly formed arterial endothelial tubes by pericytes and, eventually, smooth muscle cells. In adult tissues, arteriogenesis can occur by either remodeling of pre-existing collateral arteries or by de novo arterial formation, most likely by arterialization of a subset of capillary vasculature (Carmeliet, 2005; Simons, 2005). The need to assess arteriogenesis and the state of arterial circulation arises in diverse settings. In one typical set of experiments, normal arterial circulation is disrupted and its recovery is then evaluated over time (Tirziu and Simons, 2005). Alternatively, a genetic manipulation of a particular protein involved in regulation of arteriogenic response may need to be assessed in mice (Chittenden et al., 2006) or the extent of circulation between different mouse strains compared (Helisch et al., 2006). Retina is also frequently used as a model of vascular, and in particular, arterial circulation development, but the technique of retinal circulation analysis will not be discussed here. Arterial blood flow is the primary determinant of tissue perfusion in higher organisms. Typically, reduction in arterial blood supply results in insufficient delivery of oxygen to tissues, thereby precipitating ischemia. The functional consequences of ischemia are a function of the organ in which it occurs and its severity. In skeletal muscle tissues or the heart, ischemia at first leads to decreased functional performance and may eventually manifest itself as cell death. In other tissues or organs, ischemia can also result initially in functional impairment, the nature of which will depend on the specific situation. At the same time, an increase in blood supply above normal levels rarely has a dramatic effect on organ function with exception of skeletal muscle where performance can increase above normal levels. Thus, assessment of the state of arterial circulation in any given organ or tissue relies on a combination of anatomical and functional techniques that can provide an insight into both the visual and quantitative extent of circulation development and its functional effectiveness. Among the anatomical tools, micro-CT imaging has received considerable recent attention due to its high spatial resolution (approaching 6 mm) and the ability to provide a quantitative assessment of the extent of vasculature formation. Other more traditional anatomical tools, such as polymer casting of the vasculature and angiography, suffer from being not particularly quantitative and a limited spatial resolution. On the other hand, in certain vascular beds, such as the retina, ex vivo staining of the arterial vasculature coupled with high-resolution microscopy is particularly effective.

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Functional assessment of arteriogenesis relies of assessment of blood flow in compromised tissues. This may involve laser Doppler imaging, electron paramagnetic resonance (EPR) measurements of tissue O2 levels, nuclear magnetic resonance (NMR) assessment of perfusion and blood flow velocity and relatively little currently used radioactive or fluorescent microspheres. These techniques are discussed in detail below.

2. Anatomical Assessment of Arteriogenesis The two main approaches are microangiography using either x-ray or fluorescent dyes and micro-computed tomography (CT) imaging. The latter provides much higher resolution but is much more technically challenging.

2.1. High-resolution micro-computed tomography The goal of micro-CT imaging is to visualize the arterial circulation and to quantitate the extent of arteriogenesis. Images are obtained as a series of two-dimensional (2D) slices along a vertical axis, generating a Z stack. This series of 2D images can then be rendered as a single three-dimensional (3D) image (Fig. 14.1) or the images can be displayed in their original forms. Quantification can proceed on the basis of either 2D or 3D data sets (Zhuang et al., 2006). The technique is very reproducible and can be used to obtain detailed anatomical information about the state of arterial circulation in various mouse tissues including the heart, extremities, kidneys, and brain (Chittenden et al., 2006; Li et al., 2006; Tirziu and Simons, 2005). Examples of micro-CT analysis of mouse hindlimb arterial circulation in normal animals and in animals in which vasculature growth was stimulated by a common femoral artery ligation is illustrated in Fig. 14.2, while normal renal arterial circulation is shown in Fig. 14.3. 2.1.1. Animal preparation and image acquisition (mice) 1. Inject mice with heparin (100 units) 5 min prior to euthanasia with isoflurane. 2. To ensure contrast perfusion, the inferior vena cava is cut and the vasculature is flushed with 0.9% normal saline containing papaverine (4 mg/l) and adenosine (1 g/l) via a needle inserted into the left ventricle. 3. Once flushed, 3.7% paraformaldehyde is infused over 5 min at 100 mmHg pressure.

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Figure 14.1 Generation and quantification of 3D micro-CT images. A 3D micro-CT image of a an ischemic (panel A) and normal (panel C) mouse hindlimb arterial circulation reconstructed at 16 mm. The axial slices (z-slices, 1250 cross-sections from thigh to calf region) that were used for quantitative analysis, are shown as red lines. Representative z-slices of the upper and lower limb region are shown in panel B. The white spots show vessels and sizes within a single section.

4. The descending aorta is cannulated and injected with 0.7 ml of solution containing 50% bismuth and 5% gelatin in normal saline over 2 min with a syringe pump. The mice are then immediately chilled on ice and immersion-fixed in 2% paraformaldehyde overnight. 5. For hindlimb vasculature image acquisition, mice are placed into a GE Healthcare micro-CT (GE eXplore Locus SP) set at 8-mm detector pixel size, 80 kV x-ray tube voltage, 80 mA x-ray tube current, 1
1 detector bin mode, and 500 view angles. Images are acquired in the axial mode, covering a volume of 2.0 cm along the z-axis with 1.04-cm field of view (covering a single hindlimb). Micro-CT is calibrated using standard wires of different sizes (10, 20, 30, 40, and 50 mm). 6. Reconstruction is carried out as either 500 slices at 8 mm or 250 slices at 16 mm with a cone-beam algorithm. Bone and pre-existing vessels are used as anatomical references. A 2D maximum intensity projection (MIP) image is generated using Microview software (GE Healthcare). The desired threshold is then chosen using a modified Image Pro-Plus 5.0 algorithm Vessel Tree 0.9. The data are expressed as a number vascular segments per slice and then integrated over the entire 500-slice set.

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Figure 14.2 Micro-CT analysis of mouse hindlimb circulation. Two-dimensional micro-CT images of a normal hindlimb (Panel A) and an ischemic hindlimb 4 weeks after femoral artery ligation (Panel B) at 16 mm reconstruction. A high-resolution, twodimensional reconstruction at 8 mm resolution is shown in z-plane (panel C) and x-plane (panel D). Newly formed collateral arteries are indicated by arrows.

2.1.2. Quantitative image analysis Image-Pro Plus 5.0.1 (Media Cybernetics, Silver Spring, MD) is used for image analysis and quantification of the number, length, and volume of vessels. The algorithm was built within the AutoPro programming language (Visual Basic SAX engine) inside Image-Pro Plus. The 3D imaging data are analyzed on each of the 2D planes that make up the 3D volume. On each plane, the ‘‘spots’’ of vasculature are counted by width and area. The total number of spots at each width across all planes corresponds to the vessel length at that width. The total area at each width corresponds to the vascular volume for that vessel size. The minimum width is used in the 3D analysis.

2.2. Micro-CT coronary angiography The goal of this technique is to visualize coronary arteries. The key to success is filling the epicardial coronaries without filling the left ventricular cavity with contrast, which interferes with epicardial vessel visualization. For this reason, bismuth contrast is used.

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Figure 14.3 Micro-CTof renal circulation, 2D representation of a normal mouse renal arterial vasculature at 16 mm resolution.

2.2.1. Bismuth contrast agent preparation 1. Add 100 g bismuth oxychloride to 800 ml 18% NaCl, and mix thoroughly with stir bar (the mixture will remain turbid). 2. While stirring, add concentrated HCl until solution becomes clear (final volume will be around 1000 ml). 3. Filter through gauze to remove large particles, which can block smaller vessels. 4. Fill one or more buckets with a total of 20 l cold tap water. 5. Add the bismuth solution slowly (with a large pipette) to the tap water while vigorously stirring the solution. 6. Allow bismuth to precipitate (30 to 60 min). 7. Remove the supernatant. 8. Pool the bismuth precipitate in a wide-bore 1-l cylinder and allow to sediment further overnight. 9. Remove supernatant and store at 4  C (more supernatant can be removed over time; the bismuth contrast agent should be stable for months) and store in 20 ml aliquots at 4  C). 10. Immediately prior to performing angiography, mix bismuth 1:3 with 10% gelatin in PBS (25% bismuth and 7.5% gelatin). Keep the contrast agent at around 37 to 40  C, constantly stirring and covered to prevent evaporation. 2.2.2. Complete filling of the coronary arterial vasculature 1. Anesthetize and ventilate the mouse as described previously. 2. Open the chest, keeping both internal mammary arteries intact.

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3. Insert a 21G butterfly needle into the thoracic descending aorta; ligate all major branches arising from the aortic arch using 5-0 silk suture. Cut the inferior vena cava to allow venous drainage. 4. The heart is then arrested in diastole by a retrograde perfusion of oxygenated St Thomas cardioplegic solution at 25  C. 5. Infuse vasodilation buffer (papaverin 4 mg/l, adenosine 1 g/l in PBS) for 3 min at 120 mmHg pressure. 6. Infuse 2% paraformaldehyde (in PBS) for 5 min. 7. Flush with PBS for 2 min. 8. The bismuth contrast agent solution is then administered by retrograde injection of 0.1 ml/10 g heart weight at 0.25 ml/min using an automatic mechanical injector (mechanical injector). This step is tricky and the volume of the contrast agent is dependent on animal size, species, tip location, and the length of the catheter. Ideally, volume and injection rate should be controlled and kept constant from animal to animal in the same study. 9. Once the distal coronary arteries have been filled, quickly cover mouse heart with ice water for 5 min to allow the contrast agent to solidify in situ. 10. Finally, the heart is removed and immersed in 2% paraformaldehyde (4  C) overnight. 11. The fixed heart is then scanned in a micro-CT scanner as described previously at 6.5 to 16 micron resolution. The acquired image is then displayed as a 3D or a series of 2D projections.

3. Microangiography X-ray microangiography of mouse circulation provides a gross overview of arterial circulation, that, while lacking the details of micro-CT images, allows rapid assessment of gross differences between experimental groups. The technique is relatively straightforward, but is not quantitative and lacks the ability to detect arteries less than 20 mm in diameter. The alternative to x-ray angiography is fluorescent angiography (Fig. 14.4) that is particularly effective for visualization of coronary circulation (Chittenden et al., 2006).

3.1. X-ray microangiography: Animal preparation and image acquisition (mice) 1. For the x-ray microangiography, a 1:1 mixture of bismuth (SigmaAldrich) and gelatin (ICN Biomedicals) is used as the contrast agent.

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Figure 14.4 Mouse coronary fluorescent angiography. A microscopic image of a normal-mouse right coronary artery after injection of a fluorescent dye. Bar,0.5 mm.

2. Mice are anesthetized and a 21G butterfly needle is inserted into the thoracic aorta just above the renal arteries, the inferior vena cava is cut to allow outflow of blood and perfusate. 3. Saline containing heparin (3000 IU/l), papaverine (4 mg/l), and adenosine (1 g/l) is infused for 3 min at 100 mmHg pressure immediately prior to contrast perfusion. 4. The contrast agent (0.02 ml/g body) is infused via a syringe pump over 2 min at 100 mmHg pressure, and perfusion fixation is carried out using 2% paraformaldehyde for 5 min at the same pressure. 5. For image acquisition, animals are placed in posteroanterior view and the lower extremities are then exposed on Kodak MIN-R mammography film for 3 min at 25 kVp and 3.25 mA.

3.2. Coronary fluorescent microangiography: Animal preparation and image acquisition (mice) 1. Mice are anesthetized with the ketamine hydrochloride/xylazine mixture and the right carotid artery is then exposed. 2. One milliliter of 10 mg/ml FITC-dextran (Sigma-Aldrich) is infused into the carotid artery at 100 mmHg for 1 min.

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3. The FITC-dextran is allowed to circulate for 4 min before whole hearts are harvested and placed on ice. Images are then acquired using the M2Bio 3D fluorescent microscopy system with Axiovision 4.2 software (Zeiss).

4. Functional Assessment of Arteriogenesis Since the principal result of effective arteriogenesis is restoration of arterial circulation and tissue perfusion, techniques that assess tissue perfusion should provide an accurate assessment of arteriogenic response. Such assessment of perfusion can rely on measurement of tissue flow (laser Doppler imaging), tissue oxygen levels (EPR oximetry), contrast arrival time (T2-weighted magnetic resonance imaging [MRI]) or detection of arterial flow (time-of-flight MRI). Each of these techniques has its own advantages and challenges, and a combination of these approaches provides a more reliable assessment of perfusion than any one given technique.

5. Laser-Doppler Perfusion Imaging The basis of this technique is the user of a Doppler effect to detect blood flow in tissues. The laser light is used to illuminate the arterial blood flow in subcutaneous tissues and the magnitude of the Doppler shift reflects the velocity of the flow. The key parameters are the extent of laser light penetration and the angle of alignment to the direction of the flow. Because most of the flow examined is subcutaneous (due to a limited depth of penetration), the results are very sensitive to any changes in vasomotion due to, for example, changes in temperature or the type of anesthesia used. Despite these caveats, with proper controls the technique provides a reasonably reliable assessment of perfusion in mice feet.

5.1. Animal preparation and data acquisition 1. Mice are anesthetized with a ketamine/xylazine (80/5 mg/kg) mixture and are positioned ventrally on a heating pad set to 38  C sufficient to keep skin temperature at 36.5 to 37.5  C, and in a box to prevent air drafts that can constrict superficial blood vessels. 2. The rear legs are outstretched to allow scanning of the full plantar surface. 3. Images are acquired using a Moor Infrared Laser Doppler Imager (Moor Instruments Ltd.).

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4. The data are analyzed with Moor LDI image processing software V3.09, and reported as ratio flow right/left (R/L) hindlimb after background subtraction.

6. Electron Paramagnetic Resonance Tissue Oxymetry Electron paramagnetic resonance (EPR) technique allows assessment of tissue concentrations of oxygen and thus can be used to assess effects of changes in arterial perfusion on tissue. The basis of EPR is the paramagnetic nature of molecular oxygen, which alters the EPR spectra of other paramagnetic species in its vicinity. The magnitudes of the effects are directly related to the amount of oxygen that is present in the environment of the paramagnetic materials (Dunn and Swartz, 2003). Thus, with a pellet made of a paramagnetic material such as charcoal inserted in a tissue, EPR spectrum will reflect molecular oxygen levels in the vicinity of this pellet. Under appropriate conditions, EPR is accurate over a wide spectrum from less than 0.1% to nearly 100% oxygen levels (Khan et al., 2007). EPR is not an imaging technique and it provides no spatial information regarding distribution of the signal. However, positions of paramagnetic pellets around which the EPR signal is generated can serve as a surrogate for spatial distribution of the signal. Furthermore, once implanted, such particles, which are biologically inert, remain at the site of implantation for a very long time, thereby allowing serial studies. The EPR signal is measured with a detector placed on the surface of the skin in an appropriate combination of electromagnetic and magnetic fields. In practice, this is achieved using dedicated EPR magnets.

6.1. Electron paramagnetic resonance data acquisition 1. Mice are anesthetized with ketamine and xylasine and gastrocnemius, adductor, or any other desired muscle is exposed. 2. Under direct vision, 100 mg of activated char is injected via a 26G needle into the muscle in one or more locations as desired. 3. The incision is closed and the animal is allowed to recover for 2 weeks. 4. Tissue pO2 levels are then determined at baseline with an anesthetized animals placed in an EPR magnet. 5. Following common femoral artery ligation, pO2 in the ischemic (gastrocnemius) and nonischemic (adductor) muscle are then measured over time.

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7. Magnetic Resonance Imaging MRI provides a comprehensive set of tools for detection of arterial conduits and assessment of tissue perfusion. The principal use of MRI discussed here is the time-of-flight (TOF) imaging that can provide quantitative assessment of arterial blood flow, although other, more complex MRI techniques such as T2-weighted perfusion imaging can be used as well. The principle of TOF imaging is the ability to saturate a single image slice with a signal that is proportional to the velocity of blood flow using large flip angles and fast repetition times of the magnetic field (Wagner et al., 2004a). The technique is effective in mice and generates very reliable assessment of blood flow (Wagner et al., 2004b).

7.1. Time-of-flight magnetic resonance imaging data acquisition 1. Mice are anesthetized with ketamine and xylazine, and the legs and lower body are immobilized and secured to a holding pad that can fit inside the imaging coil. 2. The temperature of the pad and the coil is maintained at 38  C. 3. Images are then acquired using an inductively coupled low-pass, 2.5-cm long quadrature birdcage coil designed for use in mice in a 7T horizontal-bore magnet with a Varian Unity console. 4. A gradient echo MR sequence is used with TR/TE/a ¼ 0.03/0.0035/ 90 degrees, slice thickness 600 mm, 3
3-cm field of view, 512
128 matrix size, resulting in a pixel resolution of 59
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600 mm. 5. For processing, a region of interest is selected in an area of the nonischemic leg that does not include any visible vessels. The image is then windowed to include only pixels with the signal intensity that is above this region’s signal mean plus six standard deviations. The number of bright spots (conductance vessels) is counted in the ischemic leg for both groups at different time points. Taken together, the various anatomical and functional tools for assessment of arteriogenesis discussed here provide a comprehensive and complete picture of the state of arterial circulation in mice vascular models.

ACKNOWLEDGMENTS Many individuals have contributed to development of the Dartmouth arteriogenesis imaging program. Specifically, I’d like to thank Zhenwu Zhuang and Lyubomir Zagorchev (microCT); Harold Swartz, Nadeem Khan, and Armin Helisch (EPR); and Jeffrey Dunn and Justin Pearlman (MRI). Finally, I would like to thank members of my laboratory and the Dartmouth Angiogenesis Research Center for developing and testing many techniques described here.

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REFERENCES Carmeliet, P. (2005). Angiogenesis in life, disease and medicine. Nature 438, 932–936. Chittenden, T. W., Claes, F., Lanahan, A. A., Autiero, M., Palac, R. T., Tkachenko, E. V., Elfenbein, A., Ruiz de Almodovar, C., Dedkov, E., Tomanek, R., Li, W., Westmore, M., et al. (2006). Selective regulation of arterial branching morphogenesis by synectin. Dev. Cell. 10, 783–795. Dunn, J. F., and Swartz, H. M. (2003). In vivo electron paramagnetic resonance oximetry with particulate materials. Methods 30, 159–166. Helisch, A., Wagner, S., Khan, N., Drinane, M., Wolfram, S., Heil, M., Ziegelhoeffer, T., Brandt, U., Pearlman, J. D., Swartz, H. M., and Schaper, W. (2006). Impact of mouse strain differences in innate hindlimb collateral vasculature. Arterioscler. Thromb. Vasc. Biol. 26, 520–526. Khan, N., Williams, B. B., Hou, H., Li, H., and Swartz, H. M. (2007). Repetitive tissue pO2 measurements by electron paramagnetic resonance oximetry: Current status and future potential for experimental and clinical studies. Antioxid. Redox Signal. 9, 1169–1182. Li, W., Shen, W., Gill, R., Corbly, A., Jones, B., Belagaje, R., Zhang, Y., Tang, S., Chen, Y., Zhai, Y., Wang, G., Wagle, A., Hui, K., et al. (2006). High-resolution quantitative computed tomography demonstrating selective enhancement of mediumsize collaterals by placental growth factor-1 in the mouse ischemic hindlimb. Circulation 113, 2445–2453. Simons, M. (2005). Angiogenesis: Where do we stand now? Circulation 111, 1556–1566. Tirziu, D., and Simons, M. (2005). Angiogenesis in the human heart: Gene and cell therapy. Angiogenesis 8, 241–251. Wagner, S., Helisch, A., Bachmann, G., and Schaper, W. (2004a). Time-of-flight quantitative measurements of blood flow in mouse hindlimbs. J. Magn. Reson. Imaging 19, 468–474. Wagner, S., Helisch, A., Ziegelhoeffer, T., Bachmann, G., and Schaper, W. (2004b). Magnetic resonance angiography of collateral vessels in a murine femoral artery ligation model. NMR Biomed. 17, 21–27. Zhuang, Z. W., Gao, L., Murakami, M., Pearlman, J. D., Sackett, T. J., Simons, M., and de Muinck, E. D. (2006). Arteriogenesis: Noninvasive quantification with multi-detector row CT angiography and three-dimensional volume rendering in rodents. Radiology 240, 698–707.

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Methods to Study Myeloid Cell Roles in Angiogenesis Michael C. Schmid and Judith A. Varner Contents 1. Introduction 1.1. Angiogenesis 1.2. Myeloid cells: Monocytes and macrophages 1.3. Recruitment of monocytes into tumors 1.4. Integrin roles in myeloid cell trafficking to neovascular sites 1.5. Inflammation and cancer 2. Methods for the Study of Myeloid Cells in Tumor Neovascularization and Growth 2.1. Identification of myeloid cells in tissues by immunohistochemistry 2.2. Isolation of human and murine peripheral blood mononuclear cells 2.3. Isolation of murine mononuclear cells from bone marrow 2.4. Integrin activation as measured by clustering 2.5. Adhesion of myeloid cells to purified ligands 2.6. Adhesion of myeloid cells to endothelial cells in vitro 2.7. In vitro myeloid migration/invasion assays 2.8. In vivo monocyte-trafficking studies 2.9. Matrigel angiogenesis assays 2.10. Role of myeloid cells in tumor growth studies 2.11. Quantification of myeloid cells in experimental tumors by FACS, qPCR, and IHC 3. Conclusions References

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Abstract Tumor growth and metastasis depend on neovascularization, the growth of new blood vessels. Recent studies have found that bone marrow derived cells contribute to angiogenesis during tumor growth and inflammation. Tumor

Moores UCSD Cancer Center, University of California-San Diego, La Jolla, California Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03015-2

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neovascularization is regulated in part by monocytes, which are myeloid lineage cells from the bone marrow. Tumors exhibit significant monocyte infiltrates, and recent studies indicate that monocytes are actively recruited to the tumor microenvironment. Upon tumor infiltration, monocytes can participate in tumor neovascularization by differentiating into M2 macrophages, which express proangiogenic growth factors. By understanding how bone marrow– derived cells contribute to tumor growth, it may be possible to develop new approaches to cancer therapy. In this chapter, we discuss experimental methods to examine the roles of myeloid cells in tumor growth and angiogenesis, including methods to identify, isolate, purify, and characterize bone marrow– derived monocytes. We also outline methods to analyze the in vivo roles of myeloid cells in tumor growth and angiogenesis using adoptive transfer, bone marrow transplantation, tumor models and immunohistochemistry for markers of vessels and myeloid cells. Finally, we review methods to characterize myeloid cell trafficking in vitro and in vivo.

1. Introduction 1.1. Angiogenesis Neovascularization, the formation of blood vessels, plays important roles in development, inflammation, and wound repair. All cells require oxygen and nutrients for survival and are found in close proximity to blood vessels. During conditions of hypoxia or tissue injury, the formation of new blood vessels, or angiogenesis, is induced. During angiogenesis, new blood vessels originate from preexisting vessels by sprouting of new blood vessels from pre-existing vessels (Carmeliet, 2005; Folkman et al., 1971). Proangiogenic growth factors, including vascular endothelial growth factor (VEGF) and basic fibroblast growth factor (bFGF), can induce quiescent endothelial cells to undergo proliferation and migration, resulting in the formation of new vessel structures (Carmeliet, 2005; Coultas et al., 2005). Vasculogenesis, the coalescence of new blood vessels from individual endothelial cells or progenitor cells, also occurs during tumor neovascularization (Asahara et al., 1997; Lyden et al., 2001). Importantly, tumor neovasculogenesis is modulated by monocytes (reviewed by Pollard et al., 2004).

1.2. Myeloid cells: Monocytes and macrophages Monocytes and macrophages belong to the myeloid cell lineage and derive from myeloid progenitor cells. Monocytes arise in the bone marrow; upon maturation, they are released into the bloodstream. Circulating monocytes migrate into tissues where they undergo transformation into macrophages. The number of macrophages in tumor tissues is significantly greater than in

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normal tissues ( Jin et al., 2006; Lin et al., 2006, 2007). Importantly, this increase in macrophage content in tumor tissue appears to result from recruitment of circulating monocytes, rather than the expansion of resident macrophages. Tumor-associated macrophages (TAMs) have been shown to promote neovascularization by two different mechanisms. TAMs release a number of potent proangiogenic cytokines and growth factors, such as VEGF, tumor necrosis factor-a (TNF-a), interleukin-8 (IL-8), and bFGF (Lewis et al., 2000; Yamashiro et al., 1994; Sica and Bronte, 2007). Additionally, TAMs also express a broad array of proteases known to play roles in the angiogenic process. These proteases include urokinase-type plasminogen activator (uPA) and the matrix metalloproteinases MMP-2, MMP-7, MMP-9, and MMP12 (Sica and Bronte, 2007; Jodele et al., 2005, Giraudo et al., 2004). uPA and MMPs support angiogenesis by remodeling and breaking down the extracellular matrix (ECM). Degradation of ECM leads to the mobilization of growth factors and facilitates the migration of vascular cells into new environments. The proangiogenic functions of TAMs may account for the correlations observed between TAM densities and vascular densities in many tumor types. Importantly, high TAM densities indicate poor prognoses in breast, prostate, ovarian, and cervical cancer (Bolat et al., 2006; Esposito et al., 2004; Leek et al., 2002; Nishie et al., 2000; Sutsui et al., 2005; Valkovic et al., 2002).

1.3. Recruitment of monocytes into tumors Increasing evidence indicates that myeloid cells promote neovascularization. Immune cell trafficking in vivo is regulated by cytokines and by members of the integrin, immunoglobulin, and selectin adhesion molecule families (Weber and Koenen, 2006; Luster et al., 2005). Several chemokines and their receptors have been shown to recruit circulating monocytes. Monocyte chemoattractant protein-1 (MCP-1, or CCL2) and RANTES (CCL5) increase the infiltration of TAMs in several primary tumors including breast, ovarian, melanoma, and glioblastoma (Lin et al., 2001; Murdoch et al., 2004; Niwa et al., 2001; Ueno et al., 2000). Furthermore, CCL2 and CCL5 stimulate the secretion of matrix-degrading enzymes, such as MMP9 and MMP12, in macrophages. Another chemokine, interleukin-8/CXCL8 (IL-8), a proangiogenic chemokine, was identified as an autocrine growth factor in several human cancers (Zhu et al., 2004). IL-8 stimulates the adhesion of monocytes, which express low levels of the IL-8 receptors, CXCR1 and CXCR2, to vascular endothelium under flow conditions, indicating that IL-8 and CXCR-1 and -2 play roles in monocyte recruitment (Gerszetsen et al., 1999). Several growth factors and cytokines such as colony stimulating factor-1 (CSF-1), VEGF, and platelet-derived growth factor-D (PDGF-D) have also been implicated in the recruitment of monocytes into tumors (Uutela et al., 2004; Barleon et al., 1996). CSF-1 is produced by various types of human

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tumors, and is a potent chemoattractant for macrophages. Coordinated expression of CSF-1 in macrophages together with epidermal growth factor (EGF) in primary mammary tumor cells increase invasion into mammary tumors (Goswami et al., 2005). IL-1beta increases infiltration of neutrophils and macrophages as well as angiogenesis in mouse models of corneal neovascularization. Deletion of monocytes using genetic approaches or toxins suppresses IL-1beta–induced angiogenesis (Nakao et al., 2005). Importantly, CD11bþ F4/80þ cells that infiltrated sites of corneal neovascularization were COX-2þ, while macrophages that remained in the limbal vessel did not express COX-2. As signaling molecules downstream of COX-2, such as PGE2 and thromboxane A2, have been shown to enhance the production of various angiogenesis-related factors like VEGF, MMPs, and chemokines, these studies suggest that COX-2 expressing macrophages help to regulate angiogenesis (Nakao et al., 2005).

1.4. Integrin roles in myeloid cell trafficking to neovascular sites Our laboratory recently described the mechanisms by which myeloid cells adhere to tumor endothelium and extravasate into ischemic or tumor tissue. Myeloid cells express a number of functional integrins (a2b1, a4b1, a5b1, avb3, avb5, amb2 (CD11b) and aXb2 (CD11c) that could play roles in the trafficking of these cells to and within neovascular microenvironments. Our studies on the roles of integrins in circulating bone marrow–derived cell trafficking revealed important functions for integrin a4b1 ( Jin et al., 2006a,b). We found that integrin a4b1, a receptor for vascular cell adhesion molecule (VCAM) and fibronectin, selectively promotes the homing of monocytes ( Jin et al., 2006a,b) to neovascular tissue, and that this integrin is essential for the participation of these cells in angiogenesis and tumor growth. Human and murine bone marrow–derived myeloid cells (CD14þ CD11bþ) adhered to endothelial cells in vitro and tumor endothelium in vivo via integrin a4b1. Treatment of mice bearing Lewis lung carcinoma (LLC) tumors with antagonists of integrin a4b1 significantly suppressed the number of monocytes/endothelial progenitor cells within tumors and reduced blood vessel density. Our studies suggested that suppression of monocyte/macrophage homing to tumors by the application of an integrin a4b1 antagonist could be a useful supplementary approach to suppress tumor angiogenesis and growth.

1.5. Inflammation and cancer Links between chronic inflammation and cancer have been recognized for several decades (Coussens and Werb et al., 2002). Tumor-associated macrophages (TAMs) have been characterized, and studies to understand their

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recruitment and contributions to angiogenesis are ongoing. However, new studies suggest that several myeloid subpopulations play distinct roles during neovascularization such as in tumor development. These cells may form vessels, secrete growth factors, or support angiogenesis in other ways. These properties define myeloid cells/progenitor cells as putative targets for anticancer therapies. Suppression of cell homing to the tumor microenvironment offers a new strategy to inhibit tumor neovascularization, while stimulation of homing may promote tissue recovery from ischemia.

2. Methods for the Study of Myeloid Cells in Tumor Neovascularization and Growth To study the roles of myeloid cells in tumor neovascularization, we purify CD11bþ myeloid cells from human or murine peripheral blood or murine bone marrow. We then investigate the adhesive properties and receptor expression profiles of CD11bþ cells using the methods provided below. We also investigate the mechanisms by which monocytes traffic to tumor tissue, promote angiogenesis, and modulate tumor growth using the methods described below.

2.1. Identification of myeloid cells in tissues by immunohistochemistry To identify myeloid cells in tissues, we perform immunohistochemistry to detect myeloid cells using markers of monocytes (CD11b, CD14) and differentiated macrophages (F4/80). We typically cut 5-mm sections of cryo-preserved tissues and mount two sections per microscope slide, and then store the slides at –80  C. To immunostain, remove frozen sections on glass microscope slides from –80  C freezer and allow to warm to room temperature (RT). Fix slides for 2 min in cold acetone, air dry, and then draw a box around the section with a hydrophobic pen (PAP-Pen). Alternatively, encircle the section with a Pap-Pen prior to fixation in 3.7% paraformaldehyde for 2 to 5 min. Wash slides two times for 5 min each in PBS. Never allow sections to dry out. Create a humidified chamber by placing a damp paper towel in the bottom of a plastic box with a sealing lid and then place two glass pipettes in parallel on top of the paper towel. Place the slide flat on pipettes and add 100 ml of blocking buffer comprised of 8% normal human or goat serum in PBS on each encircled section. Incubate 2 h at RT or overnight at 4  C. Use of human serum helps to block Fc-receptors. Rinse the slides twice for 5 min each in PBS with shaking or agitation

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in PBS. Dry the glass portions of the slide, and remove excess liquid from the section by touching the corner of a Kimwipe to the edge of the section. Apply 100 ml of primary antibody diluted typically at 1 to 5 mg/ml in 5% normal goat serum or 2.5% bovine serum albumin (BSA) in PBS to the encircled section on the left of the slide. Apply only block buffer to the second section on the slide, which serves as a negative control section. Incubate for 2 h at RT. Wash the slides five times for 5 min in PBS with agitation. Dry the slide and wick the excess liquid from the section as described previously. Incubate in secondary antibody (such as highly cross-absorbed Alexa 488-conjugated goat anti-rat IgG) diluted 1/250 to 1/1000 in 5% NGS or 2.5% BSA in PBS, by applying 100 ml to the encircled section. Incubate 1 h at RT. If the primary antibody is directly labeled, skip this step. Wash the slide five times for 5 min in PBS. Dry the slide and wick the excess liquid from the section. Incubate in nuclear stain such as DAPI or TOPRO for 10 to 30 min. Wash once in PBS and apply mounting media containing anti-fade. Place a coverslip gently on the section. Place a drop of nail polish on the corners of the coverslip to fix it in place. Store slides in refrigerator in a dark, flat slide holder or in a slide box. To immunostain for the myeloid cell marker CD11b, we use rat antimouse/human monoclonal antibody M1/70 (BD Pharmingen) and to immunostain for the murine macrophage antigen F4/80, we use rat antimurine F4/80 clone B8M8 (eBioscience, San Diego, CA). To stain for human CD14, we use mouse anti-human CD14 clone 61D3 (eBioscience, San Diego, CA). Blood vessel density is determined after immunostaining for von Willebrand factor with antibody A0082 from Dako or for CD31 (rat anti-mouse CD31, clone Mec 13.3 from BD Pharmingen or mouse antihuman CD31, clone P2B1 from Millipore). We quantify CD11bþ or F4/ 80þ cells per high-power field by determining pixel density/200 microscopic field using MetaMorph (Version 6.3r5, Molecular Devices) to determine the degree to which myeloid cells invade normal and diseased tissues, such as tumors. At least five randomly selected microscopic fields per section are quantified. We also quantified VWF or CD31þ blood vessels/200 field (Fig. 15.1).

2.2. Isolation of human and murine peripheral blood mononuclear cells To isolate myeloid cells from peripheral blood, we first purify total mononuclear blood cells from peripheral blood. Centrifuge freshly isolated blood over a density gradient to purify peripheral blood mononuclear cells (PBMCs). Aliquot 15-ml RT Histopaque 1077 (Sigma Aldrich, St. Louis, MO) into 50-ml tubes (or 5-ml Histopaque 1083 in 15-ml tubes for rodent cells) for density gradient centrifugation. Dilute freshly isolated blood in two to four volumes of Ca2þMg2þ-free PBS containing 0.5 mM EDTA (PBS-E). Alternatively, dilute buffy coat at the same proportions in PBS-E.

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Figure 15.1 Mouse and human breast carcinomas are highly infiltrated by monocytes. Upper panel: Immunohistochemical detection of blood vessels by expression of vWF (green, white arrowhead) and myeloid cells by expression of CD11b (red, yellow arrowhead) in normal mouse breast and spontaneous breast carcinoma. Lower panel: Immunohistochemical detection of blood vessels and monocytes, respectively, by expression of vWF (green) and CD14 (red) in normal human breast and human breast carcinoma.The intensity of each staining was quantified as number of pixels per field and graphed.

Then carefully layer 35 ml diluted blood suspension on top of the Histopaque layer. Centrifuge immediately at 400 g for 25 min at RT without braking. After centrifugation, carefully aspirate the upper half of the gradient and discard. The opaque interface contains mononuclear cells (MNCs). Carefully remove the mononuclear cell layer with a 10-cc syringe and 18G needle. Transfer these cells to centrifuge tubes containing 10 ml PBS-E for collection of the cells, and centrifuged at 300 g for 10 min at 20  C to pellet cells and to remove Histopaque. It is important to note that centrifugation at low temperatures may result in cell clumping and lower recovery. Aspirate the supernatant and resuspend cells in a small amount of PBS-E (5 to 10 ml), pooling all cells into a single 50-ml tube. Adjust the volume to 50 ml with PBS-E, gently mix the cells with a pipette. Centrifuge again at 200 g for 10 min at 20  C without braking to remove platelets. Aspirate the supernatant and resuspend pellets again, pooling all cells together in total volume of 5 ml of PBS-E. Add 15 to 20 ml of ammonium-chloride, red blood cell (RBC) lysis solution (155 mM NH4Cl, 10 mM NaHCO3, and 0.1 mM EDTA), and incubate cells on ice for 5 to 10 min. In the case of large amounts of contaminating RBCs, add 20 ml ammonium chloride before incubating for 10 min on ice. Use a ratio of 4:1 ammonium chloride to PBSE in a 15-ml tube for isolation of murine cells. Add PBS-E to a final volume of 50 ml for human cells or 15 ml for murine cells. Centrifuge the cells at 1300 rpm for 5 min at 5  C to remove residual ammonium chloride. Aspirate the supernatant and resuspend in 5 ml PBS-E (or in 1 ml for murine cells), and determine the number of cells/ml using a hemacytometer (Fig. 15.2).

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Figure 15.2 Isolation of peripheral blood mononuclear cells. Human buffy coats were purchased fromthe San Diego Blood Bank and processed immediately. Bloodwas diluted in PBS-E and gently applied to a Histopaque layer. Upon centrifugation, the cell layer containing the mononuclear cells was carefully transferred into a new tube.The remaining blood cells (indicated as stars) were lysed by ammonium chloride treatment.

2.3. Isolation of murine mononuclear cells from bone marrow To purify mononuclear cells from murine bone marrow, aseptically remove femurs and tibias from the legs of euthanized mice. Trim muscle tissue from femurs. Cut off the ends of each bone and flush with PBS, pH 7.2, supplemented with 0.5% BSA and 2 mM EDTA (2 ml/femur and 1 ml/tibia) into plastic centrifuge tubes on ice. Triturate the cell suspension to break up

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clumps and form single cell suspensions. Centrifuge cells at 400 g for 10 min at RT, resuspend in 10 ml ammonium-chloride, red-cell lysis buffer (155 mM NH4Cl, 10 mM NaHCO3, and 0.1 mM EDTA), and incubate for 4 to 5 min at RT. Transfer the cell suspension to a 50-ml centrifuge tube and add 30 ml PBS without MgCl2 or CaCl2. Centrifuge cells at 400 g for 10 min at RT and resuspend pellets in 4 ml with PBS, pH 7.2, supplemented with 0.5% BSA. Transfer resuspended pellets to 15-ml Falcon tubes, add 4 ml of RT Histopaque 1083 to the bottom of the tube and centrifuge tubes at 400 g for 25 min at RT without braking. Aspirate the surface layer and then transfer the cloudy layer containing the purified bone marrow cells into new 15-ml centrifuge tubes. Wash pellets twice with PBS by resuspending in PBS and centrifuging at 400 g for 10 min at 4  C. Finally resuspend cells at 2.5  107 cells/ml in PBS. Using this method, approximately 5  107 bone marrow–derived mononuclear cells can be purified by gradient centrifugation from the femurs and tibias of a single mouse (Fig. 15.3). 2.3.1. Purification of CD11bþ cells As myeloid cells are CD11b positive, we use anti-CD11b antibody– mediated, magnetic bead–affinity chromatography (130-049-601, Miltenyi Biotec) to purify myeloid cells from human or mouse total peripheral blood mononuclear cells. Incubate single-cell suspensions of PBMCs in PBS with Fc-receptor-blocking reagent (human, 130-059-901, and mouse, 130-092575) (Miltenyi Biotec) at 2 ml per 106 cells on ice for 5 min to block nonspecific binding of primary antibody. Add anti-CD11b–coated magnetic beads (10 ml/107 cells) and incubate for 15 min on ice. To quantify the purity of the CD11bþ cell population, add fluorescently labeled anti-CD11b antibody together with the magnetic beads (1 ml/106 cells). Wash cells with Miltenyi Affinity chromatography buffer (MAC buffer) (PBS, pH 7.2, supplemented with 0.5% BSA and 2 mM EDTA), and centrifuge at 1200 rpm for 10 min. Resuspended cells and pass them through a 30-mm mesh or over a wetted prefilter and apply the final suspension to prewetted magnetic separation columns (MS columns, 130-042-201, Miltenyi Biotec) mounted on a magnetic plate. Wash columns three times in MAC buffer and remove from the magnet. Carefully elute CD11bþ cells in 1 ml MAC buffer. An aliquot of the column flow-through and of the column eluate should be evaluated by flow cytometry. The enriched fraction in the eluate should be over 99% CD11b-positive. To ensure a pure population, it is best to repeat the magnetic separation twice by applying the eluted cells to a new prefilled, positiveselection column, wash, and elute retained cells in 2.5 ml buffer (Fig. 15.4A). 2.3.2. Fluorescence-activated cell sorting analysis To evaluate the expression of myeloid cell markers and integrins on purified myeloid cells, aliquot 5  105 cells into polypropylene snap-cap tubes, one tube per antibody to be tested. One tube for a positive control and one tube

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Figure 15.3 Isolation of bone marrow^derived cells. Mice were euthanized and bones (femur/tibias) were isolated.The ends of the bones were carefully cut off, and the bone marrow was flushed out using an 18G syringe. Cells were first treated with ammonium chloride to remove red blood cells, and then mononuclear cells were separated by gradient centrifugation.

for a negative control should also be prepared. Centrifuge cells at 200 g and remove supernatants. Resuspend cells in cold PBS containing 1% BSA (RIA grade). Incubate cells in FcR blocking reagent (2 ml/106 cells) for 30 min on ice and wash in PBS by centrifugation. Add 100 ml of primary antibody per tube (e.g., 1 to 10 mg antibody, prepared in tissue culture medium containing 1% BSA) and incubate cells 20 to 60 min in the dark on ice. Wash cells twice by centrifugation with cold PBS containing 1% BSA. For antibodies that are not directly labeled, we then add 100 ml of

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fluorochrome-conjugated, highly cross-absorbed secondary antibody per tube of cells (e.g., 1 to 10 mg/ml in cold tissue culture medium) and incubate for 20 to 60 min on ice in the dark. Wash cells twice by centrifugation with cold PBS containing 1% BSA. Fix cells with 200 ml of cold 0.5% paraformaldehyde in PBS and store in the dark at 4  C until analysis. To evaluate myeloid cells markers, we use anti-human/mouse CD11b (130091-241, Miltenyi Biotec) (Fig. 15.4B). To evaluate integrin expression on human myeloid cells, we use PEconjugated mouse anti-human a4b1 (clone 9F10, BD Biosciences, San Jose, CA), PE-conjugated anti-human a4b1 (clone IIA1, BD Biosciences, San Jose, CA), PE-conjugated anti-human a2 (clone CLB-LFA 1/1, eBioscience, San Diego, CA), mouse anti-human avb3 (FITC-conjugated LM609, Millipore), mouse anti-human avb5 (PE-conjugated P1F6, Millipore), mouse anti-human b1 (clone P4C10, Millipore), and mouse antihuman b7 (clone FIB504, BD Biosciences, San Jose, CA). To evaluate integrin expression on murine myeloid cells, we use PE-conjugated antimouse a4b1 (R1-2, BD Biosciences), PE-conjugated rat anti-mouse a5b1 (5H10-27, BD Biosciences, San Jose, CA), PE-conjugated rat anti-mouse av (RMV-7, BD Biosciences), PE-conjugated rat anti-mouse b2 (clone MK 18/2, BD Biosciences), and anti-CD11b (M1/70, BD Biosciences) ( Jin et al., 2006a,b). 2.3.3. Characterization of integrin activation on myeloid cells Studies from our lab indicate that that integrin a4b1 plays a critical role during monocyte trafficking to tumors. Integrin a4b1 is expressed in an inactive state on circulating blood cells, including monocytes. Monocyte integrin a4b1 can be activated by chemokines, which results in a conformational change and clustering in the plane of the membrane (Hynes, 2002). To evaluate the effect of various chemokines on integrin activation and function, we expose monocytes to chemokines such as stromal-derived factor-1 (SDF-1 or CXCL12) and incubate monocytes in the presence of the anti-beta-1 integrin ligand–induced binding site (LIBS)–specific antibody (HUTS21, Becton Dickinson) or an anti-b1 integrin activation– independent antibody (P4C10, Chemicon), as described by Luque et al. (1996) and Carter et al. (1990). To perform this LIBS assays, resuspend mononuclear cells (2.5  106 cells/ml) in EBM-2 containing 5% FBS and 0.5% sodium azide. Block After removal of the magnet, CD11b-positive cells can be eluted from the column with buffer. (B) Total bone marrow^derived cells are labeled prior to affinity purification with anti^CD11b-APC antibody. Fluorescence levels of cells are measured before and after CD11b-affinity purification. Isotype-matched IgG-APC^antibody binding serves as a negative control.

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nonspecific binding sites by incubating cells in 10 mg/ml normal human immunoglobulin (12000C, Caltag) for 45 min on ice. Centrifuge cells at 400 g and resuspend in basal media containing 10 mg/ml normal human IgG, 2.5 mg primary antibody (HUTS21, P4C10, or IgG2 control) and 1 mg/ml CXCL12. As a positive control, incubate cells in 1 mM manganese (Mn2þ), a stimulator of integrin activation, together with 10 mg/ml normal human IgG. Incubate cells 10 min at 37  C. Wash cells twice with EBM2 containing 5% FBS and 0.5% sodium azide. Add 1/400 fluorescently labeled secondary antibody (Alexa 488 goat-anti mouse Fab’2) and incubate for 20 min on ice. Wash samples twice in EBM-2 containing 5% FBS and 0.5% sodium azide and fix them in 0.5% paraformaldehyde in PBS. Quantify integrin beta-1 activation in monocytes by determining fluorescence signal intensity after gating the monocyte population by forward- and sidescatter size determinations (Figs. 15.5 and 15.6).

2.4. Integrin activation as measured by clustering Upon activation, integrins may cluster in the plane of the plasma membrane and thereby increase avidity for ECM ligands. Additionally, integrins may co-localize with growth factor receptors and co-cluster with them. To evaluate the effect of chemokine clustering on integrin activity in CD11b myeloid cells, monocytes were incubated with chemokine-coated microspheres prior to immunolocalization of integrin a4b1. To coat microspheres, wash 9.0-mm diameter beads (Bangs Laboratories) twice in PBS. Add 2 mg/ml CXCL12 (R&D Systems) or 2 mg/ml BSA (RIA grade, Sigma) in a final volume of 50 ml and incubate with rotation overnight at 4  C in the dark. Wash beads twice with PBS and block in 5% BSA at 4  C for 1 h. Wash beads twice and incubate with cells for 5 min at 37  C. Incubate cells for 1 min on ice and centrifuge at 4  C. Fix in 1% PFA. Monocytes

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Immobilize cells on 10 mg/ml poly-L-lysine–coated microscope slides using a Cytospin as described by Wang et al. (2006). To immunostain for integrin localization and clustering, block nonspecific antibody-binding sites by

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Figure 15.7 Integrin-clustering assay. Microsphere beads are coated with chemokines or BSA overnight. Purified CD11b-positive cells are added, and the bead^cell suspension is incubated for 5 min prior to fixation and immunostaining with anti-integrin alpha4 antibodies and counterstaining with a nuclear stain (TOPRO3). Integrin clustering is observed in the presence of SDF-1a, while BSA-coated beads do not induce clustering.

incubation with 5% BSA and FC-Block (Becton Dickinson) for 30 min at RT. Immunostain cells with anti-integrin antibodies and TOPRO-3 (Invitrogen). Analyze integrin clustering by confocal microscopy (Fig. 15.7).

2.5. Adhesion of myeloid cells to purified ligands Monocytes traffic to sites of inflammation and tumor growth, whereupon they contribute to angiogenesis. Our studies have shown that adhesion molecules such as integrin a4b1 mediate adhesion of myeloid cells to vascular endothelium in vitro and in vivo. To study the ability of cytokine stimulated myeloid cells to adhere to purified integrin ligands or other adhesion proteins, coat non-tissue culture–treated 48-well plates overnight at 4  C with integrin ligands or other adhesion proteins, such as recombinant VCAM-1 (R&D Systems). Optimal coating concentrations should be determined for each protein; for VCAM-1, an optimal concentration is 5 mg/ml. The next day, block wells with 5% heat-denatured BSA for 2 h at 37  C. Resuspend murine or human CD11b-positive cells in basal endothelial growth medium (EBM-2, Cambrex) at a concentration of 2  106 cells/ml. Label 3  104 cells with calcein acetoxymethylester (AM) (25 mM, Invitrogen) for 30 min at 37  C. Wash cells twice in EBM to remove excess dye. Stimulate cells by adding cytokines of interest (e.g., TNFa, VEGF, SDF-1, interleukins) or tumor cell–conditioned medium at concentrations from 0 to 200 ng/ml. Plate stimulated cells plated precoated wells at a density of 1  105 cells/well. After 60 min, wash wells three times with prewarmed PBS, and fix in 3.7% paraformaldehyde. Incubate plates for 10 min in the dark at RT and quantify fluorescent-adherent cells using a fluorescence-activated plate reader (Fig. 15.8).

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Figure 15.8 Myeloid cell-ligand adhesion assay. Non^TC-treated, 48-well plates are coated overnight at 4  C with different ligands. The next day, plates are blocked with 5% BSA, and calcein AM labeled human or murine myeloid cells are added (1  105 cells/well).Thirty min later, wells are washed 3 with PBS to remove unbound cells and remaining cells are fixed in paraformaldehyde. Bound fluorescent myeloid cells are quantified by fluorescence-activated ELISA reader.

2.6. Adhesion of myeloid cells to endothelial cells in vitro To evaluate the effect of various chemokines on the ability of myeloid cells to adhere to endothelial cells, we culture endothelial cells as monolayers, fluorescently label purified monocytes, treat them with chemokines or tumor conditioned medium and perform short-term myeloid cell–endothelial cell adhesion assays. To culture endothelial cells for myeloid cell–adhesion assays, prewarm EGM-2 culture medium (EBM-2 containing 2% FCS, human VEGF-A, human FGF-2, human epidermal growth factor, insulin-like growth factor, gentamicin, ascorbic acid, and prednisone (Cambrex, East Rutherford, NJ) in a 37  C water bath. Rapidly thaw one vial of cryopreserved endothelial cells and pipette into warmed medium immediately at a density of 5  103 cells/cm2. Culture cells in 1 ml medium/cm2 until cells are more than 40% confluent, then culture them at 2 ml/cm2. Change culture medium every other day on a regular basis. Split cells 1:4 when they are 60 to

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80% confluent using 0.025% trypsin, 0.01% EDTA (CC-5012, Cambrex, East Rutherford, NJ). Note that this is 10 lower than usual concentrations. Add trypsin-neutralization solution (CC-5002, Cambrex, East Rutherford, NJ) and plate cells. Do not allow cells to become confluent, as they are sensitive to contact inhibition. Freeze several vials after the first passage. Cells are useful for up to six passages before they begin to senesce. To prepare HUVEC monolayers, seed 200,000 HUVECs in EGM-2 per well of a 48-well plate 24 h before adhesion assay. Do not allow cells to become confluent, as they are sensitive to contact inhibition and also may lift off the well as a sheet of cells if too confluent. To prepare monocytes for adhesion assays, pellet purified cells by centrifugation and resuspend in warmed adhesion medium containing 25 mM CMTMR (C2927, CellTrackerTM Orange CMTMR (5-(and-6)-(((4-chloromethyl)benzoyl) amino)tetramethylrhodamine)–mixed isomers, Invitrogen, Carlsbad, CA). Incubate at 37  C for 60 min. Wash the cells twice by centrifugation and resuspension in prewarmed adhesion medium. Rinse and resuspend cells at 800,000 cells/ml in adhesion buffer. Remove supernatant from HUVEC monolayers, using only a Pipetman (Gilson, Inc.) to aspirate. Add 200,000 EPCs or other bone marrow–derived cells in 0.25 ml per well of HUVEC monolayer. Incubate in 37  C incubator 30 min to 2 h. Aspirate supernatant with Pipetman. Do not use vacuum aspiration, as it will disturb the monolayer. Wash three times with warm adhesion buffer (Hanks balanced salt solution, 10 mM HEPES, pH 7.4, 2 mM MgCl2, 2 mM CaCl2, 0.2 mM MnCl2, 1% BSA). Add warm adhesion buffer to wells and remove gently with a Pipetman. Remove all medium with Pipetman. Wash once with PBS. Fix with 3.7% paraformaldehyde. Quantify bound myeloid cells by counting fluorescent cells per high-power microscope field (Fig. 15.9). To determine which molecules mediate adhesion of myeloid cells to endothelial cells in vitro, it is possible to perform the above assay using antibody, peptide, or small-molecule antagonists of potential adhesion receptors. We prepare HUVEC monolayers by seeding 200,000 HUVECs per well of a 48-well plate 24 h before the adhesion assay. Label monocytes with 25 mM CMTMR as described above, and wash and resuspend cells at 800,000 cells/ml in adhesion buffer. Remove supernatant from HUVEC monolayers using only a Pipetman to aspirate. Add antibodies at dilutions from 0 to 25 mg/ml in adhesion medium to the monolayer-coated wells. Possible antibodies include anti-VCAM, anti–E- or P-selectin, anti-integrin a4b1, anti-integrin a5b1, anti-integrin alpha v, anti-integrin aM, or anti-integrin b2 antibodies. Place 0.25 ml myeloid cells (200,000) per well (three replicates per antibody). Incubate in 37  C incubator. Aspirate supernatant of three wells per antibody with a Pipetman after 60 min; wash twice with warm adhesion buffer. Remove all medium with Pipetman, wash once with PBS, and fix with 3.7% paraformaldehyde. Quantify bound myeloid cells (fluorescent round cells on top of monolayer) per high-power microscope field.

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Seed 48-well plate (1 ⫻ 105 cells/well) Incubate 30 min to 2h, 378 Wash, fix and analyze using fluorescence microscope Basal media

SDF-1a

Figure 15.9 Myeloid cell^endothelial cell adhesion assay. Human umbilical vein endothelial cells are seeded in EGM-2 on 48-well plates 24 h prior to addition of myeloid cells. Freshly isolated murine or human CD11b-positive cells are labeled with calcein AM. After removal of excess dye, 1  105 cells/well are added to endothelial cell monolayers. Wells are washed 30 to 120 min later with PBS three times to remove unbound cells, and adherent cells are fixed in paraformaldehyde. Bound fluorescent myeloid cells are quantified in five different microscopic fields using MetaMorph software.

2.7. In vitro myeloid migration/invasion assays To determine whether specific extracellular matrix proteins or purified ligands regulate myeloid cell invasion and migration, we perform cell migration assays using Costar Transwells. For haptotaxis assays, coat the undersides of 8-mm Transwell inserts with fibronectin, CS-1 fibronectin or vitronectin (5 mg/ml) for 12 h at 4  C. Block nonspecific binding sites by incubation with 3% BSA in PBS for 1 h at 37  C. Resuspend cells in migration buffer (Hepes-buffered M199 medium containing 1% BSA, 1.8 mM CaCl2, 1.8 mM MgCl2, and 0.2 mM MnCl2, pH 7.4) and 50,000 cells are added to the upper chamber and incubated at 37  C, 5% CO2. Allow cells to migrate from the upper to the lower chamber for 4 h at 37  C. Remove

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nonmigratory cells from the upper chamber by wiping the upper surface with an absorbent tip. Fix the transwell inserts for 15 min with 3.7% paraformaldehyde for 15 min and incubate in a 2% crystal violet in sodium borate. After extensive water washing to remove excess crystal violet, count the number of cells that migrate to the bottom of the insert in five random 200 fields per replicate. These assays can also be performed with fluorescently labeled cell populations. Perform these assays three times with triplicate samples per group. Calculate the mean number of cells migrating plus/minus the standard error mean for the entire treatment group in each experiment. As basal migration of unstimulated monocytes can be low, monocytes can be prestimulated by incubation in purified chemokines or conditioned medium as described above. To determine which integrins mediate migration of particular matrix protein, add function-blocking integrin antibodies as described above to both chambers of the Transwell. To test the chemotactic response of myeloid cells to particular chemokines, coat both sides of the Transwell filter with ECM proteins before performing the migration assay (Fig. 15.10).

2.8. In vivo monocyte-trafficking studies To determine whether monocytes traffic to the tumor microenvironment and to elucidate mechanisms regulating their adhesion and invasion into tissues, we perform intravital microscopy to track the movement of fluorescently labeled myeloid cells in tumor blood vessels. To accomplish these in vivo homing assays, label murine or human peripheral blood myeloid cells or purified CD11bþ cells with CMTMR, as described above and resuspend cells at 800,000 cells/ml in culture medium. To test the roles of various integrins in cell homing to tumor neovasculature, incubate 0.5  106 CMTMR-labeled cells in EBM basal culture medium or medium containing 0 to 50 mg/ml of low endotoxin function blocking anti-integrin antibodies on ice for 30 min. Inject cells by tail vein injection in a final volume of 100 ml into anesthetized nude mice bearing murine lung or breast carcinomas implanted under dorsal skinfold windows. Capture digital images of the tumor vasculature in the dorsal skinfold chamber every few minutes over a 1-h period. After 1 hour, sacrifice animals (n ¼ 3 to 5) and excise tumors for cryo-preservation. Immunostain tissue sections of cryopreserved tumors to detect CD31. Quantify CD31þ blood vessels and CMTMRþ monocytes per high-power field using MetaMorph software, and calculate the average microvessel density in all tumors per treatment group ( Jin et al., 2006a,b). An alternate strategy to quantify short-term monocyte trafficking to tumors is to quantify fluorescently labeled monocytes in single-cell populations of tumors by Facs analysis. To accomplish this, isolate CD11bþ bone marrow–derived cells from C57B6 female animals as described above.

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Insert

A Upper chamber

Well

Lower chamber

B

Coat underside of insert with ligand (5 mg/ml). 12 h,48

Block with 3% BSA, 1 h, room temperature

Add 5 3 104 cells/insert

Incubate 4h, 378

Fix and stain

Count migrated cells

Figure 15.10 Myeloid cell migration assay.The underside of Transwell 8-mm^pore size tissue culture inserts are coated overnight with specific ligands.The next day, inserts are washed once in PBS and 600 ml of culture media are added into the lower chamber. Next, 100 ml of cells (5  104 cells/insert) is added into the upper chamber (insert). Plates are incubated for 4 h at 37  C. Medium is aspirated from the lower chamber and cells are removed from the upper side of the insert using a cotton swab. Cells on the lower side of the insert are fixed and stained with crystal violet, and then quantified.

Resuspend cells in 1  107 cells/ml in PBS and label with 1 mM of CFDA (V12883, Invitrogen) for 10 min at 37  C in water bath. Wash cells twice in PBS and resuspend in a final concentration of 0.5 to 1  107 cells in EBM

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(Cambrex, East Rutherford, NJ). Inject cells by tail vein injection in a final volume of 100 ml into mice bearing murine LLCs. Track labeled cells as they invade the tumor microenvironment by intravital microscopy after 20 min to 48 h. Isolate tumors 24 h postinjection of labeled myeloid cells and chop tumors and spleen (control organ) using a surgical razor. Incubate tissues for 2 h at 37  C in 10 ml of Hanks balanced salt solution (HBSS, GIBCO) containing 10 mg/ml collagenase type IV (Sigma), 1 mg/ml hyaluronidase type V (Sigma), and 200 units/ml DNase type IV (Sigma) according to the method described by Winter et al. (1999). To obtain a pure single-cell suspension, cells are then passed through 70-mm nylon cell strainer (Becton Dickinson, NJ). Cells are centrifuged 400 g for 5 min resuspended in PBS, and incubated for 5 min in RBC lysis buffer (eBioscience, San Diego, CA) at RT to lyse the RBCs. Cells are washed twice in PBS by centrifugation 400 g for 5 min. A total of 5  105 cells are finally fixed in 300 ml 0.5% PFA, and analyzed in the FL-1 channel (488 nm) using flow cytometry (Fig. 15.11).

2.9. Matrigel angiogenesis assays We previously found that myeloid cells embedded in Matrigel can promote angiogenesis even in the absence of added growth factors ( Jin et al., 2006). To explore the recruitment of myeloid cells in vivo in response to chemokines, subcutaneously inject 400 ml of cold Matrigel containing 400 ng/ml chemokines or growth factors such as SDF-1 or VEGF-A (n ¼ 8 to 10 animals). After 7 to 21 days, remove Matrigel plugs, embed in OCT, cryopreserve, and section. Thin sections (5 mm) should be immunostained with anti-CD31 and anti-CD11b or anti-F4/80 antibodies. At least five microscopic fields per tissue section are analyzed for quantification studies. Similar studies can be performed in mutant animals to evaluate the roles of key molecules in monocyte trafficking.

2.10. Role of myeloid cells in tumor growth studies To study the roles of bone marrow–derived cells in tumor growth, we have used several animal models. C57Bl/6 syngeneic mouse tumor models are particularly useful as they can be used to explore tumor growth in mutant mice in this genetic background. LLC cells are cultured in DMEM medium supplemented with l-glutamine and 10% fetal bovine serum. Tumor cells (5  105) in 100 ml PBS are injected subcutaneously into 6-week-old syngeneic C57/Bl6 mice (LLC). A time course of tumor growth and myeloid cell invasion can be performed from 1 to 28 days after tumor cell inoculation. In some studies, animals bearing tumors can be treated with potential inhibitors of myeloid cell trafficking. Tumor volume can be

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CD11b + cell isolation LLC growth during 7 days CFDA labeling

CD11b + cells injection (5 ⫻ 106 cells/animal)

20 min to 48 h incubation Tumor isolation

Enzymatic digest

Log fluorescence

Without labeled cells injection

With labeled cells injection

0.16%

0.00%

Flow cytometry

Forward scatter

Figure 15.11 Quantification of short-term, myeloid cell^trafficking to tumors.Tumor cells (5  105) are injected subcutaneously into wildtype mice. After 7 days, freshly isolated myeloid cells are labeled with fluorescent tracking dyes and injected into the tail vein (5 106 cells/animal).Twenty-four hours later, tumors are harvested and singlecell suspensions are made. Cells are fixed and fluorescently labeled cells are detected in the FL-1 channel. As a negative control, animals with tumors but no adoptive cell transfer are used.

determined after measuring length and width with calipers, according to the formula (l2  w)/2. After tumor removal, tumors can be dissociated into single-cell suspensions for analysis by fluorescence-activated cell sorting (FACS) or embedded in OCT, frozen and sectioned for immunohistological analysis of myeloid cell and blood vessel density. At least five microscopic fields per tissue section per animal should be analyzed for quantification studies. Mean myeloid cell and blood vessel density should

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be determined after quantifying positive staining on 5 to 10 microscopic fields per animal. Each study should be performed two to three times.

2.11. Quantification of myeloid cells in experimental tumors by FACS, qPCR, and IHC To evaluate the infiltration of myeloid cells in the tumor microenvironment, we inject LLC cells (5  105 cells in 100 ml PBS) subcutaneously in 4- to 6-week-old female C57B6 animals. After days 7, 14, and 21, mice are euthanized and tumors harvested. Tumor-derived, single-cell suspensions are prepared as described above (Fig. 15.12). Upon RBC lysis with RBC lysis buffer (eBioscience, San Diego, CA), centrifuge cells at 400 g for 5 min and wash once in PBS. Block nonspecific binding by adding 5 ml/2.5  106 cells of FC-Block and incubate the cells for 10 min on ice. Incubate cells in 2.5 ml of CD11b-APC (M1/70, eBioscience, San Diego, CA) per 5  105 cells or isotype-matched rat IgG1-APC control antibody for 60 min at 4  C with gentle shaking (light protected). Wash cells twice in cold PBS and fix in 0.5% PFA. Analyze by flow cytometry (Fig. 15.13). To quantify the relative infiltration of myeloid cells by quantitative RTPCR, chop approximately 100 mg of tumor tissue with a razor blade and then transfer into 1.5 ml of ISOGEN solution (Nippon Gene, Toyama, Japan). Use 100 mg of muscle as a control. Homogenize tissues with a Tissue Tearer (Biospec, Inc.) using 10 pulses of 5 s on ice. Isolate total RNA using ISOGEN according to manufacturer’s directions. Treat two micrograms of A

Enzymatic digest

Immunostain Flow cytometry

B

Chopping

Total RNA isolation Quantitative RT-PCR

C

Cryopreservation

Section and immunostain Fluorescence microscopy

Figure 15.12 Methods to analyze monocyte trafficking to tumors. Myeloid cell infiltration into primary tumor can be quantified from single-cell suspensions by three different methods: (A) flow cytometry, (B) qPCR, and (C) immunohistochemistry.

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CD11b-APC 1023

0

100 101

102 103 104

0

1023

34.61%

SSC-height

0.01% SSC-height

Side scatter

1023

Day 7

100 101

FL4-H

102 103 104 FL4-H

Day 14

SSC-height

IgG-APC

0 100 101

46.36% R2

102 103 104 FL4-H

Log fluorescence

CD11b expression (⫻104 AU)

4 3 2 1 0 Day 0

Day 7

Day 14

Day 21

Figure 15.13 Evaluation of myeloid cell infiltration in the tumor microenvironment by FACS. After 7, 14, and 21 days of tumor growth, mice are euthanized and tumors harvested. Single cell suspensions are immunostained with anti-CD11b-APC or isotype-matched rat IgG1-APC control antibody and then analyzed by flow cytometry. The percent of CD11b-positive cells in the tumor suspension at each time point can then be compared (graph).

total RNA with DNase (Invitrogen) and perform reverse transcriptasepolymerase chain reaction (RT-PCR) using Invitrogen Superscript First Strand Synthesis kit (Invitrogen) according to the manufacturer’s directions. We test cDNA for genomic DNA contamination by PCR using b-actin primers (forward: ACCACAGCTGAGAGGGAAATC, reverse:AGAGGT CTTTACGGATGTCAACG) resulting in a 277-bp product for cDNA and 372-bp product for genomic DNA. We perform real-time PCR reactions for GAPDH (sense CCTGCACCACCAACTGCTTA; anti-sense TCAT GAGCCCTTCCACAA; probe FAM-CCTGGCCAAGGTCATCCACBHQ1) were performed using Absolute QPCR Mix (ABgene, Rochester, NY). We perform real-time PCR reactions for CD11b (QuantiTect Primer Assay, Qiagen) using Power SYBER-Green PCR Master Mix (Applied Biosystems, Foster City, CA). We perform and analyze real-time PCR reactions using a Cepheid (Sunnyvale, CA) SmartCycler. We normalize samples from tumor tissue to GAPDH and determine relative gene expression levels, as previously described (Livak and Schmittgen, 2001). We analyze all samples further by separating the products on an 8% acrylamide gel.

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To study macrophage infiltration of tumors by immunohistochemistry, we remove tumors carefully from euthanized animals and wash tumors for 1 min in PBS. We embed and cryo-preserve tumors in O.C.T (Tissue Tek) using cryo-molds (Tissue Tek). Tumors can be sectioned as described previously and stained using anti-CD11b antibody (M1/70, BD Pharmingen). Analyze at least five microscopic fields per tissue section using MetaMorph software. 2.11.1. Bone marrow transplantation to study myeloid cells in tumor growth To evaluate the role of monocyte trafficking to tissues, including tumors, it is useful to perform transplants of bone marrow from GFP-expressing mice or from mice bearing mutations in integrins or chemokine receptors. To prepare mice for bone marrow transplants, treat recipient mice with a suspension of 800 mg/l sulfamethoxazole and 160 mg/l trimethoprim (SMZ) for 1 week prior to irradiation and 2 weeks after bone marrow transplantation. To isolate bone marrow for transplantation, euthanize donor mice and sterilize the skin with ethanol. Aseptically remove femurs, using separate sterile surgical tools for cutting soft tissues and bones. Trim muscle tissue from femurs. Cut off the ends of each bone and flush with PBS, pH 7.2, supplemented with 0.5% BSA and 2 mM EDTA (2 ml/femur and 1 ml/ tibia) into plastic centrifuge tubes on ice. Triturate to break up clumps and form single-cell suspensions. Centrifuge at 400 g for 10 min at RT. Resuspend in 10 ml red-cell lysis buffer (155 mM NH4Cl, 10 mM NaHCO3, and 0.1 mM EDTA) and incubate for 4 to 5 min at RT. Transfer cells to a 50-ml centrifuge tube and add 30 ml PBS without MgCl2 or CaCl2. Centrifuge 400 g 10 min at RT. Resuspend in 4 ml PBS, pH 7.2, supplemented with 0.5% BSA and 2 mM EDTA, and transfer to 15-ml Falcon tube. Add 4 ml of room-temperature Histopaque 1083 to the bottom of the tube. Centrifuge 400 g for 25 min without brakes. Aspirate surface layer, then transfer cloudy layer containing the purified bone marrow cells into a new 15-ml centrifuge tube. Wash twice with PBS by resuspending in PBS and centrifuging at 400 g for 10 min at 4  C. Add ice-cold PBS without MgCl2 or CaCl2. Resuspend at 2.5  107 cells/ml in PBS (store on ice until ready for injection into recipient mice). The average yield of mononuclear bone marrow–derived cells is 2  107 cells per animal. Irradiate female mice of the C57Bl6 genotype (6 weeks of age) with a single dose of 1000 rad (10 Gy). Bone marrow should be transplanted within the same day. Warm the donor bone marrow cells to RT before injecting into recipient mice. Inject RT bone marrow cells into irradiated recipient mice by tail vein injection at 5  106 cells in 200 ml per animal.

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Monitor animals for 4 weeks after bone marrow transplantation. Animals will be fully recovered and ready for in vivo angiogenesis and tumor studies after 4 weeks. To compare the effects of key mutations in myeloid cells on tumor growth and angiogenesis, perform reciprocal transplantations between wildtype (WT) animals and mutant (MUT) animals (WT!MUT and MUT!WT). Also perform WT!WT and MUT!MUT bone marrow transplants to control for the effects of irradiation and transplantation. Implant tumor cells as described above and evaluate tumor growth, angiogenesis, and myeloid cell invasion over time (from 0 to 28 days) using the methods described previously (Fig. 15.14).

Recipient

Donor

Femur/ tibias

SMZ treatment, 1 week

Cut off ends of bones

1000 rad

Flush out bones with syringe

BMDC injection (5 ⫻ 106 cells/animal) Recipient 4 weeks reconstitution (2 week with SMZ) Tumor cells can be implanted

Figure 15.14 Bone marrow transplant methods. Recipient animals are treated for 1 week with antibiotics (SMZ). Mice are lethally irradiated, and freshly isolated bone marrow derived cells (5 106) are injected into tail veins of irradiated animals. Animals are treated for another 2 weeks with antibiotics, and then kept for 2 weeks without antibiotics. Successfully transplanted animals can be used afterwards for tumor studies.

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3. Conclusions Myeloid cells play important tumor-initiating and -promoting roles. The methods outlined here will assist new researchers to explore the study of tumor-associated myeloid cells.

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Author Index

A Abbruzzese, J., 144, 147 Abe, A., 343 Abernathy, C., 342 Aboagye, E. O., 167 Abramovitch, R., 55, 56, 302 Abrams, J. S., 282, 283 Abramsson, A., 56, 67, 212 Abu-Issa, R., 211, 215 Achen, M. G., 4, 5 Acton, S. T., 257 Adair, T. H., 234 Adamis, A. P., 127 Adamson, S., 223 Adebahr, S., 95 Ades, E. W., 74 Adkins, J. N., 186, 190 Aebersold, R., 181, 182, 183, 189, 190 Agarwal, A., 88, 89 Ager, A., 263 Aghi, M., 54, 56 Agus, D. B., 128 Ahmad, S., 245, 248 Ahmed, A., 245, 248 Ahn, G. O., 55, 57 Ahn, N. G., 186 Aiello, L. P., 127 Aimes, R. T., 149 Aird, W. C., 192 Aitken, R. P., 245 Aiyer, A., 343, 344, 352, 359, 361 Akamatsu, H., 343 Akerman, M. E., 85, 98, 193 Akhtar, N., 318, 322 Akita, N., 86 Alauddin, M. M., 89, 90 Alby, L., 194 Aldo, P. B., 292 Alencar, H., 144, 147 Alessandri, G., 28 Alferez, D., 131 Algire, G. H., 126 Aliberti, J., 263, 264 Alison, M., 130 Alitalo, K., 2, 3, 4, 5, 8, 9, 10, 37, 54, 56, 282, 343 Allen, E., 302 Allen, J. S., 150, 151 Allport, J. R., 88

Allt, G., 54 Al-Mehdi, A. B., 29 Alminaite, A., 54 Alon, R., 343 Altomonte, M., 167 Alvero, A., 292 Alves, P., 190 Amann, G., 9, 10 Amano, H., 55 Ameer-Beg, S. M., 144, 145 Amiel, G. E., 305 Amin, M. A., 5 Amit, M., 304 Amoh, Y., 36, 37 Amos-Landgraf, J. M., 130 An, P., 5 Anderle, M., 190 Anderson, D. A., 54 Anderson, D. C., 193 Anderson, N. L., 182 Anderson, R. G., 195 Ando, K., 210 Andrusiak, R., 87 Angel, P., 6 Angelberger, P., 157 Angeli, V., 4, 10 Ansiaux, R., 129 Antczak, C., 151 Apel, I. J., 302 Aplin, J. D., 281, 288, 293, 294 Arap, M. A., 86, 87, 88, 95 Arap, W., 83, 84, 85, 86, 87, 88, 89, 90, 91, 94, 95, 97, 98, 100, 192, 193 Arboleda, C. E., 288 Arbones, M. L., 257 Ardelt, P. U., 85, 86, 88, 95, 98 Arfors, K. E., 254, 260, 261 Armanini, M., 127 Arnaout, M. A., 149, 367 Arnold, D. R., 234, 235, 236, 237, 238, 239, 241, 243, 244, 245, 246 Arnold, P., 29 Arnold, R. J., 190 Arsic, N., 302 Asahara, T., 54, 303, 305, 342 Asakura, H., 262 Ashmun, R. A., 85, 86, 98 Ashton, S. V., 293, 294 Assheton, R., 244

373

374

Author Index

Atala, A., 305 Atencia, D., 211 Athey, P., 151 Atkinson, B. A., 244 Atkinson, M. A., 88, 89 Ator, E., 4 Au, P., 303, 306 Audenaert, K., 156 Auerbach, R., 28, 29, 194, 303, 304, 318, 322 Ausprunk, D. H., 303 Autiero, M., 330, 331, 335 Avecilla, S. T., 54, 55, 84 Avitabile, D., 89 Avraham, H., 4 Avraham, I., 55, 56, 302 Avraham, S., 4 B Baatout, S., 319 Bachar-Lustig, E., 55, 56, 302 Bachmann, G., 339 Bacich, D. J., 86 Backer, J. M., 155, 156, 158, 160, 161 Backer, M. V., 155, 156, 158, 160, 161 Baczyk, D., 293, 295 Baez, S., 254 Baggerly, K. A., 85, 86, 98, 100 Baghi, M., 144, 146 Bai, K., 158 Bai, W., 130, 131, 133 Bailey, A. S., 54 Bailey, J., 167 Bailey, K. R., 89 Baird, A., 89 Baker, A. H., 88, 89 Baker, J., 144, 145 Baker, M., 256 Baker, P. N., 293, 294 Bal, N., 343 Balakin, K. V., 126, 127 Baldwin, M. E., 128 Ballara, S., 5 Baluk, P., 4, 8, 9, 14, 144, 145 Bander, N. H., 86 Bando, H., 343 Bandyopadhyaya, A. K., 156 Banerji, S., 3, 9, 10 Bankson, J., 144, 146 Baranov, E., 30, 31, 46 Barateau, V., 322 Barber, P. R., 144, 145 Barker, D. J., 245 Barleon, B., 343 Barnhart, K. T., 296 Baron, A., 134 Barreto Gadelha, E., 233 Barron, B., 144, 147

Barron, D. H., 230, 233 Bar-Shavit, R., 282 Bartel, G., 4 Barth, R. F., 156 Barthel, H., 167 Bartlett, J. S., 89 Bartunkova, S., 144, 145 Basevitch, Y., 304 Bassingthwaighte, J. B., 234 Bastidas, N., 56 Battaglia, F. C., 233 Battisti, C., 262 Battler, A., 95 Baudelet, C., 129, 144, 146 Baudin, B., 194 Baumhueter, S., 9, 10 Baval, B., 322 Bazer, F. W., 248 Beasley, N. J., 3 Beaudet, A. L., 257, 260, 261 Beavis, R. C., 187, 190 Beck, I., 6 Becker, A., 156 Becker, C. G., 194 Becker, C. H., 190 Beckett, M. A., 129 Bednarski, M. D., 150 Bee, J., 214 Beheshti, M., 157 Belagaje, R., 331 Belaguli, S., 211, 215 Bell, A. W., 234 Bellingard, V., 282 Belton, O., 183 Bemis, L., 245 Benezra, R., 55, 87 Benirschke, 292 Benjamin, L. E., 14, 142, 143 Benjamin, R. S., 90 Ben Mahdi, M. H., 5 Berg, E. L., 9, 10 Berg, T., 95 Bergers, G., 53, 54, 55, 56, 57, 59, 64, 67, 68, 73, 85, 132, 133, 142, 143 Bergmann, F., 343 Bergmann, J., 194 Bergsland, E., 67, 132 Berk, D. A., 37 Berkovic, S. F., 156 Berkowitz, R., 28, 292 Berleau, L. T., 126, 161 Berlin, J., 134 Bernasconi, M., 85, 86, 97, 98, 193 Berthier, R., 8, 9 Besmer, P., 55 Betsholtz, C., 55, 56, 67, 212 Bevilacqua, G., 343 Beyer, T., 148, 158

375

Author Index

Bhardwaj, J., 367 Bhatia, M., 304 Bhushan, M., 5 Bianco, A., 151 Bibby, M. C., 29 Bicknell, R., 167, 193 Biensen, N. J., 245 Biglioli, P., 56 Binder, M., 92 Binderup, L., 322 Biondini, M. E., 239, 243, 244 Birner, P., 4 Bischoff, J., 301, 304, 305, 306, 314, 315, 318, 319, 320, 322 Bisdas, S., 144, 146 Bjarnegard, M., 212 Blaikie, P., 54, 342 Blair, R., 303 Blank, M., 5 Blankenberg, F. G., 155, 156, 158, 160, 161 Blankenship, T. N., 282, 289 Blavier, L., 343 Bleeker, E. J., 152 Bloch, S. H., 154 Bock, M., 144, 145, 150 Bodenstein, C., 88 Bodmer, W. F., 130 Bogenrieder, T., 143 Bogers, A., 210, 212 Boggi, U., 343 Bohlen, P., 4, 5, 6, 10, 15, 55 Bojarski-Nagy, K., 4 Bokenkamp, R., 211 Bolat, F., 343 Bondarenko, P. V., 190 Borges, L. P., 5 Borgna, L., 87 Borgstrom, P., 343, 344, 352, 359, 361 Borlak, J., 194 Bornstein, P., 14 Borowicz, P. P., 229, 230, 231, 232, 233, 234, 235, 236, 237, 238, 239, 241, 243, 244, 245, 246, 247, 248 Bortnem, B. J., 235, 248 Boscolo, E., 306, 315 Bosse, D. C., 74 Bosse, R., 257 Bothwell, A. L., 303 Bouchard, T. S., 352 Bouvet, M., 27, 30, 31, 36, 38, 39, 40, 41, 44, 45, 46 Bover, L., 95 Boxall, J., 144, 146 Boyer, D., 29 Brady, F., 167 Brahmer, J., 134 Brain, N. J. R., 89 Bramm, E., 322

Brand, T., 211, 215 Brandt, U., 330 Brasch, R. C., 144, 146 Braun, R. D., 30 Braverman, I. M., 4 Bray, F., 142 Brechbiel, M. W., 144, 145 Bredesen, D. E., 85, 86, 87, 97, 98, 193 Breier, B. H., 245 Breier, G., 8, 9, 126 Breiteneder-Geleff, S., 9, 10 Brekken, R., 55, 132, 133 Brekken, R. A., 154, 155 Brentjens, R., 151 Breviario, F., 8, 9 Brignole, C., 87 Bristow, J., 210, 211 Brock, C. J., 182 Broermann, R., 9, 10 Bronner, M., 282 Bronte, V., 343 Brooks, P. C., 149 Brosen, I., 289 Brosnan, M. J., 89 Broughton, L., 167 Brouwers, A. H., 167 Brown, G. G., 144, 146 Brown, J. M., 55, 57 Brown, L. F., 4, 6, 8, 9, 10, 12, 14, 15 Browne, F., 303 Brun, T., 148, 158 Bruneel, A., 194 Brutsaert, T., 245 Buchholz, C. J., 89 Buck, F., 194 Bullard, D. C., 257, 261 Buning, H., 89 Bunt, S. K., 55 Bupp, K., 89 Burg, M. A., 86 Burger, C., 144, 147 Burgers, P. C., 197 Burns, P. N., 144, 147 Burrows, M. T., 268 Burton, G. J., 289 Butcher, E. C., 9, 10, 263, 264 Butler, J., 55 Butler, J. P., 190 Butros, L., 54, 342 Butz, S., 8, 9 Byars, L., 148, 158 Bylund, D. D., 190 Byzova, T. V., 149 C Caban˜as, C., 352 Cagney, G., 183, 189, 191

376 Cahill, D. J., 183 Cai, W., 141, 143, 149, 150, 151, 152, 153, 156, 158, 159, 160, 161, 162, 163, 166, 168 Calias, P., 127 Callaghan, M. J., 56 Calvin, D. P., 129 Cammer, M., 344 Campa, C., 126, 127, 128, 133 Campbell, J., 263, 264 Campochiaro, P. A., 127 Candal, F. J., 74 Caniggia, I., 288 Cao, J., 5, 268 Cao, Q., 158, 159, 160, 161, 162, 163, 166, 168 Cao, Y., 2, 3 Cao, Y. A., 158 Capla, J. M., 56 Caplice, N. M., 314 Capogrossi, M. C., 56 Carbon, J. G., 154, 155 Carbone, D. P., 54, 55 Cardo-Vila, M., 85, 86, 87, 94, 95, 98 Carmack, E., 183 Carmeliet, G., 56 Carmeliet, P., 55, 56, 125, 126, 213, 302, 330, 342 Carmona, R., 211 Carpenter, J. E., 151 Carpizo, D., 126 Carradine, C., 151 Carroll, V. A., 167 Carson-Walter, E. B., 86 Cartegni, L., 87 Carter, C., 127 Carter, W. G., 352 Cartwright, J. E., 293, 294 Cartwright, T., 134 Caruthers, S. D., 150, 151 Carvalho, J. F., 5 Carver, L. A., 179, 191, 193, 194, 195, 196, 197 Carver-Moore, K., 126 Casanovas, O., 132, 133 Case, J., 305 Catana, C., 148 Catimel, B., 5 Caton, J. S., 230, 231, 232, 233, 234, 236, 237, 238, 239, 243, 244, 245, 247, 248 Cattaneo, A., 87 Cavagna, F. M., 144, 146 Cavallari, A., 262 Caveda, L., 8, 9 Caze, R., 292 Cella, D., 134 Ceradini, D. J., 56 Cerdan, C., 304 Cerem, L. W., 5 Chadburn, A., 54, 342 Chadwick, M. P., 89

Author Index

Chait, B. T., 189 Chakel, J. A., 186 Chalkley, H. W., 126 Challice, C., 210, 211 Chamley, L. W., 292 Chan, C., 158 Chan, K. Y., 245 Chan, L., 85, 87, 98, 193 Chance, B., 156 Chandler, 246 Chandrasekharappa, S. C., 131 Chang, C. N., 144, 146 Chang, S. S., 86 Chang, W. J., 195 Chantrain, C. F., 343 Chaplin, D., 55 Chaplin, D. J., 144, 145 Chapman, S. C., 115, 116 Charara, J., 268 Charif, H., 197 Charnock-Jones, D. S., 230, 234, 244, 245 Charnsangavej, C., 144, 147 Charron, M., 148, 158 Chatterjee, S., 305 Chattopadhyay, D., 151 Chatziioannou, A. F., 158 Chau, T. C., 144, 145 Chaubal, A., 144, 145 Chawengsaksophak, K., 342 Cheezum, M. K., 263 Chelius, D., 190 Chen, C., 127 Chen, G. T., 282 Chen, H., 126, 127, 161 Chen, H. H., 268 Chen, I. Y., 154 Chen, K., 152, 153, 154, 158, 159, 160, 161, 162, 163, 166, 168 Chen, K. S., 130 Chen, L., 5, 85, 86, 88, 95, 98 Chen, P., 158 Chen, Q., 292 Chen, S., 155 Chen, T. H., 314 Chen, W. M., 314 Chen, X., 141, 143, 149, 150, 151, 152, 153, 154, 156, 158, 159, 160, 161, 162, 163, 166, 168 Chen, Y., 29, 133, 331 Cheng, X., 270 Cheng, Z., 128, 133, 151, 162 Cheresh, D. A., 149, 150 Cherry, S. R., 148 Cherubini, G., 89 Chesnutt, B. C., 268, 269, 270 Cheung, A. M., 55 Cheung, A. N., 296 Cheuvront, T. J., 108, 113, 115 Chevalllet, M., 182

377

Author Index

Chidlow, J. H., 4 Chien, S., 256, 257, 258, 268, 354 Chimenti, S., 55, 56, 302 Chin, Y., 87 Chinen, L. K., 150 Chiocca, E. A., 54, 56 Chirivi, R. G., 193 Chittenden, T. W., 330, 331, 335 Chiu, K., 280, 282 Cho, H. J., 268 Choi, J. H., 305 Choi, J. Y., 88 Choi, K., 212 Chomczynski, P., 216 Chou, C. L., 190 Choyke, P. L., 143, 144, 145, 146 Christian, S., 85, 98, 193 Christianson, 246 Christianson, D. R., 88 Christinger, H. W., 133 Christofori, G., 132 Christopher, D. A., 144, 147 Chu, T., 88 Chudek, J., 131 Chung, C., 189 Ciarrocchi, A., 55 Cichutek, K., 89 Cilli, M., 87 Cines, D. B., 288 Claes, F., 330, 331, 335 Claesson-Welsh, L., 18, 126 Claffey, K. P., 4, 14, 155, 156, 158, 160, 161 Clark, P. M., 245 Clasper, S., 9, 10 Clauss, T. R., 190 Clavel, F., 296 Cleaver, O., 302 Clements, V. K., 55 Clemmer, D. E., 190 Clendenin, C., 222 Clifton, J. G., 195 Cobleigh, M., 134 Cohen, I. R., 2, 5 Cohen, K. S., 56, 306 Cohen, P., 282 Cohen, S., 304 Cohen, T., 5 Colarusso, P., 267 Cole, N., 191 Coleman, R. E., 156 Collen, D., 302 Collignon, J., 115, 116 Collingridge, D. R., 167 Collins, D. J., 144, 146 Collins, T., 212 Condeelis, J., 55

Conlon, T. J., 88, 89 Connolly, L. M., 186 Connors, M. A., 5 Conrad, G., 214 Conrads, T. P., 189 Contag, C., 158 Contreres, J. O., 322 Conway, E. M., 302 Cook, C. M., 244 Cook, R. F., 195 Coppinger, J. A., 183 Corada, M., 8, 9 Coram, M., 190 Corbly, A., 331 Cordon-Cardo, C., 128 Cornelissen, B., 156, 157 Cornelssen, C., 268 Corry, D. B., 282, 283 Cortens, J. P., 187, 190 Corthals, G.L., 182 Corti, A., 87 Corwin, N. M., 280, 282 Costa, C., 54, 342 Cotran, R. S., 29, 126, 302 Coultas, L., 342 Coussens, L. M., 55, 67, 343, 344 Coutifaris, C., 282 Cowburn, D., 189 Cowen, S. E., 29 Cowin, S. C., 256 Cox, D., 344 Cox, G., 3 Crabtree, J. S., 131 Craig, R., 187, 190 Crawford, Y., 125, 126, 129 Crispino, J., 212 Crocker, I. P., 293, 294 Crokart, N., 129 Cron, G. O., 144, 146 Crooks, G. M., 343 Cross, F. R., 189 Cross, J., 223 Crum, R., 28 Crystal, R. G., 55 Cudd, T. A., 248 Cueni, L. N., 2, 3, 8 Cui, C., 107, 108, 113, 115 Cui, T. Y., 280 Cundari, E., 89 Cunningham, B. C., 133 Cunningham, E. T. Jr., 127 Curiel, D., 89 Curnis, F., 87 Cursiefen, C., 5 Cusick, R. A., 303 Cybulsky, M. I., 258 Czirok, A., 108, 109, 110, 111, 113, 116

378

Author Index

D Dadras, S. S., 3 Daheron, L. M., 306 Dai, H., 151, 166 Damiano, E. R., 256, 263, 269 Damico, L., 133, 134 D’Amore, P. A., 5, 127 Damsky, C. H., 280, 281, 282, 291 Dana, M. R., 5 Danese, S., 6 Danielsson, R., 190 d’Arcy, J. A., 144, 146 Dash, P. R., 293, 294 Davidoff, A. M., 129 Davidson, N. E., 134 Dawson, J., 5 Day, K., 263, 264 Dayton, P. A., 154 De, S., 149 De Bandt, M., 5 DeBusk, L. M., 54, 55 de Candia, P., 87 Deciu, C., 189 Declerck, Y. A., 343 Deconinck, A., 212 Dedkov, E., 330, 331, 335 Deeks, S. G., 296 Deen, D. F., 128 De Falco, E., 56 De Gunzburg, J., 195 Deheregoda, M., 130 Dejana, E., 8, 9, 281, 282 de Jong, J. R., 167 Dekker, L. J., 197 de la Motte, C., 6 Deleage, G., 88 deLima-Hahn, E., 89, 95 Delorme, S., 144, 145 DeMaio, E., 2 DeMars, R., 282, 291 Demetri, G., 127 Demirci, U., 270 Demirev, P. A., 189 Demirhan, B., 343 Demol, H., 183, 189 Demsar, F., 144, 146 de Muinck, E. D., 331 Denetclaw, W., 210, 211 Deng, Y., 343 Denk, W., 267 Denny, R., 191 De Palma, M., 54, 55, 302 DeRuiter, M., 210 Desclaux, D., 181 Detmar, M., 1, 2, 3, 4, 5, 6, 8, 9, 10, 12, 14, 15 Detmar, M. J., 14 Detre, J. A., 144, 146

Dettman, R., 210, 211 Deutsch, T. A., 28 Deutsch, U., 144, 145 de Vos, A. M., 133 de Vries, E. G., 167 Devy, L., 303 de Waal, R. M., 144, 145 DeWever, J., 129 Dewhirst, M., 29 Dewhirst, M. W., 30 Deyanova, E. G., 190 De Zenzo, G., 89 Diamanti, C. I., 163 Dias, S., 54, 55, 342 Dias-Neto, E., 84, 89 Dicker, A. P., 144, 147 Dickler, M., 134 Dickson, P. V., 129 Diefenbach, B., 149 Diem, K., 9, 10 Dieterlen-Lievre, F., 214 Dijkman, H., 86, 88 Dilger, I., 367 Di Maio, M., 2 Dimitrakopoulou-Strauss, L. G., 144, 147 Ding, H. A., 343 Ding, I., 5 Dings, R. P., 129 Di Paolo, D., 87 DiPietro, L. A., 5 Dishart, K. L., 89 Dixon, M., 270 Dmitriev, I., 89 Do, K. A., 85, 86, 95, 98, 100 Dobos, J., 54 Dobrila, F., 343 Doe, C. Q., 9, 10 Doillon, C., 268 Dome, B., 54 Dominiczak, A. F., 88 Donate, F., 6 Dong, C., 268 Dong, J., 128, 133 Dong, X., 211, 215, 216 Dong, X. R., 209 Donnelly, E. F., 144, 145, 147 Donner, A., 297 Dor, Y., 55, 56, 302 Doraiswamy, V., 235, 248 Dorschel, C., 191 Double, J. A., 29 Doumas, P., 182 Dove, W. F., 130 Dowd, M., 126 Dowlati, A., 134 Drinane, M., 330 Drobnjak, M., 128 Droegmueller, W., 233

379

Author Index

Du, R., 55, 56, 57, 59, 64 Duarte, C. M., 161 Dubey, P. D., 263, 264 Duda, D. G., 32, 306 Dudczak, R., 157 Dudek, A., 305 Duling, B. R., 254, 256, 261 Dumas, G., 268 Dumont, D., 211 Dunk, C., 293, 295 Dunker, R., 149 Dunn, J. F., 338 Dunn, M. J., 180, 182 Dunne, J. L., 260 Dunson, D., 268 Dupuis, M., 5 Durr, E., 179, 191, 193, 194, 196, 197 Dutrochet, H., 254 Dutt, S., 6 Dvorak, A. M., 10, 14, 195, 196 Dvorak, H. F., 10, 14, 126 Dwek, R. A., 182 Dwyer, A. J., 144, 146 Dybdal, N., 9, 10 E Eastham-Anderson, J., 131 Ebata, N., 8 Eberhard, D. A., 128 Eberhardt, C., 126 Ebrahimnejad, A., 168 Eckhoff, B., 95 Edelberg, J. M., 192 Edgemon, K. A., 131 Eferl, R., 6 Egeblad, M., 55 Ehrhardt, A., 88 Ehrhardt, R. A., 234 Eisenhut, M., 150 Elfenbein, A., 330, 331, 335 Elicker, B. M., 4, 14 Ellegala, D. B., 151 Ellerby, H. M., 85, 86, 87, 97, 98, 193 Ellerby, L. M., 87 Ellis, L. M., 154 Ellis, S. A., 282 Elner, S. G., 5 Elner, V. M., 5 Eltoum, I. A., 87 Emili, A., 183, 189 Emmert-Buck, M. R., 131 Enders, A. C., 282, 289 Eng, J. K., 183, 186, 189 Enge, M., 212 Engelson, E. T., 254 Enholm, B., 5 Enis, D. R., 303

Ens, W., 189 Enssle, R., 89 Enzelberger, M., 272 Epstein, R. J., 28 Era, S., 343 Eralp, I., 210 Ergun, S., 305 Eriksson, E. E., 265 Eriksson, U., 343 Erlebacher, A., 288, 296 Eskin, S. G., 268 Esposito, I., 343 Esser, R., 5 Essler, M., 85, 98 Esteves, R. A., 280, 282 Evans, D. B., 126 Even-Ram, S., 282 Even-Sapir, E., 148 Ewald, A. J., 54, 55, 57, 67, 68, 73 F Faber, J. J., 244 Fahrig, M., 126 Faivre, S., 127 Fan, S. T., 126 Fang, S. Y., 144, 146 Farber, E. M., 6 Farruggia, F., 262 Fassler, R., 212 Faust, N., 263, 264, 266 Federspiel, M. J., 89 Fehrenbacher, L., 134 Feinberg, 282 Feldmann, M., 5 Feng, D., 10, 14 Feng, K. T., 282 Fenn, J. B., 184 Fenoglio, A., 305, 306, 309, 311, 313, 314, 315 Fenselau, A., 29 Fenselau, C., 189 Fenton, B. T., 37 Fenyo, D., 186 Ferara, N., 126, 129 Ferguson, P. L., 190 Ferkowicz, M. J., 305, 306, 309, 311, 313, 314, 315 Ferlay, J., 142 Feron, O., 129 Ferrara, K. W., 154 Ferrara, N., 4, 5, 125, 126, 127, 128, 129, 130, 131, 132, 133, 149, 153, 154, 160, 161, 196 Ferreira, V., 126 Ferrell, C. L., 233, 234 Ferrier, C., 244 Ferris, B., 55 Ferris, G. R., 293 Fidler, I. J., 367

380 Fiebig, E., 260 Fiebiger, E., 4, 6, 10, 15 Fiederlein, R. L., 264, 268 Figueiredo, J. L., 144, 147 Filla, M. B., 107, 108, 113, 115 Fingleton, B., 54, 55 Fink, C., 144, 145 Finkel, T., 305 Fiocchi, C., 6 Fisher, A. B., 29 Fisher, D. R., 162 Fisher, K., 235, 248 Fisher, K. D., 86 Fisher, S. J., 279, 280, 281, 282, 283, 288, 291 Fishman, D., 210, 211 Fitzgerald, D. J., 183 Fitzgerald, M. L., 280, 282 Fitzgibbon, M. P., 190 Flamm, A., 85, 86, 98 Flamme, I., 302 Fleischer, A. C., 144, 145, 147 Fleming, J. B., 154, 155 Fleming, W. H., 54 Flint, A. P. F., 231 Fliss, D. M., 148 Florey, O., 263 Florin, L., 6 Flotte, T. R., 88, 89 Flower, D., 343 Flunkett, M. L., 6 Foidart, J. M., 157 Foley, E. D., 127 Folkman, J., 2, 28, 29, 84, 125, 126, 127, 142, 143, 302, 303, 342 Fons, P., 303 Fonsatti, E., 167 Ford, S. P., 236, 237, 238, 243, 245 Forlow, S. B., 265 Forrest, M. J., 260 Forrest, W. F., 130, 131 Forsberg, F., 144, 147 Forsyth, K. D., 264, 268 Foster, F. S., 55, 144, 147 Foulk, R. A., 282 Fournier, G. A., 29 Fox, B. A., 361 Fox, W. D., 128 Fraga, C. H., 129 Franco, M., 55 Frangioni, J. V., 156 Frankenne, F., 157 Frantz, G. D., 128, 133 Franza, B. R., 182 Fraser, S. E., 113 Freas, A., 189 Frech, M., 149 Fredriksson, S., 212 Freeman, A., 293, 294

Author Index

French, W., 212 Frenette, P. S., 4, 10 Friedl, P., 143 Friedlander, M., 343, 344, 352, 359, 361 Friedrich, M., 55 Frolich, T., 302 Frost, A. R., 87 Fruttiger, M., 56 Fu, L., 128 Fuchs, A., 182 Fuchs, E., 161 Fuh, G., 127, 128, 130, 131, 133, 134 Fujita, K., 128 Fujita, Y., 305 Fujiwara, Y., 212 Fukuda, K., 54, 55 Fukui, M., 343 Fukumura, D., 4, 5, 14, 29, 303, 306 Fukushi, J., 343 Funel, N., 343 Furth, E. E., 282 Fusenig, N. E., 144, 145 Fuxe, J., 8, 9 Fyles, A., 144, 146 G Gadelha Da Costa, A., 233 Gaehtgens, P., 254, 255, 257, 262, 263 Gahmberg, C. G., 86, 88 Galardy, R., 280, 282 Galiano, R. D., 56 Galkina, E., 263 Galkina, E. V., 268 Galletti, G., 262 Gallez, B., 129, 144, 146 Galli, R., 54, 55, 302 Gambhir, S. S., 141, 147, 148, 149, 150, 151, 152, 154, 156, 158, 159, 160, 161, 162, 163, 166, 168 Gambliel, H. A., 195 Ganss, R., 55, 56, 57, 59, 64 Gao, L., 331 Garcia-Garrido, L., 210, 211 Garcia-Zepeda, E. A., 343 Garin, J., 182 Garmy-Susini, B., 38, 39, 44, 45, 343, 344, 352, 359, 361 Garofalo, A., 193 Garrett-Beal, L., 131 Garvik, B. M., 183 Gasparri, A., 87 Gatlin, C. L., 182 Gaudin, P. B., 86 Gaudry, M., 5 Gay, W. J., 234 Gaynutdinov, T. I., 156, 158 Gearing, A. J., 193

Author Index

Geary, T. W., 236, 237, 238 Gehling, U. M., 305 Geisendorf, S., 9, 10 Gelb, M. H., 181, 189 Gelfanova, V., 190 Gelovani, J. G., 84, 89, 90 Genant, H. K., 145 Genbacev, O. D., 281, 282, 288 George, V. G., 74 Gerber, H. P., 4, 127, 128, 129, 130, 132, 133, 134 Gerber, S. A., 181, 189 Gerecht-Nir, S., 304 Gerhardt, H., 55, 56 Geromanos, S. J., 191 Gerszten, R. E., 343 Gertsenstein, M., 126 Getsios, S., 282 Gevaert, K., 183, 189 Gheysens, O., 154 Ghosh, S., 265 Giacca, M., 302 Giannoudis, A., 343 Giavazzi, R., 193 Gilbert, L. B., 88, 89 Giles, W. B., 244 Gill, R., 331 Gillett, N. A., 127, 128 Gilley, D., 305, 306, 309, 311, 313, 314, 315 Gilman, A. G., 195 Gimbrone, M. A. Jr., 29, 126, 302, 343 Gingrich, D. E., 161 Ginhoux, F., 4, 10 Ginsberg, M. H., 270, 271 Ginsel, L. A., 264, 268 Ginzburg, Y., 282 Giordano, R. J., 85, 86, 88, 94, 95, 98 Giraudo, E., 85, 343 Girod, A., 88, 89 Gittenberger de-Groot, A., 210, 211, 212 Giudice, L., 288, 296 Glaser, M., 167 Glasser, L., 264, 268 Gleissner, C. A., 268 Gnatovskiy, L., 343 Godfrey, K. M., 248 Goebeler, M., 367 Goerdt, S., 193 Goertz, D. E., 144, 147 Goethals, M., 183 Gokoh, M., 6 Golding, M., 56 Goldstein, 292 Golos, T. G., 288 Golub, J. S., 304 Go´mez, M., 352 Gonzalez-Iriate, M., 211 Goodlad, R. A., 130, 131

381 Goodlett, D. R., 189 Goodman, S. L., 149 Goodwin, D. A., 163 Gorenstein, M. V., 191 Goresky, C. A., 234 Gorg, A., 182 Gormley, M., 288 Gorn, M., 193 Gorski, D. H., 129 Goshe, M. B., 189 Gossage, K. W., 49, 151 Goswami, S., 344 Gotsch, U., 8, 9 Gougerot-Pocidalo, M. A., 5 Gould, K. A., 130, 144, 147 Gould, M. N., 130 Gourdie, R., 210, 211 Graemmel, P., 263, 264 Graf, S., 157 Graf, T., 263, 264, 266 Gragoudas, E. S., 127 Grahl, K., 280 Gralla, O., 303 Gralow, J., 134 Grant, D. S., 28, 303, 318, 319 Grant, R. M., 296 Grauer, L. S., 86 Gray, R., 134 Grayburn, P. A., 154, 155 Grazul-Bilska, A. T., 230, 231, 232, 233, 234, 235, 236, 237, 238, 239, 241, 243, 244, 245, 246, 247, 248 Green-Jarvis, B., 54, 55 Gregoire, V., 129 Greinix, H., 4 Grieves, J., 194 Griffin, N. M., 177, 191, 192, 196, 197 Griffin, R. J., 129 Griffin, T. J., 189 Griffing, S., 134 Griffioen, A. W., 129, 152 Griffiths, C. E., 5 Grifman, M., 88, 89 Grimm, J., 144, 147 Grisaru-Granovsky, S., 282 Grisham, M. B., 4 Grobelny, D., 280, 282 Grobholz, R., 144, 145 Groenewegen, G., 167 Groger, M., 10 Groisman, A., 253, 270, 271, 272 Grompe, M., 54 Grone, E. F., 161 Grone, H. J., 161 Grosios, K., 5 Gross, J. F., 29 Grossin, M., 5 Grosso, F., 89

382

Author Index

Grummer, R., 297 Grunewald, M., 55, 56, 302 Grunstein, J., 133 Gruss, P., 9, 10 Grzesik, D. A., 343 Guelly, C., 54 Guha, A., 158 Guilford, W. H., 263 Gulati, R., 305 Guleserian, K. J., 304, 305 Guller, S., 292 Gullino, P. M., 28 Gulyas, G., 151 Guo, X., 88 Guo, Y., 28, 303, 318, 319 Gur, E., 148 Gurtner, G. C., 56 Gustafsdottir, S., 212 Gustafsson, E., 212 Gutierrez, E., 270, 271 Gutierrez, J., 128 Guyer, D. R., 127 Gyetko, M. R., 302 Gygi, S. P., 180, 181, 182, 189 H Haag, J. D., 130 Haas, S. J. P., 95 Habets, J. W., 152 Hackett, N. R., 55 Haedicke, W., 85, 86, 97, 98, 193 Hafez, S., 229 Hafezi-Moghadam, A., 268 Hagedorn, M., 98, 192 Hagedorn, N. L., 129 Hager, J. H., 132 Haider, M. A., 144, 146 Hainsworth, J., 134 Hajitou, A., 84, 86, 87, 88, 89, 90 Hakannson, L., 167 Halcox, J. P., 305 Hale, J. E., 190 Haley, L., 303 Halin, C., 1, 4, 6, 8, 9, 10, 12, 15, 262, 263 Hall, L., 128, 131, 133 Hallek, M., 88, 89 Hallmann, R., 9, 10 Hamano, S., 344 Hamberg, L. M., 5 Hamburger, V., 116, 213 Hamilton, G., 157 Hamilton, H. L., 116, 213 Hamilton, T., 129 Hamner, J. B., 129 Hamwi, A., 157 Han, Y., 256 Hanahan, D., 55, 67, 85, 132, 133, 143, 343

Haney, J. A., 28, 303, 318, 319 Hannay, J. A., 90 Hansen, H., 211 Hanson, E. T., 304 Hanzel, W., 9, 10 Harada, N., 193 Hardy, B., 95 Hari, D. M., 129 Harlow, E., 121 Harlow, L. A., 5 Haroon, H. A., 144, 146 Harpal, K., 126 Harrell, M. I., 14 Harris, A., 193 Harris, A. L., 3, 167, 343 Harris, J. W., 281 Harris, L. K., 293, 294 Harris, T. D., 150 Hartl, I., 89 Hartman, L., 280 Harvey, N., 8, 10 Hashizume, H., 8, 9, 144, 145 Haskard, D. O., 263 Hattori, K., 54, 55, 56, 342 Haubner, R., 88 Hauschild, A., 3 Haverich, A., 194 Havlik, E., 157 Hawighorst, T., 5 Hay, W. W. Jr., 233, 234 Hayakawa, K., 6 Hayashi, K., 38, 39, 40, 41, 44, 45, 46 Hayden, M. S., 296 He, L., 161, 162, 163, 166 He, Y., 343 Hebbel, R. P., 305, 306, 314 Hebestreit, H. F., 182 Heck, S., 263, 264, 266 Hedqvist, P., 260 Hegemeier, H. H., 193 Heikkila, P., 86, 88 Heil, M., 330 Heim, W., 134 Heinsohn, H., 126 Heissig, B., 54, 55, 56, 342 Helisch, A., 330, 339 Hellerqvist, C. G., 144, 145, 147 Hellstrom, M., 67, 212 Helmchen, F., 267 Hemberger, M., 223 Hemingway, I., 193 Hemmerich, S., 9, 10 Henderson, E., 144, 146 Hendricks, R. L., 28 Hendrikx, J., 55 Henke, E., 87 Herbst, R., 144, 147 Herlyn, M., 143

383

Author Index

Hermanowski-Vosatka, A., 195 Herscovitch, P., 144, 147 Hess, B. W., 236, 237, 238 Heston, W. D., 86 Heun, H., 129 Hickler, B., 245 Hicklin, D. J., 4, 6, 10, 15, 18, 54, 55, 132, 133, 154, 304, 342 Higashi, H., 343 Higgins, B., 128 Higgs, R. E., 190 Hildebrandt, S., 95 Hildenbrand, R., 367 Hill, J. M., 305 Hill, L. R., 190 Hill, S. A., 144, 145 Hillan, K. J., 126, 127 Himes, S. R., 264 Hiraga, A., 144, 145 Hirakawa, S., 3, 4, 6, 8, 10, 15 Hirakow, R., 210 Hiraoka, N., 302 Hirohashi, S., 190 Hiruma, T., 210 Hjarnaa, P. J., 322 Hochstrasser, D. E., 180 Hodgkiss, R. J., 144, 145 Hoffman, J. A., 85, 98 Hoffman, R., 29, 32 Hoffman, R. M., 27, 29, 30, 31, 32, 34, 36, 37, 38, 39, 40, 41, 42, 43, 44, 45, 46, 47, 48 Hofmann, J., 129 Hogers, B., 211 Ho Kim, T., 303 Holifield, J., 211 Hollema, H., 167 Holmgren, E., 134 Holmyard, D., 223 Honda, K., 190 Hong, H. Y., 88 Hong, Y. K., 2, 4, 6, 8, 10, 15 Hood, J. D., 149 Hood, L., 212 Hooper, A. T., 55 Hooper, N. M., 195 Hoorelbeke, B., 183 Horlin, A., 367 Hornig, C., 161 Horoszewicz, J. S., 193 Horvat, R., 9, 10 Hospers, G. A., 167 Hottori, K., 55 Hou, H., 338 Hou, L. C., 160 Houck, K. A., 126 Houston, M. L., 281 Hovorka, A., 4 Howarth, E. S., 233

Hoyt, C. C., 49, 151 Hsieh, S. L., 314 Hsu, A. R., 151, 153, 160 Hsu, Y. Y., 144, 146 Hu, G., 150, 151 Hu, H. M., 361 Hu, K., 30 Hu, L., 129 Hua, X., 131 Huang, J., 128, 129 Huang, J. C., 144, 146 Huang, K., 189 Huang, L., 193 Huang, Q., 30 Huang, S., 367 Huang, S. T., 182 Hudlicka, O., 234, 235 Hughes, B. J., 193 Hughes, C. C., 303 Hughes, M. S., 151 Hughes, W. F., 28 Hui, C., 212 Hui, K., 331 Hulliger, S., 267 Hume, D. A., 264 Huminiecki, L., 193 Hummerich, L., 6 Hung, H., 129 Hungerford, J., 211, 222 Hunkapiller, N. M., 279 Huo, Y., 265 Huppertz, B., 245, 288 Hur, J., 305 Hurtado, R., 211 Hurwitz, H., 134 Hutchins, 239 Hutchinson, O. C., 167 Hutson, R., 217 Huttary, N., 4 Hutter, J., 254, 260 Huysmans, H., 210, 212 Hwang, K. K., 305 Hylton, D., 6, 15, 127 Hyman, M. C., 265 Hynes, R. O., 149, 352 I Iachininoto, M. G., 56 Ighiguro, Y., 128 Ikawa, M., 32 Ikenasio, B. A., 245 Ikezaki, K., 343 Ilan, N., 8, 9 Ilic, D., 282 Illerhaus, G., 92 Ilmonen, M., 54 Ilyas, M., 130

384

Author Index

Inadera, H., 343 Inagaki, M., 215, 216 Inamura, T., 343 Ingber, D. E., 270 Ingram, D. A., 302, 303, 305, 306, 309, 311, 313, 314, 315 Inoue, M., 85, 132, 343 Intaglietta, M., 29 Ip, H., 222 Irimia, D., 270 Iritani, B. M., 14 Iruela-Arispe, M. L., 126 Ish-Hotowicz, D., 193 Ishibashi, T., 344 Ishida, A., 305 Ishihama, Y., 190 Ishii, Y., 211 Ishimori, T., 144, 145 Ismail, T., 86 Isner, J. M., 54, 303, 305, 342 Isobe, T., 190 Issa, B., 144, 146 Itin, A., 55, 56, 302 Ito, M., 4, 133 Ito, W. D., 194 Ito, Y., 343 Itoh, T., 55, 132, 133 Itohara, S., 55, 132, 133 Itskovitz-Eldor, J., 304 Iwai, S., 4, 133 Iwaki, T., 343 Iwasaka, S., 319 Iwata, C., 319 Izumo, S., 212 J Jaalouk, D. E., 95 Jablonka-Shariff, A., 235 248 Jackson, A., 144, 146, 167 Jackson, D. G., 3, 4, 5, 9, 10 Jackson, E. F., 144, 146 Jackson, J. R., 3, 4 Jacobson, B. S., 196 Jaffe, E. A., 194 Jaffe, R. B., 129 Ja¨ger, S., 95 Jaggi, J. S., 151 Jahagirdar, B., 305 Jahnke, A., 95 Jahnke, K., 3 Jain, N., 268 Jain, R. K., 4, 5, 14, 29, 37, 129, 144, 145, 268, 302, 303, 306 Jaitly, N., 190 Jakobsson, L., 18 James, K. M., 144, 147 James, P., 189

Janatpour, M. J., 281, 282, 291 Janson, N., 194 Janssens, C. J., 263 Janzer, R. C., 192 Ja¨rvinen, T. A., 85, 98 Jaskowiak, N. T., 129 Jauniaux, E., 289 Jayson, G. C., 144, 146, 167 Jehning, B. T., 155, 156, 158, 160, 161 Jeltsch, M., 4, 5, 56 Jenkins, N., 217 Jeon, N. L., 270 Jerin, J., 148, 158 Jessberger, R., 4, 10 Jevremovic, D., 305 Ji, Y., 85, 86, 98, 100 Jia, W. D., 95 Jia, Z., 126 Jiang, P., 30, 31, 32, 34, 37, 38, 40, 41, 45, 46, 48 Jiang, S., 54 Jiang, X., 270 Jiang, Y., 145 Jilani, S. M., 126 Jilkine, A., 189 Jin, D. K., 55, 56, 84 Jin, G., 354 Jin, H., 343, 344, 352, 359, 361 Jiwani, A., 144, 147 Joachimiak, A., 149 Jodele, S., 343 Johnson, D. H., 134 Johnson, L. A., 10 Johnson, M. L., 230, 231, 232, 233, 234, 235, 236, 237, 238, 239, 241, 243, 244, 245, 246 Johnson, R. S., 55, 56, 57, 59, 64 Jones, B., 331 Jones, C. J., 281, 293 Jones, M., 9, 10 Jonjic, N., 343 Jordan, B. F., 129 Joseph, J., 194 Josephson, L., 88 Josic, D., 195 Joukov, V., 5 Joyce, J. A., 85 Judenhofer, M. S., 148 Jugold, M., 150 Julyan, P., 167 Jung, S., 55, 56, 263, 264, 265, 302 Jung, S. M., 144, 146 Jung, U., 257, 260, 261, 265 Jurjus, A. R., 6 K Kagiyama, M., 5 Kaiho, S., 193 Kain, R., 85, 86, 87, 97, 98, 193

Author Index

Kaipainen, A., 5 Kajiya, K., 10 Kalen, M., 67, 212 Kalman, F., 210, 211 Kanazawa, S., 5 Kandel, J. J., 128 Kaneshima, H., 296 Kang, H. J., 305 Kano, M. R., 319 Kantor, C., 86, 88 Kao, L. C., 288 Kapidzic, M., 282, 288, 296 Kapiotis, S., 157 Kaplan, B., 95 Kapp, E. A., 186 Kappel, B. J., 151 Kapturczak, M., 88, 89 Karkkainen, M. J., 2, 8 Karlhofer, F., 4 Karpanen, T., 5, 282, 343 Kaserer, K., 157 Kashefi, A., 161, 162, 163 Kashgarian, M., 303 Kasirer-Freide, A., 270, 271 Kasman, I. M., 127, 130, 131 Kassner, A., 150 Kassner, P. D., 89 Katsuoka, K., 36, 37 Kattan, J., 211 Kaufman, D. S., 292, 304 Kaufmann, P., 230, 233, 234, 244, 245 Kaul, F., 88, 89 Kaul, S., 151 Kaur, P., 352 Kaushal, S., 38, 39, 44, 45, 305 Kawai, K., 157 Kawakami, M., 128 Kawamoto, S., 144, 145 Kawashima, A., 157 Kawate, T., 144, 145 Kawinski, E., 193 Kay, M. A., 88 Kayaselcuk, F., 343 Kayton, M. L., 128 Kazemi, M. R., 150 Keh-Yen, A., 4 Keller, A., 189 Kelly, K. A., 88 Kelly, L. M., 246, 263, 264, 266 Kempe, M., 267 Kempermann, G., 95 Kendziorski, C. M., 130 Kennel, S., 193 Kenner, L., 6 Kenny, L. C., 293, 294 Keogh, R. J., 293, 294 Kerbel, R. S., 55, 126, 129, 143, 196 Keren, S., 151

385 Kerjaschki, D., 4, 5, 9, 10 Kerr, D. J., 2, 86 Kersemans, V., 157 Keshet, E., 55, 56, 302 Kevil, C. G., 4 Keyt, B. A., 126, 161 Khafif, A., 148 Khan, N., 330, 338 Khan, Z. A., 304, 306, 314, 315, 318, 319, 320, 322 Khare, P. D., 89 Khoury, N. N., 6 Kieckens, L., 126 Kiefer, G., 151 Kieffer, S., 181, 182 Kienast, O., 157 Kiessling, F., 144, 145, 150 Kiessling, L. L., 282 Kikuchi, K., 5 Kilic, N., 305 Kim, E., 129, 144, 147 Kim, H. J., 4 Kim, H. S., 305 Kim, I., 4 Kim, I. S., 88 Kim, J., 128 Kim, J. M., 314 Kim, J. W., 314 Kim, K. J., 127 Kim, M. B., 257 Kim, R., 126 Kim, S., 160, 222 Kim, S. H., 4 Kim, S. Y., 314 Kim, Y. M., 314 Kimura, M., 128 Kimura, Y. N., 344 Kinahan, P. E., 148, 158 Kincaid-Smith, P., 244 King, B. F., 282 King, G. J., 244 King, G. L., 127 Kinoshita, M., 193 Kinuya, S., 157 Kinzler, K. W., 86, 130, 193, 194 Kirby, M., 212 Kircher, C. H., 3, 4 Kiselyov, A., 126, 127 Kishimoto, S., 6 Kislinger, T., 183, 189 Kitagaki, H., 6 Kitazawa, T., 6 Kitten, G., 211 Klagsbrun, M., 125 Kleeman, J., 343, 344, 352, 359, 361 Klein, R. J., 56 Kleinman, H. K., 319 Kleinman, M. E., 56

386 Kleinschmidt, J. A., 88, 89 Klibanov, A. L., 151 Kluge, K., 305 Kluger, M. S., 303 Klumpp, C., 151 Knierman, M. D., 190 Knighton, D. R., 28, 303 Knopp, M. V., 144, 146 Kobayashi, H., 144, 145 Koch, A. E., 3, 5 Kochman, R., 288, 296 Koenen, R. R., 343 Kofler, D. M., 89 Koh, G. Y., 4 Koh, T. S., 144, 146 Koh, Y. S., 4 Koike, M., 343 Koike, N., 303 Koivunen, E., 85, 86, 88, 98, 192 Kolker, E., 189 Koller, E., 157 Koller, F., 157 Kolonin, M. G., 84, 85, 86, 87, 91, 95, 98, 100, 193 Kominami, K., 32 Konerding, M. A., 144, 145 Koning, G. A., 152 Konishi, J., 144, 145 Konje, J. C., 233 Konstantinou, G. N., 144, 146 Konttinen, Y. T., 86, 88 Koodie, L., 305 Koong, A., 168 Koons, A., 129 Kopp, H. G., 55 Korchin, B., 90 Korey, M., 28 Korpanty, G., 154, 155 Korsisaari, N., 130, 131 Kostarelos, K., 151 Kothari, S., 305 Kovacs, G., 54 Kovats, S., 282, 291 Kowalski, H., 5, 9, 10 Kowalski, J., 128, 133 Kowanetz, M., 126, 131 Koziol, J. A., 191, 192, 197 Kragh, M., 322 Krajewski, S., 85, 86, 87, 97, 98, 193 Kramer, J., 212 Krasich, R., 302, 303, 305, 306, 313, 314 Krasinska, K. M., 179, 191, 193, 194, 196, 197 Kraus-Berthier, L., 322 Krebs, 247 Kreil, A., 157 Kreutzberg, G. W., 263, 264 Krieger, S., 4 Kriehuber, E., 9, 10

Author Index

Krier, T. R., 302, 303, 305, 306, 313, 314 Krikun, G., 292 Kristjansen, P. E., 322 Krober, S. M., 4 Krombach, F., 254, 260 Kros, J. M., 197 Krtolica, A. R., 282 Kubai, L., 28, 303, 318 Kubalak, S., 217 Kubes, P., 262, 267 Kuebler, W. M., 254, 256, 260, 262 Kufe, D. W., 129 Kuhn, H., 211 Kuhn, R., 6 Kuhnast, B., 88 Kuijpers, T. W., 264, 268 Kulesa, P. M., 113 Kulkarni, A. R., 56 Kumar, M., 222 Kumar, P., 190 Kunkel, E. J., 257, 260, 261 Kunkel, S. L., 5 Kuno, J., 144, 145 Kunstfeld, R., 4, 6, 10, 15 Kuo, T. K., 314 Kuppe, H., 262 Kuratsu, J., 343 Kurman, R. J., 282, 292 Kuroi, K., 343 Kurtaran, A., 157 Kuwabara, H., 190 Kuwano, M., 343, 344 Kuwano, T., 344 Kwaspen, F., 144, 145 Kwon, M. K., 88 Kwon, T. G., 88 Kwon, T. H., 88 Kwong, L. N., 130 Kyle, C., 9, 10 Kyriakides, T. R., 14 L Laakkonen, P., 4, 85, 98, 193 Labas, V., 194 Lacerda, L., 151 Lacy, E. K., 150, 151 Ladner, R. C., 89 Lahdenranta, J., 85, 86, 87, 88, 94, 95, 98 Lahm, H., 88 Lakhani, V. T., 131 Lakso, M., 5 Lal, A., 86, 193, 194 Lamagna, C., 53, 55 Lametschwandtner, A., 144, 145 Lamki, L., 144, 147 Lammers, T., 150 Lamond, A. I., 190

Author Index

Lanahan, A. A., 330, 331, 335 Landberg, J., 211 Landerholm, T., 211, 215, 216 Landers, R. J., 343 Landsman, L., 55, 56, 302 Lane, D., 121 Lane, W. J., 304 Lange-Asschenfeldt, B., 3, 4, 6, 10, 14, 15 Langer, R., 303, 304 Langley, R., 367 Lankester, K. J., 144, 146 Lanning, R. M., 306 Lansford, R., 113, 115 Lanza, G. M., 150, 151 Lanzen, J., 30 Lanzer, G., 54 Larocca, D., 89 Lashnits, E., 8, 9 Lasky, L. A., 9, 10 Laudanna, C., 258 Lavashova, Z., 155, 156, 158, 160, 161 Lavine, K., 211, 212 Lawitts, J. A., 14, 144, 145 Lawley, T. J., 74 Lawrence, M. B., 263, 268 Lawrenson, J. G., 54 Leach, M. O., 144, 146 Leapman, S. B., 29, 126 LeCouter, J., 4, 128, 133 LeDouarin, N., 214 Lee, A., 29 Lee, C. R., 55 Lee, C. V., 128, 133, 134 Lee, E. J., 88 Lee, H., 303 Lee, H. Y., 153 Lee, J. K., 268 Lee, K. D., 314 Lee, L., 158 Lee, M. H., 314 Lee, M. K., 89 Lee, M. M., 305 Lee, O. K., 314 Lee, P. S., 144, 146 Lee, R., 193 Lee, S., 211 Lee, S. H., 4, 126 Lee, S. M., 88 Lee, T. H., 4 Lee, T. Y., 144, 146 Lee, W., 127 Leedham, S., 130 Leek, R. D., 3, 343 Legallais, F. Y., 126 Leibovich, S. J., 5 Leike, K., 88, 89 Leips, J., 55 Lemay, J., 268

387 Lengauer, C., 86, 193, 194 Lennartsson, J., 303 Leong-Poi, H., 151 Le Ricousse-Roussanne, S., 322 Lerman, H., 148 Less, J. R., 144, 145 Leu, A. J., 37 Leung, D. W., 126 Leury, 246 Lev, D. C., 90 Levac, K., 304 Levashova, Z., 158 Levenberg, S., 304 Levenson, R. M., 49, 151 Levi, B. Z., 5 Levine, A. D., 6 Levine, J. P., 56 Levinsky, R. J., 264, 268 Lewis, C. E., 343 Lewis, J., 193 Lewis, J. S., 343 Lewis, R. L., 304, 318, 322 Ley, C. D., 322 Ley, K., 253, 254, 255, 256, 257, 258, 260, 261, 262, 263, 264, 265, 266, 268, 269, 270 Li, B., 6, 15, 56, 127, 133 Li, C. Y., 30, 88, 89 Li, F., 302, 303, 305, 306, 313, 314 Li, G. Z., 191 Li, H., 303, 338 Li, J., 305, 354 Li, J. F., 343 Li, K. C., 150 Li, K. L., 144, 146 Li, L., 30, 31, 32, 34, 36, 37, 46, 222, 304 Li, P., 5 Li, Q., 126 Li, S., 157 Li, T., 54, 305, 342 Li, W., 330, 331, 335 Li, X., 303 Li, X. M., 30, 31, 46 Li, Y., 179, 191, 192, 193, 194, 196, 197 Li, Z. B., 151, 152, 153, 159, 161, 162, 163, 168 Liang, W. C., 128, 133 Liang, X., 128, 133 Liang, X. H., 133, 134 Liao, W., 144, 145 Libermann, T., 8 Librach, C., 282 Librach, C. L., 280, 282, 291 Licha, K., 156 Lichy, M. P., 144, 145 Lieberman, J., 263, 264 Lieberman, M., 296 Liebherr, G., 211 Liegler, T. J., 296 Lie-Venema, H., 210

388 Lievshitz, G., 148 Lijowski, M., 151 Lilenbaum, R., 134 Lill, J., 189 Lilley, C. E., 89, 90 Lim, K. E., 144, 146 Lim, K. H., 291 Lim, Y. C., 343 Lima e Silva, R., 127 Lin, C., 4, 6, 10, 15 Lin, E. Y., 343 Lin, H., 190 Lin, K. K., 270 Lin, P., 30 Lin, P. C., 54, 55 Lin, Y., 305, 306, 314 Lindahl, P., 67, 212 Lindblom, P., 55 Lindbom, L., 254, 260, 261, 265 Lindenmayer, A. E., 144, 145 Lindgren, C., 194 Lindner, J. R., 151 Lindsey, J. R., 14 Link, A. J., 183 Linkesch, W., 54 Linquist-Stepps, V. D., 296 Lipson, K. E., 5 Lisanti, M. P., 195 Litovsky, S., 212 Little, C. D., 107, 108, 109, 110, 111, 113, 115, 116, 211 Littman, D. R., 263, 264, 265 Litton, M. J., 282, 283 Litvosky, S., 212 Liu, H., 189 Liu, H. L., 144, 146 Liu, J., 179, 195, 196 Liu, J. B., 144, 147 Liu, L., 262 Liu, P., 55, 56, 57, 59, 64 Liu, S., 88, 149, 162 Liu, T. T., 144, 146 Liu, W., 5 Liu, X., 215, 216 Liu, Y. K., 95 Liu, Z., 151, 166 Livak, K. J., 364 Llodra, J., 4, 10 Loboda, A., 189 Locke, R., 144, 145 Locksley, R. M., 282, 283 Lockwood, C., 292 Lockwood, C. J., 292 Lockwood, G., 144, 146 Lodish, M., 212 Logothetis, C. J., 86, 87 Lohela, M., 343 Lohler, J., 6

Author Index

Loiler, S. A., 88, 89 Lombardo, C. R., 87 Long, D., 244 Long, D. S., 256, 263, 269 Long, F., 191, 192, 196, 197, 212 Long, P. V., 28, 303, 318, 319 Longhi, R., 87 Longo, L. D., 244 Lorang, D., 131 Lorber, M. I., 303 Loren, M., 129 Loughna, S., 144, 145 Lowell, C. A., 260 Lowman, H. B., 128, 133 Lu, J., 211, 215, 216 Lu, K., 53 Lu, K. V., 55, 56, 57, 59, 64 Lu, L., 126 Lu, M., 222 Lu, N., 29 Lu, P., 190 Lu, Y., 223 Lubbert, M., 95 Lub-de Hooge, M. N., 167 Lubutti, S. K., 131 Luider, T. M., 197 Lukas, T. J., 191 Lumsden, A., 115, 116 Lunardi, J., 182 Lund, E. L., 322 Lundkvist, A., 56 Luo, W. W., 191 Luongo, C., 130 Luque, A., 352 Luscinskas, F. W., 343 Luster, A. D., 343 Luster, D., 343 Luther, J. S., 231, 234, 243, 246, 247, 248 Luthra, S. K., 167 Luttun, A., 56, 303 Lutty, G. A., 29 Lutz, A. M., 154 Lutzko, C., 343 Luyten, F. P., 319 Lyden, D., 54, 55, 305, 342 Lye, S., 293, 295 Lymboussaki, A., 37 Lyubynska, N., 4 M Ma, M., 88 Ma, P. X., 303 MacCalman, C. D., 282 Macias, D., 210, 211 Mack, C., 222 Madri, J. A., 8, 9, 192, 194 Madsen, C., 222

Author Index

Magness, R. R., 232, 234 Magni, F., 87 Magnusson, P., 18 Maguire, C. T., 209 Maguire, P. B., 183 Mahalingam, B., 367 Mahmood, U., 144, 147, 156 Mahoney, J. M., 28 Maierhofer, T., 54 Maiese, K. M., 128 Mailhos, C., 193 Mailloux, A., 194 Main, E. K., 282 Maini, R. N., 5 Maio, M., 167 Majesky, M. W., 209, 210, 211, 215, 216 Majima, T., 5 Makinen, T., 5 Makowski, E. L., 233 Malik, A. K., 128 Malischnik, C., 54 Mallick, P., 187 Malmquist, G., 190 Maltepe, E., 288, 296 Mammano, F., 296 Man, S., 55 Manabe, I., 222 Mandir, N., 131 Mandl, S., 158 Manjunath, N., 263, 264 Manka, D. R., 265 Mankoff, D. A., 148, 161 Manley, C., 129 Mann, J. S., 144, 146 Mann, M., 184, 190 Manner, J., 210, 211, 215 Manova, K., 87 Manseau, E. J., 14 Mansfield, J. R., 49, 151 Mansion, M., 182 Mantovani, A., 343 Mao, H., 191 Maoz, M., 282 March, K. L., 305 Marchio, S., 86, 88 Marcotte, E. M., 190 Mari, C., 158 Marimpietri, D., 87 Marini, F. C. III, 89, 90 Mariscalco, M. M., 193 Marker, P. H., 305 Markides, K. E., 190 Marks, J. D., 89 Marks, W., 54, 342 Marme, D., 5, 343 Marsh, J. N., 151 Martens, L., 183, 189 Martin, G. R., 28, 303, 318, 319

389 Martin, T., 304 Martina, Y., 89 Martinelli, G., 262 Martinive, P., 129 Maruta, F., 86 Marx, S. J., 131 Mass, R. D., 126 Masson, F., 181 Massoud, T. F., 147, 148, 149, 156 Masuda, Y., 304 Matli, M. R., 128 Matrisian, L. M., 54, 55 Matsuda, H., 128 Matsuda, K., 8 Matsushima, K., 343 Matsuzuki, H., 190 Mauad Filho, F., 233 Mauceri, H. J., 129 Maupin, A. B., 56 Mayer, D. N., 9, 10 Mayer, J. E., Jr., 304 Mayhew, T. M., 230, 234, 244, 245 Mayo, K. H., 129 Mayumi, T., 193 Mazal, P. R., 4 Mazar, A. P., 6 Mazo, I. B., 262 McCabe, N. P., 149 McCall, M. J., 163 McCarty, M., 367 McCarville, M. B., 129 McCowan, L. M., 292 McCune, J. M., 296 McDevitt, M. R., 151 McDonald, D. M., 4, 8, 9, 14, 85, 87, 143, 144, 145 McElroy, M., 27, 38, 39, 44, 45 McHugh, M., 127 McIntire, L. V., 268 McIntosh, D. P., 195, 196 McIntosh, M., 190 McLaughlin, B., 5 McLean, J. W., 144, 145 McMahon, G., 132, 133 McMahon, J., 55 McMaster, M., 282 McMaster, M. T., 282, 291 McMichael, A. J., 282 McNabola, A., 127 McNiel, E., 129 McRedmond, J. P., 183 McTigue, M., 163 Mead, L. E., 302, 303, 305, 306, 309, 311, 313, 314, 315 Meade, V., 305, 306, 309, 311, 313, 314, 315 Means, W. J., 236, 237, 238 Meares, C. F., 163 Medina, O. P., 86, 88

390 Medvedovska, J., 89 Meeus, S., 54 Mehic, D., 6 Meidell, R. S., 155 Meininger, C. J., 248 Melato, M., 343 Melder, R. J., 268 Melero-Martin, J. M., 301, 304, 306, 314, 315, 318, 319, 320, 322 Melton, D. A., 302 Mempel, T. R., 254, 260, 262 Mende, T., 305 Menendez, P., 304 Menetink, M., 211 Meng, C. K., 184 Meng, G., 128, 133 Meng, X., 5 Meng, Y. G., 128 Menicagli, M., 343 Mentink, M., 210, 211 Merad, M., 4, 10 Mercer, A., 5 Mercer, B., 222 Merler, E., 342 Merritt, C. R., 144, 147 Mertelsmann, R., 92 Mertens, M., 262 Meschia, G., 232, 233, 234 Mestas, J., 253 Metcalfe, J., 233 Meyer-Morse, N., 67, 132 Meza, J. E., 186 Miano, J., 222 Miao, H., 354 Michel, T., 248 Michelfelder, S., 89, 92 Middlesworth, W., 129 Miettinen, M., 144, 145 Mihm, M. C., 3 Mikawa, T., 210, 211 Miller, C. A., 186 Miller, J. H., 88 Miller, J. W., 28, 127 Miller, K., 134 Miller, R. K., 288 Miller, Y., 268 Milne, J. S., 245 Milosevic, M., 144, 146 Min, W., 5 Mineo, C., 195 Minick, C. R., 194 Mintz, P. J., 85, 86, 87, 95, 98 Miodonski, A. J., 144, 145 Miotto, G., 189 Misselwitz, B., 144, 145 Mitchell, C., 56 Miura, S., 262 Miyagawa, S., 86

Author Index

Miyazawa, K., 319 Miyazono, K., 319 Mize, G. J., 183 Modis, L., 211 Modlich, U., 193 Moglich, M., 144, 146 Mohamedali, K. A., 158, 159, 160, 161, 162, 163, 166 Mohle, R., 305 Molendini, C., 8, 9 Molina, R. D., 233 Molinaro, T. A., 296 Molkentin, J., 212 Molldrem, J. J., 84, 91 Monroe, M. E., 190 Monsky, W., 4, 14 Montani, J.-P., 234 Montet, X., 144, 147 Montgomery, E., 86, 193, 194 Moon, S. O., 4 Moons, L., 303 Moore, D. B., 306 Moore, J., 3 Moore, J. T., 129 Moore, L. G., 245 Moore, M. A., 54, 55, 304, 305 Moore, M. W., 126 Moosberger, I., 4 Moossa, A. R., 30, 31, 32, 34, 36, 37, 38, 39, 40, 41, 44, 45, 46 Mor, F., 2, 5 Mor, G., 292 Morabito, A., 2 Moran, A. M., 305 Morel, J. C., 5 Morgenstern, B., 150 Morikawa, S., 144, 145 Morishita, Y., 319 Morman, M. U., 267 Morris, D. R., 183 Morriss, F. H., 233 Morrissey, L. W., 194 Mortell, K., 305, 306, 309, 311, 313, 314, 315 Mosca, F., 343 Moser, A. R., 130 Moser, C., 254, 260 Mossman, H. W., 231, 232 Motoike, T., 144, 145 Moya, C., 95 Moya, C. A., 89, 90, 95 Mroueh, K. N., 302, 303, 305, 306, 313, 314 Mueller, M. M., 150 Mulatero, C., 167 Mulder, N. H., 167 Mulder, W. J., 152 Mullani, N., 144, 147 Mu¨ller, F., 92 Muller, O., 89

391

Author Index

Mu¨ller, O. J., 89 Mu¨ller, O. L., 88, 89 Muller, W., 6 Muller, Y. A., 133 Mullilken, J. B., 306, 315 Mumby, S. M., 195 Munchbach, M., 189 Munn, L. L., 268 Munoz-Chapuli, R., 210, 211 Murakami, M., 331 Murdoch, C., 343 Murohara, T., 54, 305, 342 Murphy, G. P., 193 Murphy, T. J., 14 Murray, C. D., 239 Murthy, S. K., 270 Muschel, R. J., 29 Mustafa, D. A., 197 Mustjoki, S., 5 Mustonen, T., 5 Muta, M., 343 Muthukkaruppan, V., 29 Muzihansky, A., 3 Muzyczka, N., 88, 89 N Nachman, R. L., 194 Nagasu, T., 190 Nagata, K., 215, 216 Nagengast, W. B., 167 Nagy, J. A., 14 Nagy-Bojarski, K., 4 Nahari, D., 5 Nahirney, P., 211 Naik, P., 132 Naiyer, A. J., 304 Nakagawa, S., 193 Nakahata, T., 4, 133 Nakajima, Y., 210 Nakamura, H., 210 Nakanishi, T., 32 Nakano, A., 267 Nakao, S., 344 Nakayama, J., 86 Nakayama, N., 166 Naldini, L., 54, 55, 302 Namikawa, R., 296 Nanda, A., 86 Nanus, D. M., 86, 88 Napolitano, M., 56 Narasimhan, K., 84 Narko, K., 5 Natali, P. G., 167 Nath, A. K., 303 Nazarian, L. N., 144, 147 Needham, J., 230 Nelson, P. J., 265 Nemerow, G. R., 88

Neufeld, G., 5 Neufert, C., 15 Neurath, M. F., 15 Nevin, L. M., 150 Newitt, R., 189 Newnham, 246 Ng, C. Y., 129 Ng, E. W., 127 Ng, Y., 213 Nguyen, A. V., 343 Nguyen, H. V., 126, 161 Nguyen, L., 3 Ni, J., 9, 10 Nice, E. C., 5 Nichlin, S. A., 89 Nicholay, K., 152 Nicklin, S. A., 88, 89 Nico, B., 87 Nicotra, M. R., 167 Niederleithner, H., 10 Nielsen, G., 29 Nikolova, G. V., 126 Nishi, T., 343 Nishie, M., 343 Nishii, R., 157 Nishimune, Y., 32 Nishimura, G., 128 Niu, G., 151 Niwa, H., 343 Niwa, Y., 343 Njardarson, J. T., 151 Nodasaka, Y., 8 Noel, A., 303 Norlin, J., 212 Norman, K. E., 260, 261, 267 Normanno, N., 2 Norrby, K., 303 North, R. A., 244 Norton, S., 190 Nourshargh, S., 258 Novelli, M., 130 Novikov, V., 144, 146 Novotny, M. V., 190 Novotny, W., 127, 134 Nowak, R. M., 244 Ntziachristos, V., 144, 147, 156 Nukiwa, T., 344 Nunes, D. N., 84, 89 Nursal, T. Z., 343 Nuseir, N., 5 Nutt, R., 148, 158 Nuyens, D., 213 Ny, A., 303 O Obermeyer, N., 194 Oceandy, D., 264 O’Connell-Rodwell, C., 158

392

Author Index

O’Connor, J. P., 144, 146 Oda, Y., 189, 190 O’Dushlaine, C., 189 O’Farrell, P. H., 181 Oh, B. H., 305 Oh, P., 179, 191, 192, 193, 194, 195, 196, 197 Ohizumi, I., 193 Oka, S., 6 Okabe, M., 32 Okazaki, K., 288, 296 Olafsen, T., 166 Olive, P. L., 144, 145 Oliver, G., 9, 10 Ollivier, V., 5 Olsen, M. W., 322 Olson, E., 222 Olson, L. M., 235 Olsson, C., 133, 134 Oltenfreiter, R., 157 Omenn, G. S., 186 Omigbodun, A., 282 Onderwater, J. J., 264, 268 O’Neill, E. E., 182 Ono, M., 190, 343 Ono, N., 6 Onoue, H., 343 Onuma, E., 5 Ordahl, C., 210, 211 Orkin, S., 212 Orlandi, A., 56 Ornitz, D., 211, 212 Orpana, A., 5 Orschell, C. M., 305 Orth, R. C., 144, 146 O’Shea, K. S., 126 Osman, S., 167 Ostoros, G., 54 Ostrand-Rosenberg, S., 55 Ostrowski, M. C., 264 Ostuni, E., 270 O’Toole, K. M., 128, 129 O’Toole, T., 149 Otsubo, M., 343 Otte, M., 305 Otto, F., 92 Ouchi, Y., 319 Oude Egbrink, M. G., 263 Oukrif, D., 130 Owens, G., 222 Oz, M. C., 304 Ozaki, Y., 343 Ozawa, M. G., 84, 85, 86, 87, 88, 89, 90, 95 Ozawa, T., 128 Ozerdem, U., 54 P Paavonen, K., 343 Padera, T. P., 14

Padhani, A. R., 144, 146 Page-McGaw, A., 55 Pagnan, G., 87 Pajusola, K., 343 Paku, S., 54 Pal, N., 130, 131 Paleolog, E. M., 4, 5 Palic, R. T., 330, 331, 335 Palmer, M. S., 282 Pan, N., 270 Panetta, C. J., 314 Pang, J. Z., 95 Pantarotto, D., 151 Pantel, K., 305 Papadakis, E. D., 89 Papenfuss, H. D., 29 Park, C. C., 5, 212 Park, E. C., 29 Park, H. D., 126 Park, J., 161 Park, R. W., 88 Park, S., 189 Park, S. Y., 314 Park, Y. B., 305 Parker, A. L., 86 Parker, G. J., 144, 146 Parkin, D. M., 142 Parmacek, M., 222 Parra, E., 245 Partanen, J., 211 Partlow, K. C., 151 Paruchuri, S., 304, 306, 314, 315, 318, 319, 320, 322 Pasqualini, R., 83, 84, 85, 86, 87, 88, 89, 90, 91, 94, 95, 97, 98, 100, 192, 193 Pasquier, C., 5 Passaniti, A., 28, 303, 318, 319 Passegue, E., 55, 56, 57, 59, 64 Pastorin, G., 151 Pastorino, F., 87 Patankar, T. F., 144, 146 Patel, M., 151 Patel, V., 155, 156, 158, 160, 161 Pattarini, L., 302 Patterson, S. D., 183 Patz, A., 29 Paulmurugan, R., 154 Pauly, R. R., 28, 303, 318, 319 Pavli, P., 264 Peale, F. V., 128, 130, 131, 133 Pearce, A. C., 182 Pearlman, J. D., 330, 331 Peault, B., 210, 212 Peck-Radosavljevic, M., 157 Peeters, L. L., 245 Peichev, M., 304 Pek, D. C., 88 Peng, J., 180

393

Author Index

Penman, S., 30, 31, 32, 34, 36, 46 Pennington, S. R., 180 Perabo, L., 89 Pereira, D., 304 Peremans, K., 156 Perens, E., 144, 145 Perez, E. A., 134 Perez-Pomares, J., 210, 211 Perk, J., 87 Perk, L. R., 167 Perkins, J., 248 Perrone, F., 2 Perry, E., 299 Perry, J., 282 Perry, M. C., 134 Perry, T., 305 Perry, T. E., 304 Pertovaara, L., 5 Pesce, M., 56 Peschke, P., 144, 145 Peterson, T. E., 305 Petit, I., 55, 56 Petkovic, L., 293, 295 Petrenko, V. A., 91 Petrich, B. G., 270, 271 Petritsch, C., 55, 56, 57, 59, 64 Petrova, T. V., 2, 3, 4 Petyuk, V. A., 190 Petzelbauer, P., 10 Peyman, G. A., 28 Pezzolo, A., 87 Pfarrer, C., 223 Pfeifer, D., 95 Phillips, H. S., 127 Picard, A., 304, 306, 314, 315, 318, 319, 320, 322 Piccardi, F., 87 Pichler, B. J., 148 Pickard, J., 257 Pickens, D. R., 144, 145, 147 Piecha, G., 131 Pieczek, A., 303 Pieper, R., 182 Pierce, E. A., 127 Pierce, G. F., 89 Piersanti, S., 89 Pigott, R., 193 Pilch, J., 85, 98, 193 Pili, R., 28, 303, 318, 319 Pinto, J. T., 86 Pirich, C., 157 Pisani, P., 142 Piston, D. W., 267, 268 Pitot, H. C., 130 Pixley, F. J., 344 Planelles, V., 5 Pober, J. S., 3, 13, 303 Poelmann, R., 210, 211, 212

Poinas, A., 182 Polakis, P., 130 Politi, L. S., 54, 55, 302 Pollard, J. W., 55, 264, 342, 343 Pollefeyt, S., 126 Pollock, R. E., 90 Pollok, K., 305, 306, 309, 311, 313, 314, 315 Polverini, P. J., 5 Ponomarev, V., 87 Ponten, A., 303 Poon, R. T., 126 Porcelli, D., 56 Porkka, K., 85, 98, 193 Pospieszalska, M. K., 253 Poston, L., 244, 245 Potoczek, M., 144, 147 Poul, M. A., 89 Poulsom, R., 130, 131, 193 Powell-Braxton, L., 126 Prakobphol, A., 282, 288 Prater, D. N., 302, 303, 305, 306, 313, 314 Prato, M., 151 Prchal, J. T., 302, 303, 305, 306, 313, 314 Preda, A., 144, 146 Preece, G., 263 Preisinger, A. C., 130 Preitfellner, J., 157 Preston, S., 130 Prevo, R., 3, 10 Price, P., 167 Price, R., 144, 146 Pries, A. R., 255, 256, 262 Prise, V. E., 144, 145 Proudfoot, A. E., 265 Purvine, S., 189 Putzer, B. M., 95 Puype, M., 183 Puzon, W., 352 Pyle, A. L., 56 Pyrah, I. T., 131 Pytowski, B., 4 Q Qi, J., 148 Qian, H., 343 Qian, Y. B., 95 Qin, J., 149 Qin, L. X., 95 Quadroni, M., 189 Quake, S. R., 272 Queenan, J. T., Jr., 288 Quemeneur, L., 4, 10 Quensel, C., 212 Quigley, J. P., 149 Quintana, F. J., 2, 5 Qumsiyeh, M., 292 Quyyumi, A. A., 305

394

Author Index

R Raab, I., 4 Rabbany, S. Y., 56 Rabilloud, T., 181, 182 Rabkin, E., 305 Rabkin-Aikawa, E., 304 Racek, T., 95 Radivojac, P., 190 Radziejewski, C., 5 Raff, M. C., 192 Raffler, N. A., 268, 269, 270 Rafii, D., 54 Rafii, S., 54, 55, 56, 84, 304, 305 Rago, C., 86, 193, 194 Rahman, M., 292 Rainisalo, A., 86, 88 Raither, A., 95 Rajantie, I., 54, 343 Rajasekeran, S., 129 Rajewsky, K., 6 Rajotte, D., 85, 86, 97, 98, 192, 193 Raju, K., 28 Rambaldi, A., 193 Ramirez, J., 189 Ramirez, M. I., 10 Ramos, C. L., 265 Ramsey, E. M., 231, 232, 244, 281 Ran, S., 84 Randolph, G. J., 4, 10 Rangel, R., 89, 90 Ranson, M., 167 Rao, J., 149, 150 Rao, R., 87 Rappsilber, J., 190 Rasid, A., 126 Ratajska, A., 211, 212 Raud, J., 257, 260 Rauvala, H., 5 Raychaudhuri, A., 5 Raychaudhuri, S. K., 6 Raychaudhuri, S. P., 6 Raymond, E., 127 Razorenova, O., 149 Razvi, S., 303 Reardan, D. T., 163 Reddi, A. H., 319 Red-Horse, K., 126, 129, 282, 288, 296 Redline, R., 6 Redman, C. W., 282 Redmer, D. A., 229, 230, 231, 232, 233, 234, 235, 236, 237, 238, 239, 241, 243, 244, 245, 246, 247, 248 Regan, C., 222 Regele, H., 4 Regele, H. M., 4 Regnault, 246 Rehman, J., 305

Reich, R., 282 Reichelderfer, M., 130 Reilly, J. P., 190 Reilly, R. M., 158 Reimund, J. M., 6 Reneman, R. S., 263 Rennick, D., 6 Renz, M., 9, 10 Resing, K. A., 186 Restel, B. H., 89, 90 Reusch, P., 5 Reuter, V. E., 86 Reyes, M., 305 Reynolds, L. P., 229, 230, 231, 232, 233, 234, 235, 236, 237, 238, 239, 241, 243, 244, 245, 246, 247, 248 Reynolds, P., 89 Reynoso, J., 32, 36 Ribatti, D., 87 Rice, A., 5 Richard, L., 4, 14 Richardson, C. D., 144, 145 Richardson, K., 191 Ricordi, C., 88, 89 Riddle, L., 127 Ried, M., 88 Ried, M. U., 89 Riefke, B., 156 Rietveld, F. J., 144, 145 Riggins, G. J., 86, 193, 194 Rio, G. D., 87 Risau, W., 8, 9, 126, 144, 145, 302 Rist, B., 181, 189 Ristimaki, A., 5 Rivera, J., 288, 296 Rizzardi, C., 343 Ro, R. J., 144, 147 Roberge, S., 144, 145 Roberts, A. B., 319 Roberts, C., 144, 146 Roberts, T. P., 144, 146 Robertson, D. A., 233, 234 Robertson, H. A., 244 Robertson, J. D., 150 Robertson, W. B., 289 Roche, N. S., 319 Roche, S., 214 Rochon, Y., 182 Rockl, W., 161 Roddy, R., 148, 158 Rodrigo Mora, J., 262, 263 Rodriguez, W., 270 Rodriguez-Porcel, M., 154 Rohde, E., 54 Rohren, E. M., 156 Rolny, C., 18 Roman, B. L., 144, 145 Romano, T., 8, 9

395

Author Index

Romans, K. E., 86, 193, 194 Romero, R., 292 Rong, H., 127 Rongish, B. J., 108, 109, 110, 111, 113, 115 Roos, D., 264, 268 Rosales, A. G., 89 Rosen, N., 87 Rosen, S. D., 9, 10, 282 Rosenberg, R. D., 192 Rosenblum, M. G., 158, 159, 160, 161, 162, 163, 166 Rosenfeld, C. R., 233 Rosenfield, K., 303 Rosengren-Beezley, S., 260 Rosenmaier, A., 4 Rosenquist, T., 212 Rosenzweig, A., 343 Ross, J., 131 Rossant, J., 293, 295, 342 Rossignol, M., 182 Roth, I., 282, 283 Roth, M. J., 89 Rothberg, K. G., 195 Rothlein, R., 193 Rouleau, A., 304 Rouquie, D., 182 Rowe, D. H., 128 Rowland, A. M., 126 Rowlands, D. C., 86 Roy, S., 190 Rubart, M., 268 Rubenstein, J. L., 128 Ruddell, A., 14 Rudge, J. S., 6, 15, 127 Rudloff, H. E., 193 Ruel, J., 268 Ruggeri, B., 161 Ruhrberg, C., 56 Ruiter, D. J., 144, 145 Ruiz de Almodovar, C., 330, 331, 335 Ruoslahti, E., 84, 85, 86, 87, 88, 97, 98, 192, 193 Rupp, P. A., 107, 108, 109, 110, 111, 113, 116 Russell, R. G., 343 Russell, S. J., 89 Rustin, G. J., 144, 146 Ryan, A. J., 131 Ryder, U., 190 Rygaard, J., 322 Ryu, H. M., 314 S Sacchi, A., 87 Sacchi, N., 216 Sachs, J. R., 190 Sackett, T. J., 331 Sadygov, R. G., 189 Saeed, M., 144, 146

Saffari, N., 144, 147 Saga, T., 144, 145 Saggio, I., 89 Saha, P. K., 85, 87, 98, 193 Sahai, E., 344 Saharinen, P., 2, 8 Saijo, Y., 344 Saishin, Y., 127 Saji, H., 343 Sakamoto, M., 85, 86, 98 Saksela, O., 5 Sakurai, Y., 4, 133 Salloum, R. M., 129 Salo, T., 86, 88 Salone, B., 89 Salven, P., 54, 343 Sampaolesi, M., 54, 55, 302 Sa´nchez-Madrid, F., 352 Sanders, L. C., 149 Sandhu, J., 158 Sandler, A., 134 Sandra, A., 211 Sanford, J. S., 270, 271 Sano, D., 128 Sans, M., 6 Santoni, V., 181, 182 Sanyal, M., 6 Sapi, E., 292 Sarelius, I. H., 257 Sarembock, I. J., 265 Sargent, I. L., 282 Sargent, W., 127 Sargiacomo, M., 195 Sarkis, A. S., 86, 87 Sasmono, R. T., 264 Sasso, F., 343 Sato, N., 144, 145 Sato, T., 190 Sato, T. N., 144, 145 Sauvage, L. R., 305 Sawa, Y., 8 Sawamura, T., 89 Sawano, A., 4, 133 Scacheri, P. C., 131 Scadden, D. T., 56, 306 Schacht, V., 4, 6, 8, 10, 15 Schafer, B., 305 Schaff, U. Y., 270 Schafhausen, P., 305 Schainfeld, R., 303 Schanz, A., 288, 296 Schanz, S., 128 Schaper, W., 330, 339 Scharffetter-Kochanek, K., 260 Schatteman, G., 54, 305, 342 Schatz, F., 292 Scheaffer, 246 Schechner, J. S., 303

396 Scheibe, J., 95 Scheinberg, D. A., 151 Schenke, W. H., 305 Scher, H. I., 128 Scherer, P. E., 195 Scheuch, H., 6 Schieltz, D. M., 183, 189 Schiffman, S. R., 151 Schiller, J. H., 134 Schima, W., 157 Schinke, M., 212 Schipper, M., 151 Schirner, M., 5 Schlaeger, T. M., 144, 145 Schlingemann, R. O., 86, 88, 144, 145 Schluter, J., 211, 215 Schmalstieg, F. C., 193 Schmid, M., 157 Schmid, M. C., 341 Schmid-Schonbein, G. W., 254 Schmidt, A., 95 Schmieder, A. H., 150 Schmittgen, T. D., 364 Schneebaum, S., 148 Schneider, R. M., 89 Schnitzer, J. E., 177, 179, 191, 192, 193, 194, 195, 196, 197 Schnurch, H., 8, 9 Schober, A., 265 Schoen, F. J., 304, 305 Schoenwolf, G. C., 115, 116 Schon, M. P., 4, 14 Schroeder, D. A., 54 Schuch, G., 305 Schulte, I., 211, 215 Schulze-Osthoff, K., 193 Schumacher, U., 305 Schutz, F., 186 Schwartz, A., 148 Schwartz, H. M., 338 Schwartz, R., 211, 215 Schwarz, E. M., 5 Schwickert, H. C., 144, 146 Scollard, D. A., 158 Scott, D. L., 149 Scott, J. K., 84, 90 Scott, M. J., 144, 146, 150, 151 Seandel, M., 55 Seed, B., 29 Seed, M. P., 3, 4 Seetharam, S., 129 Sekine, K., 270 Selig, M., 29 Semmler, W., 150, 156 Senekowitsch-Schmidtke, R., 88 Sergeeva, A., 84, 91 Sergi, L. S., 54, 55, 302 Sessa, W. C., 3, 13

Author Index

Sevick, E. M., 144, 145 Seymour, L. W., 86 Shaffrey, M. E., 151 Shah, V., 305 Shaked, Y., 55 Shaler, T. A., 190 Shames, D. M., 144, 146 Shan, S., 30 Shanker, P., 263, 264 Shapira, O. M., 305 Shapiro, L. H., 85, 86, 98 Sharma, S., 194 Sharpe, E. E., 56 Shashkin, P., 268 Shaw, D. E., 6 Shen, R. F., 190 Shen, W., 331 Shen, Y., 88 Shenkier, T., 134 Shepard, H. M., 5 Shepherd, B. R., 303 Sher, A., 263, 264 Sherman, D., 128 Shi, Q., 305 Shi, W. F., 89 Shibuya, M., 4, 126, 133 Shido, K., 54, 55 Shields, D. C., 189 Shientag, L., 29 Shih, I. M., 282, 292 Shima, D., 56 Shima, D. T., 127 Shimada, H., 343 Shin, D. W., 151, 152 Shin, J. W., 2 Shinde-Patil, V. R., 88 Shinners, B. L., 318, 322 Shiohara, T., 6 Shitara, K., 4, 133 Shitashige, M., 190 Shively, V., 5 Shmelkov, S. V., 54, 55 Shoenfeld, Y., 5 Shohet, R. V., 154, 155 Shojaei, F., 126, 128, 129, 304 Shono, T., 343 Shore, S., 191, 192, 196, 197 Shriver, M., 245 Shukla, D., 4 Shultz, L. D., 296 Shuman, M. A., 128 Shyr, Y., 54, 55 Sica, A., 343 Siddique, T., 191 Sidky, Y. A., 194 Sidman, R. L., 84, 88, 89 Siegal, G. P., 87 Sierra-Honigmann, M. R., 303

397

Author Index

Silberhumer, G., 4 Silva, J. C., 191 Silver, M., 54, 305, 342 Sim, D., 29 Simari, R. D., 305 Simmons, W. W., 192 Simon, M., 161 Simon, S. I., 270 Simons, M., 329, 330, 331, 332, 333 Simper, D., 314 Simpson, D. H., 144, 147 Simpson, R. J., 186 Sims, T. L., 129 Sin, A., 270 Singbartl, K., 263, 264, 266 Singer, M. S., 9, 10, 282 Singh, H., 282 Singh, M., 85 Singh, R., 151 Sinha, P., 55 Sinigaglia, M., 302 Sipkins, D. A., 150 Sitartchouk, I., 144, 146 Sjogren, C., 211 Skalak, R., 256, 257, 258, 268 Skalak, T. C., 144, 145, 254, 256 Sklenar, J., 151 Skobe, M., 4, 5, 10 Skornick, Y., 148 Slaaf, D. W., 263 Slegers, G., 157 Smart, E. J., 195 Smith, B. R., 263 Smith, C., 211, 212 Smith, C. W., 193 Smith, D. F., 265, 268, 269, 270 Smith, G. P., 84, 90, 91 Smith, L. E., 127 Smith, M. L., 256, 263, 268, 269, 270 Smith, N., 167 Smith, N. R., 131 Smith, R. D., 189, 190 Smith, T. W., 192 Smits, R., 130, 131 Smitt, P. A., 197 Snyder, E. Y., 88 So, P. T., 14 Soffer, S. Z., 129 Soghomonyan, S., 89, 90 Soker, S., 305 Soleiman, A., 4, 9, 10 Solit, D. B., 87 Solovey, A., 305, 306, 314 Somia, N., 88 Song, H., 55, 56, 57, 59, 64 Song, S., 54, 57, 67, 68, 73, 88, 89, 132 Sonoda, K. H., 344 Sorg, C., 193, 367

Soriano, P., 212 Sorsa, T., 86, 88 Sosa-Pineda, B., 9, 10 Souza, G. R., 88, 95 Sozzani, S., 343 Spana, E. P., 9, 10 Spara, P., 233 Speece, P., 88, 89 Spencer, D. M., 6 Spencer, S. S., 156 Spencer, T. E., 231, 234, 243, 247, 248 Sperandio, M., 257, 268 Stacker, S. A., 4, 5 Staelens, L., 157 Staes, A., 183, 189 Stainier, D. Y., 144, 145 Stalboerger, P. G., 314 Stallcup, W., 54, 57, 67, 68, 73 Stallcup, W. B., 86 Standing, K. G., 189 Staquicini, F. I., 88, 90 Starkey, P. M., 282 St Croix, B., 86, 193, 194 Steen, H., 184 Stehle, T., 149 Steinbrink, K., 367 Steiner, S., 182 Stetler-Stevenson, W. G., 149, 302 Stewart, C. F., 129 Stewart, P. A., 192 Stinson, W. G., 28 Stirling, J. J., 144, 146 Stock, M. K., 233 Stockton, B., 263, 264 Stoddart, C. A., 296 Stolar, C. J., 128, 129 Stoll, B. R., 14 Stone, A., 144, 146 Stone, P. R., 292 Storb, R. F., 305 Storm, G., 152 Straino, S., 56 Straub, J., 130 Strauss, J. F. III, 282, 288 Strauss, L. G., 144, 147 Streilein, J. W., 5 Streiter, R. M., 344 Stride, E., 144, 147 Strieter, R. M., 5 Strijkers, G. J., 152 Strittmatter, E. F., 190 Stromblad, S., 149 Strunk, D., 54 Stryhn, A., 85, 86, 98 Stubblebine, M., 282 Stulting, R. D., 28 Stupack, D., 343, 344, 352, 359, 361 Stutte, H. J., 367

398

Author Index

Su, J., 9, 10, 343, 344, 352, 359, 361 Su, L. K., 130 Su, Q., 182 Suarez, Y., 303 Suchting, S., 193 Sugiura, T., 6, 29 Sullivan, A., 54, 305, 342 Sullivan, M. J., 191 Sumen, C., 262, 263 Sumi, H., 343 Summers, S., 74 Sun, F. X., 30, 31, 46 Sun, H. C., 95 Sun, J., 85, 86, 95, 98, 100 Sun, L., 88 Sun, S., 153 Sun, X., 166 Sun, Y., 89, 90 Sundararajan, S., 92 Sundberg, C., 14 Sundd, P., 253 Sunderkotter, C., 367 Sunshine, M. J., 263, 264 Sutherland, F. W., 304, 305 Suzuki, K., 343 Svoboda, K., 267 Swartz, H. M., 330, 338 Swartz, M., 5 Swerlick, R. A., 74 Symes, J. F., 303 Szabo, S., 28 Szekanecz, Z., 3 T Tabata, T., 190 Tabuchi, A., 262 Tafuro, S., 302 Tagashi, K., 144, 145 Tahara, K., 343 Takada, Y., 352 Takagi, H., 127 Takahashi, K., 127, 343 Takayama, S., 270 Takemoto, M., 212 Takeya, M., 343 Taksir, T., 29 Tallquist, M., 212 Tamaki, K., 55, 132, 133 Tammela, T., 2, 3, 4, 8 Tammi, R., 9, 10 Tan, X., 14 Tanaka, H., 305, 306, 309, 311, 313, 314, 315 Tanaka, M., 212 Tang, H., 190 Tang, K., 190 Tang, L., 127 Tang, Q., 88, 89

Tang, S., 331 Tang, Z., 195 Tang, Z. Y., 95 Tangelder, G. J., 263 Taniguchi, K., 193 Tanzawa, K., 55, 132, 133 Tarpey, J., 280 Taylor, D. J., 233 Taylor, J. N., 144, 146 Taylor, P. C., 5 Taylor, R. M., 28, 303, 318, 319 Tedder, T. F., 257 Tedjarati, S., 367 Teichmann, G., 144, 145 Teillet, M., 214 Tejada, M., 133 Tejada, M. L., 128, 133 Tejada, R., 54, 55 Teleron, A. A., 56 Tellides, G., 303 Temm, C. J., 302, 303, 305, 306, 313, 314 Tenenbaum, A., 95 Tepper, O. M., 56 Ter-Pogossian, M. M., 144, 147 Testa, J. E., 179, 191, 193, 194, 196, 197 Tevosian, S., 212 Thacker, N. A., 144, 146 Thaler, D., 54 Thatte, J., 263 Theilmeier, G., 213 Theis, H., 161 Theodore, W. H., 156 Thirsk, G., 144, 145, 147 Thirumamagal, B. T., 156 Thng, C. H., 144, 146 Thomas, C. E., 88 Thomas, G. R., 183 Thomas, K. L., 268 Thompson, R., 128 Thomson, J. A., 304 Thoren, P., 265 Thorlacius, H., 257, 260 Thornburg, K. L., 244 Thorpe, P., 55, 132, 133 Thorpe, P. E., 84 Thum, T., 194 Thurberg, B., 212 Thurman, G. B., 144, 145, 147 Thurston, G., 14, 144, 145 Tice, D. A., 162, 166, 168 Timar, J., 54 Tinkle, C. L., 55 Tirziu, D., 330, 331 Titulaer, M. K., 197 Tjarks, W., 156 Tjwa, M., 303 Tkachenko, E. V., 330, 331, 335 Tkachenko, S. E., 126, 127

399

Author Index

Tobelem, G., 322 Tobler, N. E., 4, 6, 8, 9, 10, 12, 15 Toi, M., 343 Tolentino, M. J., 127 Tomanek, R., 210, 211, 213, 330, 331, 335 Tomita, K., 38, 40, 41, 45, 46 Tomlinson, I. P., 130 Tompkins, R. G., 270 Toner, M., 270 Tong, B., 189 Tong, W., 264 Tong, W. P., 86 Tool, A. T., 264, 268 Toomey, S., 183 Torella, A. R., 56 Torrealba, J., 130 Totsuka, Y., 8 Tovari, J., 54 Townend, K., 244 Townsend, D. W., 148, 158 Tozawa, K., 29 Tozer, G. M., 144, 145 Tran, H., 144, 147 Traub-Weidinger, T., 157 Traverso, G., 86, 193, 194 Treese, F. A., 29 Trepel, M., 83, 84, 85, 86, 87, 88, 89, 90, 92, 95, 98 Troncoso, P., 86, 87 Trotter, M. J., 144, 145 Troxel, A., 128 Trudinger, B. J., 244 Truffa, S., 56 Tschachler, E., 6, 9, 10 Tse, V., 160 Tseng, J. R., 168 Tsourkas, A., 88 Tsuchiya, H., 38, 40, 41, 45, 46 Tsuchiya, M., 262 Tsukuda, M., 128 Tsunoda, T., 5 Tsutsui, S., 343 Tsutsumi, Y., 193 Tsutsumi-Miyahara, C., 344 Tu, M., 194 Tu, Y. H., 195 Tuma, R. F., 254, 261 Turecek, F., 181, 189 Turetschek, K., 144, 146 Turkington, T. G., 156 Turner, A. J., 195 Turner, M. A., 288 Tyrrell, J. A., 306 U Ueda, S., 344 Ueno, T., 343

Ulaszek, R. R., 189 Ulloa-Aguirre, A., 288 Underiner, T. L., 161 Underwood, J. C., 343 Unger, R. H., 155 Unnikrishnan, S., 257 Unutmaz, D., 263, 264 Urba, W. J., 361 Usami, S., 256, 268, 354 Ushio, Y., 343 Utsunomiya, H., 303 Uutela, M., 343 Uziely, B., 282 V Vacanti, J. P., 303 Valdembri, D., 86, 88 Valent, P., 157 Valkovic, T., 343 Valtanen, H., 86, 88 Van Arsdall, M., 367 Van Bruggen, N., 131 Van Damme, J., 183, 189 Vandekerckhove, J., 183, 189 van de Kerkhof, P. C., 144, 145 van den Akker, N., 210 Vandenberg, S., 55, 56, 57, 59, 64 Van Den Bossche, B., 156 Vandenhoeck, A., 126 Vanderheyden, J. L., 158 van der Schaft, D. W., 152 van der Schoot, C. E., 264, 268 van der Zee, R., 54, 305, 342 Van de Wiele, C., 156 van Dijke, C. F., 144, 146 Vandlen, R., 126 van Dongen, G. A., 167 van Trappen, P., 3 Varas, F., 263, 264, 266 Vargas, E., 245 Varner, J. A., 38, 39, 44, 45, 341, 343, 344, 352, 359, 361 Vasile, E., 14 Vaubourdolle, M., 194 Vaughey, J. N., 126 Veelken, H., 92 Veeravagu, A., 160 Veikkola, T., 5 Velasco, P., 3, 4, 6, 10, 14, 15 Velculescu, V., 86, 193, 194 Venneri, M. A., 54, 55, 302 Verfaillie, C. M., 305 Verhoeven, A. J., 264, 268 Verma, I. M., 88 Vestweber, D., 8, 9, 257, 261 Vickers, M. H., 245 Vicovac, L., 281

400

Author Index

Vidal, C. I., 85, 86, 98, 100 Vider, J., 87 Vidugiriene, J., 195 Vieira Santana Netto, P., 233 Vigl, B., 4, 6, 8, 9, 10, 12, 15 Vile, R. G., 305 Vincent, P., 127 Vinh, J., 194 Vink, H., 256 Viragh, S., 210, 211 Virgolini, L., 157 Visintin, I., 292 Vlodavsky, I., 282 Vo, K. C., 288, 296 Vogel, C., 190 Vogel, H., 160 Vogelstein, B., 86, 130, 193, 194 Vogtle, F. N., 92, 95 Vojnovic, B., 144, 145 Vollmer, E., 193 von Andrian, U. H., 14, 262, 263, 264, 343 von Haller, P., 189 von Hundelshausen, P., 265 von Kriegsheim, A. F., 182 Vonnahme, K. A., 230, 231, 232, 233, 234, 236, 237, 238, 239, 243, 244, 245, 247, 248 von Schalscha, T. L., 149 Von Seggern, D. J., 88, 89 Vora, A., 3 Vrancken-Peeters, M., 210, 211 Vu, T. H., 55, 132, 133 Vukicevic, S., 319 W Waclawiw, M. A., 305 Wagenblast, J., 144, 146 Wagener, C., 305 Wagle, A., 331 Wagner, E. F., 6 Wagner, R., 254 Wagner, S., 330, 339 Wagstaff, J., 167 Wainwright, B. J., 264 Wait, R., 182 Waku, K., 6 Waldo, K., 212 Walker, W. F., 263 Walker-Samuel, S., 144, 146 Wall, D., 191 Wallace, J. M., 230, 231, 232, 233, 234, 236, 239, 243, 244, 245, 246, 247, 248 Waller, J. L., 130 Walsh, K., 303 Wamhoff, B., 222 Wang, F., 303 Wang, G., 190, 331 Wang, H., 159, 161, 162, 163, 168

Wang, J. W., 30, 31, 46, 158 Wang, L., 95, 126, 304 Wang, M., 134 Wang, P., 190 Wang, R., 190 Wang, S., 314 Wang, S. X., 9, 10 Wang, W., 190 Wang, X., 30, 31, 46, 88 Wang, Y., 88, 354 Wang, Y. P., 196 Wang, Z., 155 Ward, 247 Ward, J. M., 131 Ward, N., 211 Wareing, M., 293, 294 Warren, R. S., 128 Warrington, K. H., 88, 89 Washburn, M. P., 181, 183, 189 Watabe, T., 319 Watanabe, K., 6 Waterbury, L. D., 28 Waterkamp, D. A., 89 Waterton, J. C., 144, 146 Watkins, A. J., 131 Watkins, D. N., 86 Watschinger, B., 4 Watson, A. L., 289 Watson, S. P., 182 Watt, S., 29 Watt, S. L., 194 Watts, J. D., 189 Wayland, H., 256 Wayner, E. A., 352 Weber, C., 265, 343 Weber, K. S., 265 Webster, S. J., 343 Wedge, S. R., 131 Wei, J., 215, 216 Wei, X., 4, 6, 10, 15 Weich, H. A., 157, 161, 343 Weichert, J., 130 Weigmann, B., 15 Weiler-Guettler, H., 192 Weinbaum, S., 256 Weinstein, B. M., 144, 145 Weisdorf, D. J., 305, 306, 314 Weiss, C., 95 Weiss, J. B., 5 Weiss, S. J., 302 Weiss, W., 182 Weissleder, R., 88, 144, 147, 156 Weissman, I. L., 296 Weitzman, M. D., 88, 89 Wells, J. A., 133 Wells, S. A., 131 Welson, S. Y., 182 Wen, L. P., 88

401

Author Index

Wendtner, C. M., 89 Weninger, W., 9, 10, 14 Werb, Z., 54, 55, 56, 57, 59, 64, 67, 68, 73, 132, 133, 280, 282, 344 Werr, J., 265 Wesseling, P., 86, 88 West, 239 Westbrook, J. A., 182 Wester, H. J., 88 Westmore, M., 330, 331, 335 Westphal, M., 128 Wethmar, K., 257, 263 Wheelock, M., 281, 282 White, A., 211, 212 White, D. L., 145 White, E. J., 268, 269, 270 White, S., 222 White, S. J., 89 Whitehouse, C. M., 184 Whiteley, K., 223 Whitesides, G. M., 270 Whitley, G. S., 293, 294 Wick, N., 4 Wickline, S. A., 150, 151 Wiecek, A., 131 Wiedeman, M. P., 236 Wiegand, S. J., 5, 127 Wiener, M. C., 190 Wiesmann, C., 133, 134 Wigle, J. T., 9, 10 Wiig, H., 343 Wijelath, E. S., 305 Wikner, C., 18 Wiley, M. J., 192 Wilhelm, S. M., 127 Wilkie, D., 127 Wilkins, M. R., 180 Wilkinson, R. W., 131 Willenbring, H., 54 Williams, B. B., 338 Williams, D. S., 144, 146 Williams, G., 342 Williams, L. E., 166 Williams, M., 10, 304 Williams, S. K., 192, 194 Williams, T. A., 150 Willman, J. K., 154 Willoughby, D. A., 3, 4 Wilmes, T., 89, 95 Wilner, W., 212 Wilson, I., 144, 145 Wilson, M., 245 Wilson, M. E., 245 Wilson, S. R., 144, 147 Winer, J., 126, 127 Winkler, J. D., 3, 4 Winn, V., 288, 296 Winter, H., 361

Winter, P. M., 150 Winterhager, E., 293, 295, 297 Wirtz, S., 15 Wirzenius, M., 343 Wise, L., 5 Wiseman, D. M., 5 Witte, A., 89 Witte, L., 54, 55, 304, 342 Witzenbichler, B., 54, 305, 342 Wobus, C., 88 Woenne, E. C., 150 Wojcicki, W. E., 144, 145, 147 Wolf, G. L., 5 Wolf, K., 143 Wolf, R. L., 144, 146 Wolfram, S., 330 Wolitzky, B. A., 261 Woll, P. J., 343 Wolleschensky, R., 267 Wolters, D., 181, 183, 189 Wong, J., 126 Wong, J. W., 191 Wong, M. H., 54 Wong, S. F., 184 Wong, W. P., 245 Woo, K., 282 Wood, C. G., 95 Wood, J., 5 Woodard, W., 306 Wooding, F. B. P., 231 Woodson, A. K., 151 Work, L. M., 88, 89 Wree, A., 95 Wright, K. C., 95 Wright, N., 130 Wu, A. M., 166 Wu, G., 231, 234, 243, 247, 248 Wu, J. C., 151 Wu, M. H., 305 Wu, S., 215, 216 Wu, S.-P., 209 Wu, W., 367 Wu, W. W., 190 Wu, X., 128, 131, 133, 134, 168, 304, 306, 314, 315, 318, 319, 320, 322 Wu, Y., 4, 5, 6, 10, 15, 18, 54, 55, 148, 156, 162, 166, 342 Wyckoff, J. B., 344 X Xavier, R., 29 Xia, Y. P., 6, 15 Xiang, W., 263, 264 Xie, D., 133, 134 Xie, J., 153 Xie, K., 126 Xie, X., 265

402

Author Index

Xing, L., 151 Xing, M. M., 270 Xiong, J. P., 149, 367 Xiong, Z., 151, 162, 168 Xu, C., 153 Xu, Y., 95 Xue, X. N., 343 Xun, Z., 190 Y Yagmurdur, M. C., 343 Yamada, T., 190 Yamagishi, T., 210 Yamamoto, N., 38, 40, 41, 45, 46 Yamamoto, S., 210 Yamaoka, Y., 8 Yamashiro, D. J., 128, 129 Yamashiro, S., 343 Yamashita, Y., 128 Yamauchi, K., 38, 40, 41, 45, 46 Yan, J. X., 182 Yancopoulos, G. D., 6, 15 Yang, L., 54, 55 Yang, M., 30, 31, 32, 34, 36, 37, 42, 43, 44, 46, 47, 48 Yang, W., 88 Yang, Y., 131 Yang, Z. Q., 282 Yao, J. C., 126, 131 Yao, V. J., 85, 86, 87, 98 Yao, X., 189, 190 Yasuda, K., 343 Yates, J. R., 191, 194, 196, 197 Yates, J. R. III, 181, 183, 189 Yates, N. A., 190 Yayon, A., 161 Ye, W., 127 Yeung, I., 144, 146 Yi, E. C., 189, 190 Yin, G., 5 Ying, Y., 89 Ying, Y. S., 195 Yla-Herttuala, S., 4 Yoder, M. C., 302, 303, 305, 306, 309, 311, 313, 314, 315 Yokoi, A., 129 Yokoyama, K., 157 Yoo, E. S., 88 Yoon, C. H., 305 Yoon, G. S., 88 Yoshida, M., 343 Yoshida, S., 8 Yoshimoto, M., 157 Yoshimura, T., 343 You, Y. N., 131 Young, J., 148, 158 Young, L. M., 55 Young, P. P., 56

Youngs, C. R., 245 Yu, J., 179, 191, 192, 193, 194, 196, 197 Yu, K., 211 Yu, L., 128, 133, 134 Yuan, H., 128 Yuan, J., 268 Yue, X., 211 Z Zacchigna, S., 302 Zalos, G., 305 Zaloudek, C., 129 Zamir, E. A., 108 Zamir, L., 270 Zaoui, K., 89, 90 Zarbock, A., 253, 260, 263, 266 Zarska, M., 212 Zavaleta, C. L., 151 Zdravkovic, 290 Zeng, W., 190 Zentgraf, H., 150 Zentilin, L., 302 Zenz, R., 6 Zhai, Y., 331 Zhang, C., 88, 150 Zhang, F., 55 Zhang, G. Y., 280 Zhang, H., 150, 151, 190 Zhang, J., 126 Zhang, J. B., 95 Zhang, L., 85, 280 Zhang, M., 88, 128 Zhang, R., 149 Zhang, X., 127, 151, 162, 166, 168, 211, 256 Zhang, Y., 182, 331 Zhao, J., 145 Zhao, Y., 257, 258, 268 Zheng, L., 303 Zhong, C., 128 Zhou, D., 343 Zhou, H., 190 Zhou, J. M., 303 Zhou, Y., 281, 282, 288, 291, 296 Zhou, Y. T., 155 Zhu, L., 343 Zhu, M. J., 236, 237, 238 Zhu, X. P., 144, 146 Zhu, Y. M., 343 Zhu, Z., 55, 126, 304 Zhuang, Z. W., 331 Ziegelhoeffer, T., 330, 339 Zimmermann, B., 267 Ziskind, A., 304 Zoldan, J., 304 Zurita, A. J., 84, 86, 87, 88, 89, 95 Zweifach, B. W., 254 Zweit, J., 167

Subject Index

A

B

Adeno-associated virus phage, see Phage display Arteriogenesis, see also Avian embryo vascular morphogenesis models; Coronary vasculogenesis anatomical assessment computed tomography angiography bismuth contrast agent preparation, 334 coronary vasculature filling, 335 micro-computed tomography image analysis, 333 mouse preparation and scanning, 331–333 overview, 331 microangiography coronary fluorescent microangiography, 336–337 X-ray microangiography, 335–336 functional assessment electron paramagnetic resonance tissue oxymetry, 338 laser-Doppler perfusion imaging, 337–338 time-of-flight magnetic resonance imaging, 339 overview, 330 Avian embryo vascular morphogenesis models chamber preparation culture chamber, 109–113 electroporation chamber, 113, 115 coronary vasculogenesis studies embryo preparation, 213 proepicardium microdissection, 213–215 whole-mount immunostaining, 220 culture dish preparation, 115–116 electroporation of fluorescent protein plasmids embryo whole-mounts, 117–119 in ovo electroporation, 119–120 embryo staging and preparation, 116 endothelial cell and extracellular matrix tagging, 116–117 fixation and processing, 120–121 immunolabeling of whole mounts, 121 insert preparation, 109 microscope incubator construction, 113 overview, 108–109 plastic embedding and sectioning, 121–122

Bioinformatics, vascular proteomics, 186–187 Biopanning and rapid analysis of selective interactive ligands differential centrifugation and unbound phage removal, 95–96 microtiter plate preparation, 95 phage recovery and amplification, 96 phage selection, 95 preselection, 95 principles, 94–95 second round selection, 96–97 single-clone binding assays on cells, 97 Blood-derived endothelial progenitor cells, see Endothelial progenitor cells Bone marrow-derived cells angiogenesis role in tumors, 54–55 markers, 54–56 ontogeny, 54 tumor analysis cell subpopulation visualization in tumors cardiac perfusion, tissue fixation, and processing, 59–61 immunohistochemical staining of modulatory cells in tumor sections, 61–63 flow cytometry quantification of cell subpopulations in tumors, 63–67 green fluorescent protein-expressing bone marrow transplantation, 57–59 pericyte progenitor analysis differentiation culture, 73–76 immunofluorescence microscopy, 70 immunomagnetic separation, 72–73 markers, 67–69 materials, 69–70 mature cell detection after differentiation, 76–78 single-cell suspension preparation, 71–72 BRASIL, see Biopanning and rapid analysis of selective interactive ligands Bromodeoxyuridine, proliferating endothelial cell analysis, 11–12, 17 C CD11b, magnetic bead purification of myeloid cells, 349, 351

403

404 CD31, see Platelet endothelial cell adhesion molecule–1 CD34, angiogenesis marker, 10 CD144, see VE-cadherin Chick embryo, see Avian embryo vascular morphogenesis models Computed tomography arteriogenesis assessment computed tomography angiography bismuth contrast agent preparation, 334 coronary vasculature filling, 335 micro-computed tomography image analysis, 333 mouse preparation and scanning, 331–333 overview, 331 tumor angiogenesis functional imaging, 146–147 structural imaging, 145 Confocal microscopy coronary vasculogenesis, 221–222 leukocyte–endothelium interactions autoperfused flow chamber, 268–270 microfluidic flow chambers, 270–272 principles, 266–268 tumor angiogenesis imaging in fluorescent protein mouse models multi-color imaging, 49 single color imaging, 48–49 Coronary vasculogenesis antigen retrieval, 221 artery formation and smooth muscle cell recruitment, 212–213 avian embryo preparation, 213 whole-mount immunostaining, 220 casting, 223–225 computed tomography angiography and coronary vasculature filling, 335 confocal microscopy, 221–222 coronary fluorescent microangiography, 336–337 epicardium explant culture, 217–218 scanning electron microscopy in vivo, 218–219 b-galactosidase whole-mount immunostaining, 222 platelet endothelial cell adhesion molecule whole-mount immunostaining in mouse embryo, 220–221 proepicardium development, 210–211 gene expression analysis reverse transcriptase–polymerase chain reaction, 217 RNA isolation, 216–217 microdissection in avian embryos, 213–215

Subject Index

mouse embryo explant culture, 215–216 scanning electron microscopy in vivo, 218–219 smooth muscle marker whole-mount immunostaining, 222 subepicardium, 211–212 CT, see Computed tomography CTB, see Cytotrophoblast Cytotrophoblast adhesion molecules, 282 culture, 287 endothelial cell coculture first trimester cytotrophoblasts in Matrigel, 292–293 jar choriocarcinoma cells with endothelial monolayers, 292 endovascular invasion studies with explanted spiral arterioles first-trimester decidual explants, 295 myometrial spiral artery segments, 293–294 human placental villous explant isolation and culture, 288–289 isolation dissection, 285 enzymatic digestions, 285, 287–288 materials, 284 Percoll gradient centrifugation, 286–287 mouse xenograft studies of vascular remodeling choriocarcinoma xenograft of skin, 297 mammary fat pad and kidney capsule xenografts, 296–297 remodeled blood vessel identification in tissue sections, 289–291 uterine invasion overview, 280–282 D DOTA-VEGF121, see Positron emission tomography E Electron microscopy, proepicardium scanning electron microscopy in vivo, 218–219 Electron paramagnetic resonance, tissue oxymetry for arteriogenesis assessment, 338 Endothelial cell cytotrophoblast coculture first trimester cytotrophoblasts in Matrigel, 292–293 jar choriocarcinoma cells with endothelial monolayers, 292 fluorescence-activated cell sorting studies of (lymph) angiogenesis, 12, 18–20 myeloid cell adhesion assay, 356–357 proliferation analysis in inflammation bromodeoxyuridine, 11–12, 17 Ki67, 17–18

405

Subject Index

proteomics analytical approaches, 196–197 angiogenesis markers, 193, 196 applications in vasculature analysis, 192–193 mass spectrometry, 201–202 phenotypic drift in culture, 194–195 plasma membrane isolation cell monolayers, 200–201 luminal endothelial cell plasma membrane purification, 200 lung perfusion ad processing, 199–200 solutions, 198–199 tagging in avian embryo vascular morphogenesis models, 116–117 therapeutic targeting in tumor angiogenesis, 179 Endothelial progenitor cells, see also Bone marrow-derived cells expansion, 314–316 flow cytometry characterization, 310, 314–317 isolation from blood adult blood-derived cells, 309, 311–312 cord blood-derived cells, 308–309 magnetic bead purification of CD31positive cells, 312–313 materials, 306–308 overview, 305–306 phenotypic characterization, 310 postnatal vasculogenesis model, 304–305 role, 302 smooth muscle cell coculture smooth muscle cell growth, 318 vasculogenic assay in vivo, 318–323 EPCs, see Endothelial progenitor cells Epicardium explant culture, 217–218 scanning electron microscopy in vivo, 218–219 Flow cytometry bone marrow-derived cells and quantification of cell subpopulations in tumors, 63–67 inflammation and analysis of (lymph) angiogenesis, 12, 18–20 myeloid cell studies in tumor angiogenesis, 349–350, 352, 363 F Fluorescence-mediated tomography, tumor angiogenesis imaging, 147 FMT, see Fluorescence-mediated tomography G GFP, see Green fluorescent protein Green fluorescent protein bone marrow-derived cells in tumors, see Bone marrow-derived cells tumor angiogenesis models, see Tumor angiogenesis

H Histone H2B, green fluorescent protein vector production for tumor angiogenesis studies, 42 I Inflammation cancer association, 344–345 fluorescence-activated cell sorting studies of (lymph) angiogenesis, 12, 18–20 immunofluorescence studies of (lymph) angiogenesis, 10–12, 15–18 mouse models, 5–6, 8, 14–15 vascular permeability analysis, 13–14 vascular remodeling, 3–5 Inflammatory bowel disease angiogenesis, 4 mouse models, 5–6 Integrins avb3 imaging dual-modality imaging, 152–153 probes, 149–150 single-modality imaging, 150–152 myeloid cells activation assays clustering assay, 353–355 ligand-induced binding site assay, 352–353 expression and neovascular site trafficking, 344 Intravital microscopy leukocyte–endothelium interactions adhesion, 258, 260 fluorescence intravital microscopy, 262–264 large vessels, 265 mixed chimeric mouse studies, 265–266 Peyer’s patch lymphocyte rolling and adhesion, 264–265 rolling, 257–258 surgical considerations, 260–261 transmigration, 260 transillumination intravital microscopy principles, 254–257 tumor angiogenesis mouse fluorescent protein models, 30–31, 46–47 K Ki67, proliferating endothelial cell analysis, 17–18 L Laser-Doppler perfusion imaging, arteriogenesis assessment, 337–338 Leukocyte–endothelium interactions adhesion, 258, 260 confocal/two-photon microscopy autoperfused flow chamber, 268–270

406

Subject Index

Leukocyte–endothelium interactions (cont.) microfluidic flow chambers, 270–272 principles, 266–268 intravital microscopy fluorescence intravital microscopy, 262–264 large vessels, 265 mixed chimeric mouse studies, 265–266 Peyer’s patch lymphocyte rolling and adhesion, 264–265 surgical considerations, 260–261 rolling, 257–258 transmigration, 260 Lymphangiogenesis chronic inflammation, 3–5, 10 immunofluorescence studies, 10–12, 15–17 markers, 2 tumors, 3 Lymphatic vascular endothelial hyaluronan receptor–1 antibody conjugation, 43–44 immunofluorescence microscopy, 15–16 lymphangiogenesis marker, 10 LYVE–1, see Lymphatic vascular endothelial hyaluronan receptor–1 M Macrophage, see Myeloid cells; Tumor-associated macrophage Magnetic resonance imaging arteriogenesis assessment with time-of-flight magnetic resonance imaging, 339 tumor angiogenesis functional imaging, 145–146 structural imaging, 145 integrin avb3 imaging, 150, 152–153 Mass spectrometry, see Proteomics MECA–32 angiogenesis marker, 10 immunofluorescence microscopy, 15–16 Metastasis, mouse fluorescent protein models experimental lymphatic metastasis model, 37–39, 44–45 imaging cell trafficking in lymphatics, 37–39 spontaneous metastasis model, 39–41, 44–45 Monocyte, see Myeloid cells MRI, see Magnetic resonance imaging Myeloid cells angiogenesis factors, 343 differentiation, 342 integrins in neovascular site trafficking, 344 monocyte recruitment to tumors, 343–344 tumor angiogenesis studies adhesion assays endothelial cell adhesion, 356–357 purified ligand adhesion, 355 angiogenesis assays in Matrigel, 361

bone marrow mononuclear cell isolation, 348–349 bone marrow transplantation studies, 365–366 CD11b-positive cell isolation with magnetic beads, 349, 351 cell quantification in tumors flow cytometry, 363 immunohistochemistry, 365 reverse transcriptase–polymerase chain reaction, 363–364 flow cytometry, 349–350, 352 immunohistochemistry, 345–346 integrin activation assays clustering assay, 353–355 ligand-induced binding site assay, 352–353 migration/invasion assays, 358–359 monocyte trafficking studies in vivo, 359–361 peripheral blood mononuclear cell isolation, 346–347 tumor growth studies, 361–363 O Oximetry, see Electron paramagnetic resonance P PCR, see Polymerase chain reaction PECAM–1, see Platelet endothelial cell adhesion molecule–1 Pericyte progenitor cells, see Bone marrowderived cells PET, see Positron emission tomography Phage display adeno-associated virus phage vector system, 89 animal vascular diversity screening in vivo animals, 98 phage selection, 98 principles, 97–98 second round selection, 99–100 single-clone binding assays, 100 synchronous serial selection, 100 tissue isolation, phage recovery, and amplification, 98–99 biopanning and rapid analysis of selective interactive ligands differential centrifugation and unbound phage removal, 95–96 microtiter plate preparation, 95 phage recovery and amplification, 96 phage selection, 95 preselection, 95 principles, 94–95 second round selection, 96–97 single-clone binding assays on cells, 97

407

Subject Index

cell-free screening of libraries on isolated receptors microtiter plate preparation, 92 overview, 91–92 phage selection, recovery, and amplification, 92–93 preselection, 92 second round selection, 93–94 single-clone binding assays on immobilized protein, 94 materials for vascular targeted peptides, 90 phage-derived peptide applications diagnostics, 88 targeted therapies, 87–88 vascular targeted gene delivery and molecular imaging, 88–90 principles, 84–85 single-clone sequencing, 91 titering of phage, 91 vascular ligand targeting in humans, 86–87 vascular specialization in single organs, 87 Placental angiogenesis, see Cytotrophoblast; Uteroplacental angiogenesis Platelet endothelial cell adhesion molecule–1 endothelial marker, 8 immunofluorescence microscopy, 16–17 magnetic bead purification of endothelial progenitor cells, 312–313 mouse embryo whole-mount immunostaining, 220–221 Podoplanin immunofluorescence microscopy, 15–16 lymphangiogenesis marker, 10 Polymerase chain reaction myeloid cell quantification in tumors with reverse transcriptase–polymerase chain reaction, 363–364 proepicardium gene expression analysis with reverse transcriptase–polymerase chain reaction, 217 Positron emission tomography integrin avb3 imaging, 151–153 tumor angiogenesis imaging, 145, 147 vascular endothelial growth factor receptor imaging animal models, 163–164 biodistribution studies, 166 cell-binding assays, 163 copper–64 labeling of DOTA-VEGF121, 164 DOTA conjugation, 162–163 immunofluorescence staining for validation, 167 micro-PET imaging, 164–166 overview, 158–161 VEGF121 preparation, 162 Proepicardium coronary vasculogenesis, 210–211

gene expression analysis reverse transcriptase–polymerase chain reaction, 217 RNA isolation, 216–217 microdissection in avian embryos, 213–215 mouse embryo explant culture, 215–216 scanning electron microscopy in vivo, 218–219 Proteomics bioinformatics, 186–187 databases, 187–188 endothelial cell studies analytical approaches, 196–197 angiogenesis markers, 193, 196 applications in vasculature analysis, 192–193 mass spectrometry, 201–202 phenotypic drift in culture, 194–195 plasma membrane isolation cell monolayers, 200–201 luminal endothelial cell plasma membrane purification, 200 lung perfusion ad processing, 199–200 solutions, 198–199 liquid chromatography–tandem mass spectrometry, 182–183 mass spectrometry principles, 183–185 overview, 180–181 quantitative proteomics label-free approaches, 189–192 overview, 188–189 two-dimensional gel electrophoresis principles, 181–182 Prox1 immunofluorescence microscopy, 16–17 lymphangiogenesis marker, 10 Psoriasis, mouse models, 6, 8, 14–15 Q Quail embryo, see Avian embryo vascular morphogenesis models Quantum dot, integrin avb3 imaging, 151–152 R Reverse transcriptase–polymerase chain reaction, see Polymerase chain reaction Rheumatoid arthritis, angiogenesis, 4 S Sildenafil, compromised pregnancy management, 248 Single-photon emission computed tomography, vascular endothelial growth factor receptor imaging, 156–157 SMC, see Smooth muscle cell

408

Subject Index

Smooth muscle cell coronary vasculogenesis marker immunostaining in whole-mounts, 222 recruitment, 212–213 endothelial progenitor cell coculture smooth muscle cell growth, 318 vasculogenic assay in vivo, 318–323 SPECT, see Single-photon emission computed tomography Subepicardium, coronary vasculogenesis, 211–212 T Tadalafil, compromised pregnancy management, 248 TAM, see Tumor-associated macrophage Tumor angiogenesis bone marrow-derived cells, see Bone marrow-derived cells clinical significance, 142–143, 178 imaging functional imaging computed tomography, 146–147 fluorescence-mediated tomography, 147 magnetic resonance imaging, 145–146 positron emission tomography, 147 ultrasound, 145, 147 integrin avb3 imaging dual-modality imaging, 152–153 probes, 149–150 single-modality imaging, 150–152 molecular imaging principles, 148–149 overview, 143–144 positron emission tomography of vascular endothelial growth factor receptor animal models, 163–164 biodistribution studies, 166 cell-binding assays, 163 copper–64 labeling of DOTA-VEGF121, 164 DOTA conjugation, 162–163 immunofluorescence staining for validation, 167 micro-PET imaging, 164–166 overview, 158–161 VEGF121 preparation, 162 prospects, 167–168 single-photon emission computed tomography of vascular endothelial growth factor receptor, 156–157 structure, 145 vascular endothelial growth factor imaging optical imaging, 156 overview, 153–154 targeted ultrasound, 154–155

mouse fluorescent protein models fluorescence microscopy imaging handheld flashlight as light source, 47 Olympus BH 2-RFCA microscope, 47–48 Olympus IV–100 confocal microscope, 48–49 Olympus OV–100 imaging system, 48 fluorescent protein transduction of tumor cell lines, 42–43 green fluorescent protein nude mouse, 32–35 histone H2B-green fluorescent protein vector production, 42 intravital microscopy, 30–31, 46–47 lymphatic vascular endothelial hyaluronan receptor–1 antibody conjugation, 43–44 metastasis models experimental lymphatic metastasis model, 37–39, 44–45 imaging cell trafficking in lymphatics, 37–39 spontaneous metastasis model, 39–41, 44–45 nestin-driven green fluorescent protein nude mouse, 36–37 overview, 29–30 retrovirus production green fluorescent protein, 41–42 red fluorescent protein, 42 skin flap elevation for imaging, 46 surgical orthotopic implantation, 44 tumor cell injection, 44 tumor tissue sampling for high-resolution imaging, 47 whole-body imaging, 45–46 myeloid cell studies adhesion assays endothelial cell adhesion, 356–357 purified ligand adhesion, 355 angiogenesis assays in Matrigel, 361 bone marrow mononuclear cell isolation, 348–349 bone marrow transplantation studies, 365–366 CD11b-positive cell isolation with magnetic beads, 349, 351 cell quantification in tumors flow cytometry, 363 immunohistochemistry, 365 reverse transcriptase–polymerase chain reaction, 363–364 flow cytometry, 349–350, 352 immunohistochemistry, 345–346 integrin activation assays clustering assay, 353–355

409

Subject Index

ligand-induced binding site assay, 352–353 migration/invasion assays, 358–359 monocyte trafficking studies in vivo, 359–361 peripheral blood mononuclear cell isolation, 346–347 tumor growth studies, 361–363 rodent model overview, 29 Tumor-associated macrophage angiogenesis factors, 343 recruitment, 343–344 Two-dimensional gel electrophoresis, see Proteomics Two-photon microscopy leukocyte–endothelium interactions autoperfused flow chamber, 268–270 microfluidic flow chambers, 270–272 principles, 266–268 U Ultrasound, tumor angiogenesis imaging, 145, 147 Uteroplacental angiogenesis, see also Cytotrophoblast anatomy, 280–281 comparative biology, 231–232 evaluation compromised pregnancy studies, 244–248 early pregnancy, 235 late pregnancy, 235–237 normal pregnancy studies, 238–244 techniques, 235 history of study, 230 placental uptake equation, 233 V Vardenafil, compromised pregnancy management, 248 Vascular endothelial growth factor inflammatory response, 4–5 inhibitors clinical trials in cancer, 134–135 mouse models for testing Apcþ/min mouse, 130–131

humanized vascular endothelial growth factor mouse model, 133–134 MEN1 mouse, 131 RIP-Tag mouse, 132–133 xenografts, 127–129 strategies, 126–127 isoforms, 126 molecular imaging optical imaging, 156 overview, 153–154 positron emission tomography of vascular endothelial growth factor receptor animal models, 163–164 biodistribution studies, 166 cell-binding assays, 163 copper–64 labeling of DOTA-VEGF121, 164 DOTA conjugation, 162–163 immunofluorescence staining for validation, 167 micro-PET imaging, 164–166 overview, 158–161 VEGF121 preparation, 162 prospects, 167–168 single-photon emission computed tomography of vascular endothelial growth factor receptor, 156–157 targeted ultrasound, 154–155 therapeutic targeting in compromised pregnancy, 245, 248 tumor expression, 126 Vascular permeability, inflammation analysis, 13–14 Vascular phage-derived peptides, see Phage display Vasculogenesis heart, see Coronary vasculogenesis postnatal vasculogenesis model systems, 303–305 VE-cadherin, endothelial marker, 8, 10 VEGF, see Vascular endothelial growth factor X X-ray microangiography, arteriogenesis assessment, 335–336