T-Cell Trafficking: Methods and Protocols (Methods in Molecular Biology 616)

ME T H O D S

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MO L E C U L A R BI O L O G Y

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For other titles published in this series, go to www.springer.com/series/7651

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T-Cell Trafficking Methods and Protocols

Edited by

Federica M. Marelli-Berg Department of Immunology, Division of Medicine, Imperial College London, London, UK

Sussan Nourshargh William Harvey Research Institute, Barts and the London School of Medicine & Dentistry, Queen Mary University of London, London, UK

Editors Federica M. Marelli-Berg Department of Immunology Division of Medicine Imperial College London Hammersmith Hospital Du Cane Road London UK W12 0NN [email protected]

Sussan Nourshargh William Harvey Research Institute Barts and the London School of Medicine and Dentistry Queen Mary, University of London Charterhouse Square London UK EC1M 6BQ [email protected]

Additional material for this book can be downloaded from http://extras.springer.com ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-460-9 e-ISBN 978-1-60761-461-6 DOI 10.1007/978-1-60761-461-6 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010921222 © Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Both the development of a functional immune system and the establishment of effective immunity are orchestrated by a series of complex and coordinated migratory events. In the last decade a large number of major discoveries have shed light on the molecular mechanisms of lymphocyte migration and the anatomy of immune responses, including the development of organized lymphoid tissue, the compartmentalized recirculation of lymphocyte subsets, and their targeted access to inflammatory antigenic sites. This has been made possible by the development and increasing availability of novel techniques, particularly in the field of real-time imaging and genetic manipulation. Unlike many other research areas where the use of standard techniques and off-theshelf kits is often sufficient and satisfactory, the study of lymphocyte trafficking requires considerable expertise. The methods compiled in this volume are contributed by internationally recognized experts in their respective fields who have many years of experience not only in using these techniques but also in troubleshooting and perfecting them. Each chapter contains a step-by-step description of the method and, more importantly, it provides invaluable tips and tricks to safeguard against potential mishaps and pitfalls. The volume is introduced by an excellent comprehensive review of the current knowledge of lymphocyte trafficking, accessible to the non-specialist. The methods in these chapters are divided into three parts. The first part covers state-of-the-art protocols to study lymphocyte migration and T-cell–endothelial cell interactions in vitro. The second part covers various approaches used for the direct visualization of the development of the lymphoid system, lymphocyte recirculation, and effector responses in experimental models in vivo. Finally, a third section is dedicated to the study of lymphocyte migration and inflammation in the human system and how such investigations can lead to the identification of prognostic markers and the identification of novel therapeutic approaches. We are confident that this book will be an essential manual for newcomers to this ever-expanding and exciting area of research as well as a valuable addition to more specialized laboratories. We would like to express our gratitude to all the authors who have made this book possible and thank Professor John Walker, the Series Editor, for his guidance in editing this volume. We would also like to express our appreciation to Abigail Woodfin for providing the cover art for this volume. F. Marelli-Berg’s lab is generously supported by the British Heart Foundation, The Gates’ Foundation and the Medical Research Council of the UK. S. Nourshargh’s work is supported by generous funds from The Wellcome Trust and the British Heart Foundation. Please note that additional material for this book can be downloaded from http://extras.springer.com Federica Marelli-Berg Sussan Nourshargh

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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SECTION I 1.

INTRODUCTORY REVIEW

How T Cells Find Their Way Around . . . . . . . . . . . . . . . . . . . . . . . Alf Hamann

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SECTION II MIGRATION OF T CELLS IN VITRO 2.

Live Imaging of Leukocyte–Endothelium Interactions . . . . . . . . . . . . . . . Olga Barreiro, Francisco Sánchez-Madrid, and María Yáñez-Mó

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Leucocyte Adhesion Under Haemodynamic Flow Conditions . . . . . . . . . . . Charlotte Lawson, Marlene Rose, and Sabine Wolf

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Influence of Stromal Cells on Lymphocyte Adhesion and Migration on Endothelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Helen M. McGettrick, Chris D. Buckley, G. Ed Rainger, and Gerard B. Nash

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Discriminating Between the Paracellular and Transcellular Routes of Diapedesis . . Jaime Millán, Eva Cernuda-Morollón, and Severine Gharbi

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6.

Monitoring RhoGTPase Activity in Lymphocytes . . . . . . . . . . . . . . . . . Marouan Zarrouk, David Killock, and Aleksandar Ivetic

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Visualisation of Signalling in Immune Cells . . . . . . . . . . . . . . . . . . . . Leo M. Carlin, Konstantina Makrogianneli, Melanie Keppler, Gilbert O. Fruhwirth, and Tony Ng

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Methods for Quantitation of Leukocyte Chemotaxis and Fugetaxis . . . . . . . . 115 Fabrizio Vianello, Elda Righi, and Mark C. Poznansky

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Analysis of CXCR3 and Atypical Variant Expression and Signalling in Human T Lymphocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 125 Anna Korniejewska, Malcolm Watson, and Stephen Ward

10. Transfection of Indoleamine 2,3 Dioxygenase in Primary Endothelial Cells . . . . 149 Petros XE Mouratidis and Andrew JT George SECTION III MIGRATION OF T CELLS IN VIVO 11. Visualisation of Lymphoid Organ Development . . . . . . . . . . . . . . . . . . 161 Henrique Veiga-Fernandes, Katie Foster, Amisha Patel, Mark Coles, and Dimitris Kioussis 12. Single-Cell Analysis of Cytotoxic T Cell Function by Intravital Multiphoton Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Thorsten R. Mempel

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13. Imaging Interactions Between the Immune and Cardiovascular Systems In Vivo by Multiphoton Microscopy . . . . . . . . . . . . . . . . . . . . . . . . 193 Owain R. Millington, James M. Brewer, Paul Garside, and Pasquale Maffia 14. Applying an Adaptive Watershed to the Tissue Cell Quantification During T-Cell Migration and Embryonic Development . . . . . . . . . . . . . . . . . . 207 D. Zhu, S. Jarmin, A. Ribeiro, F. Prin, S.Q. Xie, K. Sullivan, J. Briscoe, A.P. Gould, Federica M. Marelli-Berg, and Y. Gu SECTION IV MONITORING T-CELL MIGRATION IN HUMAN DISEASES 15. Identifying Homing Interactions in T-Cell Traffic in Human Disease . . . . . . . 231 Patricia F. Lalor, Stuart M. Curbishley, and David H. Adams 16. Tracking Antigen-Experienced Effector T Cells In Vitro and In Vivo . . . . . . . 253 Claire L. Gorman, Claudia Monaco, Enrico Ammiratti, Anna-Chiara Vermi, Federica M. Marelli-Berg, and Andrew P. Cope 17. Preclinical Testing of Strategies for Therapeutic Targeting of Human T-Cell Trafficking In Vivo . . . . . . . . . . . . . . . . . . . . . . . 267 Caroline Coisne and Britta Engelhardt Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 281

Contributors DAVID H. ADAMS • Liver Research Group, Division of Medicine, Institute of Biomedical Research, MRC Centre for Immune Regulation, University of Birmingham, Birmingham, UK ENRICO AMMIRATTI • Clinical Cardiovascular Biology Research Centre, Vita-Salute San Raffaele University and San Raffaele Scientific Institute, Milan, Italy OLGA BARREIRO • Servicio de Inmunología, Hospital de la Princesa, Universidad Autónoma de Madrid; Centro Nacional de Investigaciones Cardiovasculares, Madrid, Spain. JAMES M. BREWER • Centre for Biophotonics, Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde, Glasgow, UK JAMES BRISCOE • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK CHRIS D. BUCKLEY • Centre for Cardiovascular Sciences and Centre for Immune Regulation, The Medical School, University of Birmingham, Birmingham, UK LEO M. CARLIN • Cancer Studies Division/Randall Division of Cellular and Molecular Biophysics, Richard Dimbleby Department of Cancer Research, Guy’s Medical School Campus, King’s College London, London, UK EVA CERNUDA-MOROLLÓN • Unidad de Histocompatibilidad, Hospital Universitario Central de Asturias, Oviedo, Spain CAROLINE COISNE • Theodor Kocher Institute, University of Bern, Bern, Switzerland MARK COLES • Division of Molecular Immunology, MRC National Institute for Medical Research, The Ridgeway, Mill Hill, London; Centre for Immunology and Infection, Department of Biology & HYMS, University of York, York, UK ANDREW P. COPE • The Kennedy Institute of Rheumatology, Faculty of Medicine, Imperial College London, UK STUART M. CURBISHLEY • Liver Research Group, Division of Medicine, Institute of Biomedical Research, MRC Centre for Immune Regulation, University of Birmingham, Birmingham, UK BRITTA ENGELHARDT • Theodor Kocher Institute, University of Bern, Bern, Switzerland KATIE FOSTER • Division of Molecular Immunology, MRC National Institute for Medical Research. The Ridgeway, Mill Hill, London, UK GILBERT O. FRUHWIRTH • Cancer Studies Division/Randall Division of Cellular and Molecular Biophysics, Richard Dimbleby Department of Cancer Research, Guy’s Medical School Campus, King’s College London, London, UK PAUL GARSIDE • Centre for Biophotonics, Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde, Glasgow, UK ANDREW JT GEORGE • Department of Immunology, Hammersmith Campus, Imperial College London, London, UK

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SEVERINE GHARBI • Department of Immunology and Oncology, Centro Nacional de Biotecnología. CSIC, Cantoblanco, Madrid, Spain CLAIRE L. GORMAN • Faculty of Medicine, Imperial College London, The Kennedy Institute of Rheumatology, London, UK ALEX P. GOULD • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK YAN GU • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK ALF HAMANN • Experimentelle Rheumatologie, CC12, Charité Universitätsmedizin Berlin, Germany ALEKSANDAR IVETIC • Cytoskeleton Research Group, Division of Cardiovascular Sciences, Faculty of Medicine, Imperial College London, National Heart and Lung Institute, Hammersmith Hospital Campus, DuCane Road, London, UK SARAH JARMIN • Department of Immunology, Faculty of Medicine, Imperial College London, Hammersmith Hospital Campus, London, UK MELANIE KEPPLER • Cancer Studies Division/Randall Division of Cellular and Molecular Biophysics, Richard Dimbleby Department of Cancer Research, Guy’s Medical School Campus, King’s College London, London, UK DAVID KILLOCK • Cytoskeleton Research Group, Division of Cardiovascular Sciences, Faculty of Medicine, Imperial College London, National Heart and Lung Institute, Hammersmith Hospital Campus, DuCane Road, London, UK DIMITRIS KIOUSSIS • Division of Molecular Immunology, MRC National Institute for Medical Research, The Ridgeway, Mill Hill, London, UK ANNA KORNIEJEWSKA • Department of Pharmacy and Pharmacology, University of Bath, Claverton Down, Bath, Slough, UK PATRICIA F. LALOR • Liver Research Group, Division of Medicine, Institute of Biomedical Research, MRC Centre for Immune Regulation, University of Birmingham, Birmingham, UK CHARLOTTE LAWSON • Veterinary Basic Sciences, Royal Veterinary College, London, UK PASQUALE MAFFIA • Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde, Glasgow, UK; Department of Experimental Pharmacology, School of Biotechnological Sciences, University of Naples Federico II, Naples, Italy KONSTANTINA MAKROGIANNELI • Cancer Studies Division/Randall Division of Cellular and Molecular Biophysics, Richard Dimbleby Department of Cancer Research, Guy’s Medical School Campus, King’s College London, London, UK FEDERICA M. MARELLI-BERG • Division of Medicine, Department of Immunology, Faculty of Medicine, Imperial College London, Hammersmith Hospital Campus, Du Cane Road, London, UK HELEN M. MCGETTRICK • Centre for Cardiovascular Sciences and Centre for Immune Regulation, The Medical School, University of Birmingham, Birmingham, UK THORSTEN R. MEMPEL • Center for Immunology and Inflammatory Diseases and Center for Systems Biology, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA, USA JAIME MILLÁN • Centro de Biología Molecular Severo Ochoa, CSIC-UAM, Cantoblanco, Madrid, Spain

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OWAIN R. MILLINGTON • Centre for Biophotonics, Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde, Glasgow, UK CLAUDIA MONACO • Faculty of Medicine, Imperial College London, The Kennedy Institute of Rheumatology, London, UK PETROS XE MOURATIDIS • Department of Immunology, Hammersmith Campus, Imperial College London, London, UK GERARD B. NASH • Centre for Cardiovascular Sciences and Centre for Immune Regulation, The Medical School, University of Birmingham, Birmingham, UK TONY NG • Cancer Studies Division/Randall Division of Cellular and Molecular Biophysics, Richard Dimbleby Department of Cancer Research, Guy’s Medical School Campus, King’s College London, London, UK AMISHA PATEL • Division of Molecular Immunology, MRC National Institute for Medical Research, Mill Hill, London, UK MARK C. POZNANSKY • Infectious Diseases Unit and DFCI/Harvard Cancer Center, Harvard Medical School, Massachusetts General Hospital, Charlestown, MA, USA. FABRICE PRIN • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK G. ED RAINGER • Centre for Cardiovascular Sciences and Centre for Immune Regulation, The Medical School, University of Birmingham, Birmingham, UK ANA RIBEIRO • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK ELDA RIGHI • Infectious Diseases Unit and DFCI/Harvard Cancer Center, Harvard Medical School, Massachusetts General Hospital, Charlestown, MA, USA. MARLENE ROSE • Veterinary Basic Sciences, Royal Veterinary College, London NW1 0TU; National Heart and Lung Institute, Imperial College London, Harefield Hospital & Heart Science Centre, Hill End Road, Harefield, Middlesex, UK FRANCISCO SÁNCHEZ-MADRID • Servicio de Inmunología, Hospital de la Princesa, Universidad Autónoma de Madrid; Centro Nacional de Investigaciones Cardiovasculares, Madrid, Spain KATE SULLIVAN • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK HENRIQUE VEIGA-FERNANDES • Division of Molecular Immunology. MRC National Institute for Medical Research, Mill Hill, London, UK; Immunobiology Unit., Faculdade de Medicina de Lisboa, Instituto de Medicina Molecular, Lisboa, Portugal. ANNA-CHIARA VERMI • Clinical Cardiovascular Biology Research Centre, Vita-Salute San Raffaele University and San Raffaele Scientific Institute, Milan, Italy FABRIZIO VIANELLO • Department of Haematology, Imperial College of Medicine, Hammersmith Hospital, London, UK STEPHEN WARD • Department of Pharmacy and Pharmacology, University of Bath, Claverton Down, Bath; Bath Road, Slough, UK MALCOLM WATSON • Department of Pharmacy and Pharmacology, University of Bath, Claverton Down, Bath; Bath Road, Slough, UK SABINE WOLF • Veterinary Basic Sciences, Royal Veterinary College, Royal College Street, London, UK

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Contributors

SHEILA Q. XIE • MRC Clinical Sciences Centre, Faculty of Medicine, Imperial College London, Hammersmith Hospital Campus, London, UK MARÍA YÁNEZ-MÓ • Servicio de Inmunología, Hospital de la Princesa, Universidad Autónoma de Madrid; Centro Nacional de Investigaciones Cardiovasculares, Madrid, Spain. MAROUAN ZARROUK • Cytoskeleton Research Group, Division of Cardiovascular Sciences, Faculty of Medicine, Imperial College London, National Heart and Lung Institute, Hammersmith Hospital Campus, DuCane Road, London, UK DAAN ZHU • Developmental Neurobiology, National Institute for Medical Research, MRC, London, UK

Section I Introductory Review

Chapter 1 How T Cells Find Their Way Around Alf Hamann Abstract Among diverse cellular systems of the body, the immune system is unique in representing a network of interacting cells of enormous complexity yet based on single cells travelling around. Only the advanced visualization technologies of the recent years have brought to everybody’s attention the fact that what we see is usually a snapshot of a dynamic system, where the majority of players are highly motile and become coordinated by diverse signals provided by their environment. This introductory chapter touches a selection of aspects that address predominantly the functioning of the system as such. It attempts to provide a framework of how migratory mechanisms are regulated to ensure that various cell populations reach their destination and that the appropriate interaction partners find each other. Key words: T cells, recirculation, tissue entry, tissue exit, homing, inflammation, adhesion molecules, chemokines, chemokine receptors, antigen, α4 β7 integrin, memory, epigenetics, imprinting.

1. Recirculation: Come and See Fifty years ago J. Gowans discovered that lymphocytes possess the unique property to recirculate continuously between blood, lymphoid tissues and lymph (1). In contrast to myeloid cells, which, by and large, are only known to travel unidirectionally, the recirculation of naive lymphocytes ensures that antigen presented locally is seen by as many as possible cells carrying the enormous diverse repertoire of T and B cell receptors. Two major travelling routes through lymphoid tissues mutually complement each other that are regulated by different migratory mechanisms: the circulation through the blood with a stopover in the spleen on the one side and through both the F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_1, © Springer Science+Business Media, LLC 2010

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blood and the lymphatic system via lymph nodes, Peyer’s patches and tertiary lymphoid tissue arising in chronically inflamed tissues as gateways. Much research has focussed on the receptors guiding lymphocyte entry into lymph nodes and Peyer’s patches. This process is mediated by a well-known set of adhesion molecules assisted by the integrin-activating function of chemokine receptors that are triggered by chemokines presented on the endothelial surface. In fact, L-selectin, which makes the first contacts for naive lymphocytes with the high endothelial venules of (predominantly, but not exclusively peripheral) lymph nodes, was the first homing-related molecule to be identified (2). Subsequent research discovered the integrins LFA-1 and α4 -integrins as indispensable components of the more complex process of transmigration that also contribute to organ-specific properties of migration. L-selectin was found to direct especially naïve lymphocytes into lymph nodes, whereas α4 β7 guides cells into mucosa-associated lymphoid tissues and also into the gut wall itself. LFA-1 is less selective and contributes to transmigration in most tissues including inflamed sites. The sequential operation of these molecules during migration from blood to tissue had led to the proposal of the multi-step model of transmigration (3), which is now part of every textbook. In contrast to the lymphoid tissues named above, entry into the spleen is not dependent on homing or chemokine receptors; all evidence so far available suggests that the entry is solely driven by mechanics of blood flow and cell motility (4, 5). Yet, localization within the tissue and different compartments therein are regulated by chemotaxis and likely also by interactions with stromal cells or extracellular matrix.

2. Within Tissue: Come and Go (or One Comes, One Leaves)

Rather recent is the insight that not only the entry into tissue but also the exit requires defined molecular systems. As a simple matter of fact, the frequency of cells disposed of in a tissue can efficiently be regulated not only by the rate of entry but also by modulating the exit rate. Findings of the last years have shown that exit from the tissue is an active process controlled by chemotactic mechanisms. The chemokine receptor CCR7 was shown to be required for T cell, including Treg, exit from inflamed peripheral tissue (6, 7). Another chemotactic agent sphingosine1-phosphate (S1P) and its receptors are required for the exit from lymph nodes, a finding emerging from studies with the drug FTY 720 which displays immunosuppressive effects. Both CCR7 and S1P receptors are modulated in the course of T cell activation and

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thereby might cause the transient retention of recently activated T cells in the lymph node (8). When CCR7 is knocked out, the number of T cells retained in an inflamed tissue doubles, confirming its importance for continuous circulation (9). By technical reasons, quantification of exit rates for specific subsets of cells and specific tissues is more difficult. However, a variety of data are available from early studies applying cannulation of the thoracic duct or even single lymph nodes, which provided clear evidence that not only naive cells entering a lymph node via the high endothelium pass the tissue within half a day and exit it but also that large numbers of effector/memory cells attracted to an inflamed tissue or generated by local proliferation exit the tissue via the efferent lymph (10). It is conceivable that the process of emigration underlies a variety of regulatory influences, too; T cell activation upon antigen encounter within the tissue might be one factor, but also an influence of inflammation-generated mediators such as prostaglandins has been described (11).

3. Disposal of Combatants Infiltration of leucocytes into sites of an immune reaction is the hallmark of inflammation. It can be assumed that the evolution of leucocyte trafficking in the context of innate cellular reactions started well before the emergence of lymphocytes. In contrast, the specific variant of recirculation of T and B cells is a rather late invention. Various mechanisms contribute to the rapid delivery of defence cells and their accumulation at sites of risk. Enhanced expression of adhesion molecules, such as MAdCAM-1 in mucosal sites or ICAM-1 in many tissues, as well as induction of additional adhesion molecules such as P- and E-selectin or VCAM-1 on endothelium under conditions of inflammation, are the starting points for a massive adhesion and transmigration of leucocytes from blood into the inflamed tissue, assisted by chemotactic signals. But who tells the endothelium to become activated? Most endothelia do respond to TLR signals such as LPS, but for most inflammatory processes it is more likely that other cells receive the danger signals. Resident leucocytes such as mast cells and dendritic cells are highly sensitive sentinels for stress signals, tissue cells such as fibroblasts or epithelial cells respond to pathogenderived signals, and, in later stages, antigen-specific T cells translate recognition into local conditioning for high-rate recruitment of effector leucocytes. Both innate receptors for pathogen structures and recognition of antigen by T cells lead to activation and

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secretion of a variety of mediators, notably cytokines such as TNF, IFNγ and a large variety of chemokines, that are crucial within the multi-step transmigration process and subsequently guide immigrated leucocytes into the centre of inflammation. In addition, the complement system, activated by Ig-dependent or alternative pathways, contributes significantly to the stimulation of recruitment by production of the chemotactic and activating cleavage products C3a and C5a. The entire process shows properties of a positive feedback loop; the first signals lead to the attraction of a few pioneer cells (45), which, by release of activating cytokines and chemokines, greatly accelerate the recruitment of large numbers of further effector cells. The endothelium represents the interface between inflammatory environment and circulation: its sensitive reaction towards activating cytokines results in the expression of adhesion molecules on the surface. Moreover, its capacity to transport chemokines from the abluminal tissue side to the vessel surface, where they become fixed on glycosaminoglycans, generates tags from the inflammatory environment detectable for circulating leucocytes. It should be mentioned that the immune system also provide mechanisms to shut off the proinflammatory response in order to avoid inappropriate damage and immunopathology. Different types of regulatory T cells, including Foxp3+ Tregs and IL-10 producing Tr1 cells, appear to be generated or expanded during the immune reaction and contribute to limit the response after the acute reaction (12, 13). In addition, mediators such as IFNγ, while fuelling the inflammation at the beginning, induce feedback mechanisms negatively regulating the response and activate inhibitory reactions in myeloid cells such as production of NO, IDO or others (14). Whether these negative feedback mechanisms also directly down-regulate the recruitment machinery, or whether recruitment and activation merely fades away when the effector cells become silenced, remains to be seen. So far, there is little evidence that, e.g., Tregs affect directly the migratory process (Doebis et al., unpublished data).

4. Are T Cells Attracted by Antigen?

As mentioned above, antigen-driven activation of immigrated T effector cells is a major factor in the establishment of a full inflammatory condition. It is a longstanding question to what extent the accumulation of lymphocytes is directly related to their capacity to recognize the antigen. It has been repeatedly reported that antigen-reactive T and B cells become concentrated within a tissue offering the cognate antigen (antigen-induced trapping),

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which can even lead to the complete disappearance of the reactive cells from the circulation (15, 16). On the other side, convincing data were provided excluding an influence of antigen on the entry of lymphocytes into a given tissue (17), although, under certain conditions, endothelium might also present antigen. Rather, contact with the antigen and consequent activation might lead to an alteration of so far not identified cellular properties in the cognate lymphocytes that result in altered exit rates and a prolonged residence of the cells in the respective environment. Evidence for antigen-specific trapping has been presented for lymphoid tissue (15, 16), for the liver (18, 19) and for peripheral tissue (20). In our studies, using a transgenic DTH model, we found an enhanced recruitment of both antigen-specific and non-specific effector T cells into the inflamed cutaneous tissue upon preceding encounter of the specific T cells with cognate antigen, but no selective trapping (45). However, the specific T cells that arrived in the site started to proliferate locally after a few days, resulting in a cellular infiltrate that is strongly enriched for cognate T cells (Doebis et al., submitted).

5. The Paradigm of Organ-Specific Homing

Already in the pioneer years of cellular immunology, the capacity of distinct subpopulations of T or B cells to travel back selectively into compartments of initial antigen contact was recognized and referred to as “homing” (21). Selective homing to the gut mucosa was observed for activated (and later also for memory) cells (blasts) in the blood (21, 22), which, in healthy animals, predominantly originate from the gut environment continuously exposed to a huge burden of bacterial antigen. Application of the famous “Stamper-Woodruff” assay, a crude approach using frozen tissue sections to test for selective adhesion of lymphocytes to the remnants of endothelium (23), was surprisingly effective and led to the detection of L-selectin as major determinant of peripheral lymph node homing (2) and of the integrin α4 β7 as a mucosal homing receptor (24). Albeit this work gave rise to the assumption that distinct homing receptors guide lymphocytes into different tissues, the population of (predominantly) naive lymphocytes used in that studies is – ironically – just the one population that does not show organ-specific homing, apart from the fact that naive lymphocytes are specialized to recirculate through any lymphoid tissue, but cannot enter other types of tissues. It is therefore important to consider that lymphocytes gain organ-specific homing properties only upon activation and differentiation into effector/memory cells. This differentiation step is associated with a major change in the molecular equipment

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required for trafficking: L-selectin as well as the chemokine receptor CCR7, also important for lymph node entry, is downregulated with differentiation into end-stage memory cells, leading to a non-recirculating population with a high capacity to enter inflamed, peripheral tissues. Down-regulation of molecules important for recirculation, acquisition of organ-specific homing receptors and inflammation-seeking receptors and functional differentiation might be, but are not necessarily, synchronized. The widely cited distinction of “central” memory cells from “effectormemory” T cells is misleading and rather picks out two of several possible combinations of functional (cytokine) and homing properties. For example, both, recirculating, CCR7+ T cells and terminally differentiated CCR7-effector/memory cells which localize exclusively in peripheral tissue are ready to produce effector cytokines and can be simultaneously detected in vivo (25, 26). On the other side, upregulation of organ-specific homing receptors starts already with the first divisions upon antigen encounter in vivo (27). Early expectations were to discover a series of homing receptors, each specific for a distinct organ or type of tissue. On the level of adhesion molecules, this only holds true for the interaction partner α4 β7 -integrin and MAdCAM, which guide lymphocytes almost exclusively into gastrointestinal (“mucosal”) sites. The sister integrin α4 β1 was found to be a dominant mediator of T cell migration into the brain, a fact allowing the use of anti α4 antibodies (natalizumab) as the most efficient drug against multiple sclerosis today. However, it is not excluded, that the α4 β1 integrin, which binds predominantly to its ligands VCAM-1 or fibronectin, is also involved in the recruitment of cells to other sites of the body, one definitely being the bone marrow (28, 29), and possibly other sites of inflammation. Similar might apply for the adhesion pair E-selectin (expressed on cutaneous endothelium) and CLA (“cutaneous lymphocyte antigen”, a carbohydrate epitope, expressed on lymphocytes) being considered as a “skin” specific system, at least in humans. However, E-selectin is also frequently expressed in other tissues upon inflammation. Together with P-selectin, with which it shares some common ligands (glycosylated PSGL-1) and acts in a largely redundant way, E-selectin might rather be considered as an inflammationspecific system with some preference for the skin (30). Additional adhesion molecules, such as VAP-1 (31), CD44 (32) and others, might contribute to a significant diversity of potential address codes, but, after all, selectins, α4 -integrins and β2 -integrins and their respective ligands appear to be the working horses of recognition and adhesion to endothelium, with differential, but also widely overlapping use in various destinations of lymphocyte trafficking. As discussed above, the chemokine system pro-

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vides a greater degree of selectivity, although true organ-specific chemokine receptors also might not exist.

6. Sniffing the Way After their discovery 30 years ago, the chemokine family has attracted great interest as being major players in the determination of migratory pathways. The large number of family members of these cytokine-like mediators (more than 40 chemokines in humans) is multiplying the degree of variation provided by adhesion molecules. Interestingly, this diversity has evolved rather recently (with vertebrates) and appears to be driven along the evolution of the adaptive immune system (33, 34), requesting for sophisticated mechanisms to direct the multiple cell types, differentiation and activation stages to appropriate sites and conditions of immune reactivity within the body. Apart from the structural homologies, the chemokine family is characterized by its preponderant reactivity with a – also very diverse – subgroup of receptors (almost 20 receptors in humans) belonging to the huge superfamily of G-protein-coupled receptors. Interestingly, within this superfamily, chemokine receptors share a subtree with the olfactory receptors, the largest subfamily among G-proteinlinked receptors (almost 500 members in man; 33) that allows us to orient ourselves within a complex diversity of chemical signals of the external environment.

7. Topographical Memories For a long period, the paradigm of organ-specific homing included the assumption that T cell priming within a specific tissue environment led to an imprinting of the expression of specific homing receptors. Albeit even recent Nature papers use the term imprinting, what they refer to is only induction of certain homing receptors (35) by cells and mediators. In fact, the same group provided experimental data challenging the concept of permanent imprinting and favouring the assumption of flexibility in the expression of homing receptors (36). Indeed, organ-specific homing could also be explained by continuing selection or reinduction of a given receptor upon recirculation through selected tissues providing antigen exposure and re-stimulatory capacity associated with additional, organ-specific co-signals (37). Studies to proof the stability of differentially expressed homing receptors in vivo were largely lacking until recently. We

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analysed the stability of ligands for E/P-selectins that serve as homing receptors for inflamed tissue, notably inflamed skin. We could provide clear evidence that recently induced selectin ligands are stable only on a subfraction of T cells. However, upon repeated stimulation under ligand-inducing conditions (presence of IL-12) the fraction of cells stably expressing selectin ligands for at least several weeks in vivo increased and ex vivo isolated selectin ligand-positive effector/memory cells turned out to be almost completely stable (38). This shows that imprinting of a stable homing phenotype appears possible, but requires repeated restimulation under permissive conditions, similar to what has been found for the imprinting of a cytokine memory in T cells (39). The above-mentioned studies on the mucosal homing receptor α4 β7 in CD8+ T cells suggested that expression of this receptor is not permanent after initial induction (36). We are presently investigating this issue in CD4+ T cells. Studies not yet completed suggest that, as for selectin ligands, repeated stimulations in the presence of retinoic acid are required to achieve expression of α4 β7 which persist upon restimulation (Szilagyi et al., unpublished). In contrast to the selectin ligands, a continuous drop of the expression of α4 β7 is, however, observed on all adoptively transferred cells in vivo, suggesting that stability subsists only for a limited period, unless appropriate restimulation re-establishes the imprinted phenotype. For the chemokine receptor CCR9, which is also induced (on CD8+ cells) by retinoic acid and considered to contribute to mucosal homing (35), we could not observe a stable expression phenotype (Szilagyi et al., unpublished). These data suggest that variable degrees of a topographical memory might exist, depending on the respective receptors: complete stability, as in case of selectin ligands established after appropriate instructive differentiation of the memory cells; partial stability, which slowly fades away in the absence of permissive signals as in case of α4 β7 ; and lack of stability, as for CCR9 in CD4+ T cells. However, it has to be considered that, even in the absence of a stable expression, an imprinted memory might exist. The term “commitment” is used, for example, for T effector cells, when polarized Th1 or Th2 cells are predestined to secrete the respective typical cytokines, IFNγ or IL-4, but require restimulation to do so. Importantly, a committed cell only needs a reduced set of stimulatory signals to re-acquire the specific phenotype. Further investigations have to show whether this type of memory also exists in case of homing receptors, as some data suggest (38). What mechanisms could allow imprinting of a certain phenotype or commitment for facilitated re-expression? For the cytokine memory, proposed mechanisms include the establishment of a metastable signalling condition, involving positive feedback loops

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in the induction and action of key transcription factors such as GATA-3 (39) and imprinting of information by epigenetic modification in histones (“histone code”) or especially in DNA by differential methylation of CpGs (40). Epigenetic modification leads to alterations in chromatin structure resulting in different degrees of accessibility of a given gene locus. Especially DNA methylation is reproduced upon synthesis of new DNA strains during mitosis and therefore allows inheritance of acquired properties. This mechanism appears to have a key role in the imprinting of developmental changes and determination of lineage decisions (41– 43). So far, only indirect evidence for a role of epigenetic mechanisms in the imprinting of homing receptors has been published (44). However, it is evident that epigenetic imprinting would be ideally suited to match requirements for the acquisition of stable homing phenotypes.

8. Concluding Remarks Every decade of immunological research appears to uncover novel functional subsets of T cells; the latest ones being, e.g., the Th17 cells or the diverse types of regulatory T cells. How this increasing universe of specialists becomes coordinated and appropriately targeted to the hot spots of immunoreactivity would remain a mystery if not, at the same time, our knowledge about the mechanisms of cell trafficking would have greatly expanded. Cooperating adhesion molecules and chemokine receptors equip the migrating cells with an almost unlimited combinatorial diversity to recognize signatures defining tissues and compartments, to distinguish inflammatory processes of multiple flavours that might depend on the kind of triggers, site of inflammation or involved cell populations and so on. That chemotaxis, haptotaxis and cell contacts not only are important to regulate the macroscopic distribution of cells within the body but are equally important to guide cells through the jungle of a tissue environment – and even might support the marriage of individual cell partners destined to interact in a given environment and functional stage – that insight was greatly nourished by recent findings. The present book might provide a number of wonderful pieces of knowledge from this field (46).

Acknowledgements Special thanks to present and past coworkers of my group who contributed to ideas and findings discussed above:

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G. Debes, C. Doebis, S. Floess, S. Ghani, J. Huehn, S. Jennrich, A. Menning, B. Ratsch, K. Siegmund, C. Siewert, U. Syrbe, and B. Szilagyi. Our work was continuously supported by the Deutsche Forschungsgemeinschaft. References 1. Gowans JL. (1959) The recirculation of lymphocytes from blood to lymph in the rat. J Physiol 146, 54–69. 2. Gallatin WM, Weissman IL, Butcher EC. (1983) A cell-surface molecule involved in organ-specific homing of lymphocytes. Nature 304, 30–4. 3. Von Andrian UH, Chambers JD, McEvoy LM, Bargatze RF, Arfors KE, Butcher EC. (1991) Two-step model of leukocyteendothelial cell interaction in inflammation: distinct roles for LECAM-1 and the leukocyte beta 2 integrins in vivo. Proc Natl Acad Sci USA 88, 7538–42. 4. Nolte MA, Hamann A, Kraal G, Mebius RE. (2002) The strict regulation of lymphocyte migration to splenic white pulp does not involve common homing receptors. Immunology 106, 299–307. 5. Grayson MH, Hotchkiss RS, Karl IE, Holtzman MJ, Chaplin DD. (2003) Intravital microscopy comparing T lymphocyte trafficking to the spleen and the mesenteric lymph node. Am J Physiol Heart Circ Physiol 284, H2213–26. 6. Bromley SK, Thomas SY, Luster AD. (2005) Chemokine receptor CCR7 guides T cell exit from peripheral tissues and entry into afferent lymphatics. Nat Immunol 6, 895–901. 7. Debes GF, Arnold CN, Young AJ, et al. (2005) Chemokine receptor CCR7 required for T lymphocyte exit from peripheral tissues. Nat Immunol 6, 889–94. 8. Matloubian M, Lo CG, Cinamon G, et al. (2004) Lymphocyte egress from thymus and peripheral lymphoid organs is dependent on S1P receptor 1. Nature 427, 355–60. 9. Menning A, Höpken UE, Siegmund K, Lipp M, Hamann A, Huehn J. (2007) CCR7 is crucial for the functional activity of both naïve- and effector/memory-like regulatory T cells subsets. Eur J Imm 37, 1575–83. 10. Seabrook T, Au B, Dickstein J, Zhang X, Ristevski B, Hay JB. (1999) The traffic of resting lymphocytes through delayed hypersensitivity and chronic inflammatory lesions: a dynamic equilibrium. Semin Immunol 11, 115–23. 11. McConnell I, Hopkins J, Lachmann P. (1980) Lymphocyte traffic through lymph

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nodes during cell shutdown. Ciba Found Symp 71, 167–95. Belkaid Y. (2007) Regulatory T cells and infection: a dangerous necessity. Nat Rev Immunol 7, 875–88. Li MO, Flavell RA. (2008) Contextual regulation of inflammation: a duet by transforming growth factor-beta and interleukin-10. Immunity 28, 468–76. Feuerer M, Eulenburg K, Loddenkemper C, Hamann A, Huehn J. (2006) Self-limitation of Th1-mediated inflammation by IFN{gamma}. J Immunol 176, 2857–63. Sprent J, Miller JF, Mitchell GF. (1971) Antigen-induced selective recruitment of circulating lymphocytes. Cell Immunol 2, 171–81. Arnold CN, Butcher EC, Campbell DJ. (2004) Antigen-specific lymphocyte sequestration in lymphoid organs: lack of essential roles for alphaL and alpha4 integrindependent adhesion or Galphai proteincoupled receptor signaling. J Immunol 173, 866–73. Ager A, Drayson MT. (1988) Lymphocyte migration in the rat. In: Husband AJ, ed. Migration and Homing of Lymphoid Cells. Boca Raton: CRC Press, 19–49. Bertolino P, Schrage A, Bowen DG, et al. (2005) Early intrahepatic antigen-specific retention of naive CD8+ T cells is predominantly ICAM-1/LFA-1 dependent in mice. Hepatology 42, 1063–71. John B, Crispe IN. (2004) Passive and active mechanisms trap activated CD8+ T cells in the liver. J Immunol 172, 5222–9. Reinhardt RL, Bullard DC, Weaver CT, Jenkins MK. (2003) Preferential accumulation of antigen-specific effector CD4 T cells at an antigen injection site involves CD62Edependent migration but not local proliferation. J Exp Med 197, 751–62. Gowans JL, Knight EJ. (1964) The route of recirculation of lymphocytes in the rat. Proceed Roy Soc London, B 159, 257–82. Smith ME, Martin AF, Ford WL. (1980) Migration of lymphoblasts in the rat. Monogr Allergy 16, 203–32. Stamper HBJ, Woodruff JJ. (1976) Lymphocyte homing into lymph nodes: in

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vitro demonstration of the selective affinity of recirculating lymphocytes for high endothelial venules. J Exp Med 144, 828. Holzmann B, McIntyre BW, Weissman IL. (1989) Identification of a murine Peyer’s patch-specific lymphocyte homing receptor as an integrin molecule with an alpha chain homologous to human VLA-4. Cell 56, 37–46. Debes GF, Bonhagen K, Wolff T, et al. (2004) CC chemokine receptor 7 expression by effector/memory CD4+ T cells depends on antigen specificity and tissue localization during influenza A virus infection. J Virol 78, 7528–35. Hamann A, Arnold CA, Debes GF. (2005) Trafficking of lymphocyte subpopulations. In: Hamann A, Engelhardt B, eds. Leukocyte Trafficking. Weinheim: WILEY-VCH Verlag GmbH & Co. KGaA, 154–73. Campbell DJ, Butcher EC. (2002) Rapid acquisition of tissue-specific homing phenotypes by CD4+ T cells activated in cutaneous or mucosal lymphoid tissues. J Exp Med 195, 135–41. Berlin-Rufenach C, Otto F, Mathies M, et al. (1999) Lymphocyte migration in lymphocyte function-associated antigen (LFA)-1deficient mice. J Exp Med 189, 1467–78. Papayannopoulou T. (2003) Bone marrow homing: the players, the playfield, and their evolving roles. Curr Opin Hematol 10, 214–9. Ley K, Kansas GS. (2004) Selectins in T-cell recruitment to non-lymphoid tissues and sites of inflammation. Nat Rev Immunol 4, 325–35. Jalkanen V, Andersson BM, Bergh A, Ljungberg B, Lindahl OA. (2008) Explanatory models for a tactile resonance sensor system-elastic and density-related variations of prostate tissue in vitro. Physiol Meas 29, 729–45. Nandi A, Estess P, Siegelman M. (2004) Bimolecular complex between rolling and firm adhesion receptors required for cell arrest; CD44 association with VLA4 in T cell extravasation. Immunity 20, 455–65. Fredriksson R, Schioth HB. (2005) The repertoire of G-protein-coupled receptors in fully sequenced genomes. Mol Pharmacol 67, 1414–25. DeVries ME, Kelvin AA, Xu L, Ran L, Robinson J, Kelvin DJ. (2006) Defining the origins and evolution of the

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chemokine/chemokine receptor system. J Immunol 176, 401–15. Mora JR, Bono MR, Manjunath N, et al. (2003) Selective imprinting of gut-homing T cells by Peyer’s patch dendritic cells. Nature 424, 88–93. Mora JR, Cheng G, Picarella D, Briskin M, Buchanan N, von Andrian UH. (2005) Reciprocal and dynamic control of CD8 T cell homing by dendritic cells from skinand gut-associated lymphoid tissues. J Exp Med 201, 303–16. Campbell DJ. (2005) Control of homing receptor expression during lymphocyte differentiation, activation, and function. In: Hamann A, Engelhardt B, eds. Leukocyte Trafficking. Weinheim: WILEY-VCH Verlag GmbH & Co. KGaA, 131–53. Jennrich S, Ratsch BA, Hamann A, Syrbe U. (2007) Long-term commitment to inflammation-seeking homing in CD4+ effector cells. J Immunol 178, 8073–80. Loehning M, Richter A, Radbruch A. (2002) Cytokine memory of T helper lymphocytes. Adv Immunol 80, 115–81. Tykocinski LO, Hajkova P, Chang HD, et al. (2005) A critical control element for interleukin-4 memory expression in T helper lymphocytes. J Biol Chem 280, 28177–85. Lee GR, Kim ST, Spilianakis CG, Fields PE, Flavell RA. (2006) T helper cell differentiation: regulation by cis elements and epigenetics. Immunity 24, 369–79. Wilson CB, Makar KW, Shnyreva M, Fitzpatrick DR. (2005) DNA methylation and the expanding epigenetics of T cell lineage commitment. Semin Immunol 17, 105–19. Reiner SL. (2005) Epigenetic control in the immune response. Hum Mol Genet 14 Spec No 1:R41-6. Syrbe U, Jennrich S, Schottelius A, Richter A, Radbruch A, Hamann A. (2004) Differential regulation of P-selectin ligand expression in naïve versus memory T cells: evidence for epigenetic regulation of involved glycosyltransferase genes. Blood 104, 3243–8. Ghani S, Feuerer M, Doebis C, Lauer U, Loddenkemper C, Huehn J, Hamann A, Syrbe U. (2009) T cells as pioneers: antigenspecific T cells condition inflammed sites for high-rate antigen-non-specific effector cell recruitment. Immunology 128:e870–880. See also recent reviews in: Focus on Leukocyte Trafficking; Nature Immunology 9, 947–1000 (2008).

Section II Migration of T Cells In Vitro

Chapter 2 Live Imaging of Leukocyte–Endothelium Interactions Olga Barreiro, Francisco Sánchez-Madrid, and María Yáñez-Mó Abstract Leukocyte extravasation is a highly dynamic, interactive, and coordinated process that plays a central role during the inflammatory response of innate immunity. The interaction of leukocytes with the activated endothelium under shear forces is comprised of many sequential events, each involving specific leukocyte and endothelial receptors, as well as chemokines and adaptor and signaling molecules. Because of its complexity, researchers studying leukocyte extravasation at the subcellular level have been forced to search for appropriate in vitro models that mimic pathophysiological conditions at sites of inflammation. We report methods for direct visualization of cellular and molecular processes of critical importance to spatiotemporally dissect the different steps in the adhesion cascade. These methodologies include techniques for the study of the dynamics of individual molecules involved in a discrete part of the process, as well as simple procedures to label molecules and cells in order to observe the extravasation process. Key words: Endothelial cells, leukocytes, adhesion, flow, fluorescent proteins, fluorescent probes, confocal microscopy.

1. Introduction Leukocyte transendothelial migration is dependent on the productive interaction of leukocytes with the activated endothelial monolayer of the vasculature supplying the inflamed tissue. This interaction takes place through a series of sequential steps, which confer selectivity to the extravasation process (1). The first step is the interaction of selectins with their carbohydratebased ligands (2) and allows the leukocyte to roll on the endothelial cell wall. Leukocyte rolling increases the chances that

F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_2, © Springer Science+Business Media, LLC 2010

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leukocytes will encounter chemokines presented on the apical endothelial surface. These cytokines activate leukocyte integrins (3) and in cooperation with integrin-dependent signals induce the polarization of the leukocyte (4). Polarized leukocytes firmly adhere to the endothelial monolayer, mainly through the integrin receptor–counterreceptor pairs LFA1/ICAM-1,2 and VLA4/VCAM-1 (5). In this firm adhesion step, binding to endothelial adhesion receptors initiates intracellular signaling cascades that lead to the cytoskeletal rearrangements necessary for the formation of a three-dimensional docking structure that concentrates both ICAM-1 and VCAM-1 and virtually surrounds the adhered leukocyte, preventing its detachment under flow conditions (6). After the adhesion steps, leukocytes cross to the interstitial space either by diapedesis through endothelial lateral junctions or by transcellular migration (7). Leukocyte extravasation involves the coordinated action of multiple receptors, cytoskeletal adaptors, and signaling molecules and results in drastic morphological changes in both leukocytes and endothelial cells. Direct visualization of cellular and molecular dynamics is therefore of critical importance for understanding this process. This report summarizes several approaches to the spatiotemporal analysis of the different steps of the adhesion cascade. These approaches include techniques for the study of individual molecules involved in specific steps as well as simple procedures for labeling molecules and cells to allow observation of the extravasation process in conditions as close to physiological as possible.

2. Materials 2.1. Cell Models

1. For HUVEC culture, 199 medium is used supplemented with 10% FBS, antibiotics, heparin (100 μg/ml), and ECGF (50 μg/ml). Cells are routinely grown in culture flasks coated with 0.5% gelatin 2. Recombinant human TNF-α (R&D Systems) 3. Ficoll-Hypaque high-density medium (Sigma) 4. PHA-L (Sigma) and IL-2, provided by the National Institutes of Health AIDS Research and Reference Reagent program, Division of AIDS. RPMI 1640 (Gibco) supplemented with 10% FBS 5. RPMI 1640 supplemented with 10% FBS, G418 (Calbiochem), and MnCl2

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1. BCECF-AM (Invitrogen) 2. HBSS (BioWhittaker) supplemented with 1% BSA 3. Plastic seal

2.2.1. Static Adhesion Measurements

4. 0.1% SDS in 50 mM Tris–HCl pH 8.5

2.2.2. Static Adhesion Staining

1. FN (20 μg/ml) (Sigma) 2. 4% paraformaldehyde in PBS supplemented with 2% sacarose 3. Tris-buffered saline (TBS): 50 mM Tris–HCl pH 7.5, 150 mM NaCl

2.3. Detachment Experiments

1. A parallel flow chamber (Glycotech) 2. Glass-bottomed Petri dishes (WillCo Wells) 3. A programmable syringe pump (Harvard Apparatus) 4. HBSS supplemented with 2% FBS at 37◦ C 5. Real-time monochrome camera coupled to a video recorder system 6. Water bath

2.4. Leukocyte– Endothelium Interactions Under Flow Conditions

1. A parallel flow chamber (Glycotech) 2. A programmable syringe pump (Harvard Apparatus) 3. Glass-bottomed Petri dishes (WillCo Wells) 4. HBSS supplemented with 2% FBS 5. Water bath

2.5. Transfection Procedures for Fluorescently Tagged Fusion Proteins 2.5.1. Transfection of Endothelial Cells

1. 199 culture medium supplemented with growth factors 2. 1.5 M NaCl 3. DNA plasmids 4. Electroporator suitable for mammalian cells Gene Pulser Xcell (Bio-Rad Laboratories) – Electroporation cuvettes (4 mm) (Bio-Rad) – Coated coverslips

2.5.2. Transfection of Leukocytes

R 1. Opti-MEM medium (Life Technologies)

2. DNA plasmids 3. Electroporator suitable for mammalian cells Gene Pulser Xcell (Bio-Rad Laboratories) 4. Electroporation cuvettes (4 mm) (Bio-Rad) 5. Ficoll-Hypaque high-density medium (Sigma) 6. RPMI 1640 (Gibco) supplemented with 10% FBS

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2.6. Staining of Living Cells with Fluorescently Labeled Primary Antibodies or Fluorescent Probes

1. A kit to stably couple antibodies with fluorescent tags (Invitrogen) 2. Alternatively use a Zenon kit to directly stain primary antibody supernatant noncovalently (Invitrogen)

2.6.1. Fluorescently Labeled Antibodies 2.6.2. Cell Labeling with Fluorescent Probes

1. CMAC (blue), BCECF (green), CMTMR (red), or another permeable fluorescent probe with an ester bond that stabilizes the probe in the cytoplasm (nonspecific probes or organelle-specific ones such as mitotracker, lysotracker) (Invitrogen) 2. HBSS supplemented with 0.5% BSA

2.7. Time-Lapse Fluorescence Microscopy

1. Glass-bottomed Petri dishes 2. Laser scanning confocal fluorescence microscope or widefield epifluorescence microscope equipped with a piezoelectric focusing system that allows z-axis sectioning 3. Incubation system (La-con GBr Pe-con GmbH) 4. A parallel flow chamber (Glycotech) 5. A programmable syringe pump (Harvard Apparatus) 6. Glass-bottomed Petri dishes (WillCo Wells) 7. Phenol red-free medium

3. Methods Methods for visualizing interactions between leukocyte and the endothelium at the subcellular level aim to mimic as far as possible the in vivo situation. The molecular behavior observed in the in vitro models is very much dependent on the flow rate used, the receptor expression profile and migratory capability of the particular leukocyte subset, and the activation state of the endothelium. Primary cells are the best choice since immortalized cell lines do not usually upregulate adhesion receptors to the same levels as primary cells and may become partially dedifferentiated, losing some specific endothelial or leukocyte markers. However, to investigate a specific event (rolling, adhesion, locomotion, transmigration), transfected cell lines can be a valuable tool. In addition, static adhesion experiments can give information on the molecular dynamics or signaling cascades triggered

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by intercellular contact; however, details on the reorganization of some receptors, relevance of avidity processes, etc., are only unveiled under shear flow conditions. 3.1. Cell Models

1. HUVEC are most commonly used as a source of human primary macrovascular endothelial cells. Alternatively, primary cultures can be obtained of microvascular, lymphatic, blood– brain barrier, and high endothelial venule cells. 2. HUVEC are activated by changing to medium 199 10% FBS supplemented with 20 ng/ml TNF-α for 20 h (see Note 1). 3. PBMCs are obtained from peripheral blood donated by healthy volunteers. The PBMCs are isolated by centrifuging the blood for 30 min at 1,800 rpm on a Ficoll gradient. The isolated cells are then washed thoroughly and maintained in RPMI 1640 medium 10% FBS. For a crude preparation of PBLs, the PBMC population is depleted of monocytes by adhesion to a plastic flask for 30 min at 37ºC. 4. T lymphoblasts are derived from PBLs by activation for 48 h with PHA-L (1 μg/ml). After extensive washing, T cells are cultured for 7–14 days in RPMI 1640, 10% FBS containing 50 U/ml rhIL-2. 5. K562 transfectants (expressing α4β1 or αLβ2 integrins for VCAM-1 or ICAM-1 independent binding, respectively, see Note 2) are grown in RPMI 1640, 10% FBS, 1 mM G418.

3.2. Static Adhesion Measurements and Staining 3.2.1. Static Adhesion Measurements

1. To quantify leukocyte adhesion to an endothelial monolayer under static conditions, HUVEC are grown to confluence in 96-well plates and treated with TNF-α (for activation, see Note 1) in combination with the inhibitors to be tested (see Note 3). 2. Leukocytes are labeled with a fluorescent probe (usually BCECF-AM) as described in Section 3.6.2. Cells are usually pretreated for 15 min with blocking antibodies (approximately 10 μg/ml). For experiments with chemical inhibitors, the appropriate incubation times and doses should be determined in each case (see Note 4). 3. 105 cells/well in HBSS+1% BSA are placed in each HUVEC-coated well and incubated for 15–30 min at 37ºC (see Note 5). – Wells are filled with HBSS, sealed with an adhesive plastic seal, and maintained in an inverted position for 20 min. – Buffer is removed and cells lysed in 0.1% SDS, Tris 50 mM pH 8.5. Fluorescence is measured in a fluorescent microplate reader. (If the fluorescent probe is BCECF-AM, the excitation and emission wavelengths are 488 and 520 nm, respectively, see Note 6.)

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3.2.2. Static Adhesion Staining

– For staining of leukocyte interactions with endothelial cells under static conditions, a confluent monolayer of HUVEC is grown on a gelatin- or FN-coated coverslip (see Notes 7 and 8) and activated with TNF-α (Note 1) (Fig. 2.1).

Fig. 2.1. Visualization of leukocyte–endothelial interactions under static conditions by confocal microscopy. HUVEC were transfected with a fluorescent membrane protein and activated with 20 ng/ml TNF-α for 20 h. α4 integrin K562 transfectants were allowed to adhere to the endothelial monolayer under static conditions. Samples were fixed and analyzed by confocal microscopy. Endothelial cells extend filopodial projections around the adherent leukocyte in what has been called docking structure.6 Confocal optical sections can be used to create three-dimensional reconstructions and rotations of the complete cell volume, as shown in the side-view image.

– Leukocytes are allowed to adhere for different times in complete medium at 37ºC (see Note 5). – Medium is removed and samples are fixed with 2% paraformaldehyde in PBS for 5 min and extensively washed with TBS (see Note 9). – Samples are then stained with the appropriate specific antibodies. 3.3. Detachment Experiments

Experiments to investigate the detachment of leukocytes from the apical surface of endothelial monolayers are performed in a parallel flow chamber. The parallel-plate flow chamber used for leukocyte adhesion and transmigration under defined laminar flow is described in detail on the Glycotech web site (http://www.glycotech.com/apparatus/parallel.html). These assays measure the resistance of leukocyte–endothelial adhesion to increasing flow stresses. – HUVEC are grown in an FN-coated glass-bottomed petri dish and activated with TNF-α (see Notes 1, 8, 10, and 11). – Leukocytes (PBL or K562 transfectants) are allowed to adhere under static conditions, in complete medium, for 15 min at 37ºC (see Note 5).

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– The flow chamber is carefully mounted in the petri dish (see Note 12). HBSS+2% FBS (37ºC) is pulled through the flow chamber with a programmable syringe pump at an initial flow rate of 1 dyn/cm2 , which is increased up to 30 dyn/cm2 at 30 s or 1 min intervals (see Note 13). In the final seconds of each flow rate interval, six to eight 20× fields of view are recorded (see Note 14). Cell detachment is measured from the differences in the numbers of adherent cells after each flow rate interval (see Note 15). 3.4. Leukocyte– Endothelial Interactions Under Flow Conditions

The study of leukocyte–endothelium interactions under shear stress allows quantification of several functional parameters, such as rolling velocity, locomotion rate or percentage of rolling, detachment, adhesion, and transmigration (Fig. 2.2 and Supplemental Video 1).

Fig. 2.2. Leukocyte tracking under flow conditions by time-lapse confocal microscopy. A HUVEC monolayer was activated with 20 ng/ml TNF-α for 20 h. PBLs were perfused at 1.8 dyn/cm2 for 3–4 min and then cell-free HBSS buffer containing 2% FBS was perfused for the rest of the experiment. Bright-field images were acquired every 30 s over a period of 16 min (see Supplemental Video 1). The figure shows four representative frames from the video sequence. Each leukocyte was assigned a letter code denoting its migratory state: R for rolling, A for adhesion, L for locomotion, D for detachment. No cells transmigrated during this experiment. Cellular traks depicting the path followed by each cell during the whole experiment are overlayed in the last image.

1. PBLs (106 per ml) in HBSS 2% FBS at 37◦ C are drawn across TNF-α-activated confluent monolayers (see Note 1) at an estimated wall shear stress of 1.8 dyn/cm2 (see Note 16) for perfusion times from 30 s to 10 min (see Note 17). 2. Lymphocyte rolling on the endothelium is easily visualized because the adhered cells travel more slowly than freeflowing cells (Fig. 2.2R). Rolling velocity, frequency, and accumulation can be calculated after the experiment with the use of dedicated software. 3. Lymphocytes are considered to be adherent after 20 s of stable contact with the monolayer (Fig. 2.2A). 4. To track leukocytes that move on the apical endothelial surface (locomotion, Fig. 2.2L) in search of a suitable site for transmigration, time-lapse recording can be more illustrative of the process (8).

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5. Transmigrating lymphocytes can be distinguished because of their polarized morphology and changes in brightness (see Note 18). 6. Transmigrated lymphocytes are detected beneath the endothelial monolayer. 7. Lymphocytes are considered to be detached when they have returned to the free-flowing state after having been completely arrested on endothelium (Fig. 2.2D). 8. The number of rolling, adhered, transmigrating, transmigrated, and detached cells is quantified by direct visualization of four different fields (20× phase contrast objective) for each time point of every independent experiment (9) (see Note 19). When a specific parameter is to be calculated (rolling velocity, locomotion distance, etc.), the use of a higher magnification objective is recommended (40×–60×). 3.5. Transfection Procedures for Fluorescently Tagged Fusion Proteins

3.5.1. Transfection of Endothelial Cells

To visualize molecular dynamics, a common approach is to transfect cells with proteins labeled with fluorescent tags (EGFP, YFP, CFP, RFP, etc.) (Figs. 2.1 and 2.3). It is important to confirm that the fusion proteins have the same subcellular localization and function as the endogenous protein (see Note 20). The relatively easy protocols described below are considered to provide a rapid transient expression in a reasonable percentage of cells. Alternatives such as nucleofection or viral vector transduction can also be used. 1. HUVEC are trypsinized and resuspended in complete 199 medium to a concentration of 1.5 × 106 cells in a final volume of 200 μl. Cells are placed in a electroporation cuvette (4 mm) 2. 5 μl of 1.5 M NaCl is added (see Note 21)

Fig. 2.3. Study of molecular dynamics during leukocyte–endothelium interactions under shear flow by time-lapse fluorescence confocal microscopy. The T-lymphoblastic cell line CEM was transfected with a fluorescent cytoplasmic marker, while HUVEC were co-transfected with a fluorescent membrane protein and a intracellular marker. Endothelial cells were treated with 20 ng/ml TNF-α for 20 h before microscopy observation. Lymphocytes were perfused at 1.8 dyn/cm2 for 3–4 min and then cell-free HBSS buffer containing 2% FBS was perfused for the rest of the experiment. Fluorescent signals and bright-field images were acquired sequentially through a z-stack. A representative time point is shown, and each frame shows the maximal projection of the whole fluorescence stack for the channel. The best focused bright-field image was selected for the corresponding time point. (see Supplemental Video 2).

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3. 20 μg of plasmid DNA is added (see Notes 22 and 23) 4. Cells are electroporated at 200 V and 975 μF using an electroporator suitable for mammalian cells 5. The cuvette is filled with 800 μl fresh complete medium 199 (see Note 24) 6. Cells are seeded as droplets onto several glass-bottomed Petri dishes previously coated with 20 μg/ml FN (Notes 8 and 11) 7. Cells are allowed to adhere before filling the Petri dishes with complete medium supplemented with growth factors 8. Cells are grown for 24–48 h (see Notes 25 and 26) 3.5.2. Transfection of Leukocytes

1. Lymphoid cell lines are transfected in serum-free medium (Opti-MEM). Resuspend 10 × 106 cells in 400 μl OptiMEM 2. 20 μg plasmid DNA is added (see Notes 22 and 23) 3. Electroporation is performed at 250–280 V, 1,200 μF using an electroporator suitable for mammalian cells (see Note 24) 4. Cells are incubated in a final volume of 5 ml for 12 h, and dead cells are removed by centrifugation on a Ficoll gradient (see Note 26).

3.6. Staining of Living Cells with Fluorescently Labeled Primary Antibodies or Fluorescent Probes 3.6.1. Fluorescently Labeled Antibodies

An alternative to transfection with fluorescent protein constructs is staining of endothelial cells or leukocytes, either with neutral primary antibodies (see Note 27) directly coupled to fluorescent tags or with intracellular fluorescent probes.

1. The antibodies can be stably coupled to fluorescent tags using an appropriate kit (for example, Invitrogen) following the manufacturer’s instructions. 2. Alternatively, antibodies can be transiently tagged with a secondary Fab antibody already fluorescently labeled (Zenon kit, Invitrogen). This labeling procedure is very easy, rapid, and quite useful for short-term experiments in living cells (see Note 28). 3. Briefly, purified antibody or supernatant is incubated with the fluorescent Fab anti-mouse. 4. The reaction is stopped by adding an excess of mouse immunoglobulins. 5. The mixture is incubated with cells for 5–10 min and then washed to remove all free mouse Ig.

3.6.2. Cell Labeling with Fluorescent Probes

If the aim is not to label specific molecules but instead simply to label cells, cells can be loaded with intracellular fluorescent

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probes immediately before microscopy observation. Appropriate cell probes, with characteristic excitation and emission frequencies, can be combined with fluorescent proteins or antibodies. Usually these permeable probes contain an ester modification that is cleaved once they traverse the cell plasma membrane so that they are retained intracellularly. There are also probes that specifically label mitochondria, lysosomes, etc. 1. Leukocytes are washed with serum-free medium and resuspended at 5–10 × 106 cells/ml in serum-free medium containing 1 μM fluorescent probe. 2. Cells are incubated for 15 min at 37ºC, centrifuged, and washed to remove excess fluorescent probe (see Note 29). 3. For EC labeling, probes can be dissolved in serum-free medium or HBSS+0.5% BSA and added directly to the confluent monolayer. 3.7. Time-Lapse Fluorescence Microscopy

1. HUVEC transfected or not with fluorescent fusion proteins and/or labeled with antibodies or probes are grown to confluence on glass-bottomed dishes precoated with FN (20 μg/ml) (see Note 24) (Fig. 2.3 and Supplemental Video 2). Cells are then activated with TNF-α for the appropriate time (see Notes 1 and 11), transferred to phenol red-free medium, and placed on the microscope stage (see Note 30). 2. For experiments under static conditions, K562 transfectants or leukocytes resuspended in 500 μl of complete medium 199 are added. Labeling with fluorescent probes can facilitate observation of intercellular contacts with the endothelial monolayer. 3. Plates are maintained at 37◦ C in a 5% CO2 atmosphere using an incubation system. Alternatively, the parallel flow chamber can be coupled to the microscope stage for experiments under flow conditions. 4. Series of transmitted light and confocal or widefield fluorescence images, distanced 0.4–1 μm in the z-axis, are continuously obtained using a 40× or a 63× oil immersion objective (see Note 31). Images can be processed and assembled into movies using dedicated software (see Note 32).

4. Notes 1. Standard TNF-α treatment is 20 ng/ml for 20 h because the expression of ICAM-1 and VCAM-1 is maximal at this time. However, for maximal expression of E-selectin,

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treatment with 20 ng/ml TNF-α for 4 h would be optimal. Alternatively, leukocyte capture under flow can be achieved with a combination of a suboptimal TNF-α dose and the addition of exogenous chemokines such as SDF-1, which get immobilized at the apical glycosaminoglycans of the endothelium (10). Other proinflammatory cytokines, such as IL-1β, are also commonly employed. 2. Adhesion of K562 LFA-1 transfectants needs to be done in the presence of 1 mM Mn2+ to achieve full integrin activation. 3. If inhibitors of HUVEC function are to be tested, it is essential to measure ICAM-1 and VCAM-1 induction by flow cytometry in a parallel sample. 4. The use of allosteric inhibitors to specifically inhibit either VLA-4 (BIO5192 (Biogen Idec; Cambridge, MA)) or LFA-1 (BIRT377 (Boehringer-Ingelheim Pharmaceuticals; Ridgefield, CT)) is explained as an example. Integrin inhibitors (BIO5192 (10 μg/ml) or BIRT377 (10 μM)) are added to leukocytes 5 min before adhesion assay under static or flow conditions. 5. Adhesion times are very dependent on the leukocyte transmigratory capacity. For example, for PBLs, average adhesion times might be approximately 10 min, whereas K562 can be allowed to interact with the endothelial monolayer for more extended periods, since they do not transmigrate across the EC monolayer. 6. To provide the 100% adhesion reference, a separate well is loaded with the total input of labeled leukocytes and directly lysed. 7. 24-well plates and 12-mm coverslips are suitable for routine staining. 8. Coverslips can be coated with 1% gelatin, 20 μg/ml FN, or other ECM preparations. With gelatin coating, better confluence is achieved if the gelatin is fixed with 0.5% glutaraldehyde and extensively washed with TBS before seeding the EC. 9. Wash aldehydes with Tris-containing saline buffer to block the fixation reaction with an excess of amine groups. Alternatively, glycine solutions can be used. 10. The results are more easily quantified if the transmigration rate is low: use K562 transfectants, which do not transmigrate, or PBLs, rather than T lymphoblasts or neutrophils. 11. Since the observation area is usually limited (the center of the coverslip in the flow gasket or just a few fields in a motorized confocal time-lapse microscope), it is not

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necessary to seed large area with HUVEC. HUVEC can be conveniently seeded as a droplet, which also favors immediate confluence. Once HUVEC have spread, cells are activated with TNF-α. 12. It is important to ensure that the coverslip does not dry at any moment and to avoid bubbles. 13. An initial wash at 1 dyn/cm2 for 1 min can be performed to remove unbound cells. Alternatively, adhered cells are counted only after the first 2 dyn/cm2 interval. 14. It is convenient to record the same fields each interval, especially if initial adhesion is not homogeneous on the preparation. 15. If the incidence of transmigration is high during the experiment, transmigrated cells in each field have to be counted and subtracted from the total count to yield the number of detached cells. 16. The flow rate of 1.8 dyn/cm2 is similar to the shear force generated in the human postcapillary venules. 17. When using extracellular blockers (such as antibodies or peptides) under flow conditions, you should assess that the perfusion with buffer does not wash them away significantly. 18. A phase contrast objective is useful for direct observation of changes in leukocyte morphology during extravasation because the initially bright round cells on top of the endothelium spread and darken while migrating across and beneath the monolayer. 19. Coverslips can be immediately fixed under flow with 4% PFA at room temperature for 10 min and then washed with TBS and stained for markers of interest. 20. If antibodies against the protein of interest are available, it is convenient to confirm that they recognize the fusion protein. The molecular weight of the expressed fusion protein should also be assessed by Western blot to exclude the presence of partial degradation products, which might give false subcellular localizations. 21. Efficiency of electroporation has been shown to increase with higher osmotic strength. 22. Try to use DNA constructs at as high a concentration as possible to reduce the volume to be added to the cell suspension. 23. In co-transfection experiments, the amount of total DNA should not exceed 20 μg/ml; in making adjustments,

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decrease the amount of plasmid encoding the protein that is more efficiently expressed. 24. After electroporation, allow cells to recover from electric shock and close micropores prior to seeding. 25. Allow cells to grow after electroporation, but ideally for no more than 24 h; this will ensure the highest possible fluorescent protein content in the culture. 26. Transfection efficiency can be routinely quantified by direct flow cytometry. 27. The fluorescently tagged primary antibodies should be functionally neutral in order not to interfere with cell functions. 28. The Zenon kit is not advisable for long-term staining since the tag is not covalently bound to the primary antibody and can become detached with time, decreasing specific signal and increasing the background signal. 29. Usually cell labeling is evident since the cell pellet will be colored. 30. The use of medium without phenol red is important to avoid autofluorescence. 31. Specific acquisition conditions (in terms of scanning velocity, zoom, photomultiplier gain, offset, number of series, time-lapse parameters, etc.) need to be adjusted according to the fluorescence intensity of the samples, the cell types, and cell processes being studied. Photobleaching and phototoxicity also need to be minimized in each experimental setup. 32. Images acquired using a confocal microscope do not need further processing, but deconvolution is required prior to analysis of images acquired with a widefield fluorescence microscope.

Acknowledgments This work was supported by grants BFU2005-08435/BMC from the Ministerio de Educación y Ciencia, and European Network MAIN LSHG-CT-2003-502935 to FSM, by ContratoInvestigador FIS 0019 from Instituto de Salud Carlos III to MY-M. The authors thank Giulia Morlino and Francesc Baixauli for providing samples for the time-lapse experiments and Simon Bartlett for editing the manuscript.

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References 1. Ley K, Laudanna C, Cybulsky MI, Nourshargh S. (2007) Getting to the site of inflammation: the leukocyte adhesion cascade updated. Nat Rev Immunol 7, 678–89. 2. Ley K, Kansas GS. (2004) Selectins in T-cell recruitment to non-lymphoid tissues and sites of inflammation. Nat Rev Immunol 4, 325–35. 3. Laudanna C, Alon R. Right on the spot. (2006) Chemokine triggering of integrin-mediated arrest of rolling leukocytes. Thromb Haemost 95, 5–11. 4. Sanchez-Madrid F, del Pozo MA. (1999) Leukocyte polarization in cell migration and immune interactions. EMBO J 18, 501–11. 5. Barreiro O, de la Fuente H, Mittelbrunn M, Sanchez-Madrid F. (2007) Functional insights on the polarized redistribution of leukocyte integrins and their ligands during leukocyte migration and immune interactions. Immunol Rev 218, 147–64.

6. Barreiro O, Yáñez-Mó M, Serrador JM, et al. (2002) Dynamic interaction of VCAM-1 and ICAM-1 with moesin and ezrin in a novel endothelial docking structure for adherent leukocytes. J Cell Biol 157, 1233–45. 7. Vestweber D. (2007) Adhesion and signaling molecules controlling the transmigration of leukocytes through endothelium. Immunol Rev 218, 178–96. 8. Shulman Z, Pasvolsky R, Woolf E, et al. (2006) DOCK2 regulates chemokinetriggered lateral lymphocyte motility but not transendothelial migration. Blood 108, 2150–8. 9. Goetz DJ, Greif DM, Shen J, Luscinskas FW. (1999) Cell-cell adhesive interactions in an in vitro flow chamber. Methods Mol Biol 96, 137–45. 10. Cinamon G, Alon R. (2004) Real-time in vitro assay for studying chemoattractanttriggered leukocyte transendothelial migration under physiological flow conditions. Methods Mol Biol 239, 233–42.

Chapter 3 Leucocyte Adhesion Under Haemodynamic Flow Conditions Charlotte Lawson, Marlene Rose, and Sabine Wolf Abstract Vascular endothelial cells (EC) line the luminal side of all blood vessels and act as a selective barrier between blood and tissue. EC are constantly exposed to biochemical and biomechanical stimuli from the blood and underlying tissue. Fluid shear stress acts in parallel to the vessel wall, resulting from friction of blood against EC. Despite the importance of flow on normal EC function, much of the information regarding EC function and dysfunction has been derived from cells harvested, grown and studied in static culture. In order to study the effects of shear stress on EC function, a number of in vitro models have been developed. This chapter provides methodology for use of a system which enables recirculation of leucocytes and cell culture medium over the endothelium for a period of several minutes to days and enables investigation of the effects of prolonged leucocyte co-culture on both the endothelial and leucocyte populations. Key words: Endothelium, shear stress, parallel-plate flow chamber.

1. Introduction 1.1. The Endothelium

Vascular endothelial cells (EC) line the luminal side of all blood vessels and act as a selective barrier between blood and tissue. EC are constantly exposed to biochemical and biomechanical stimuli from the blood and underlying tissue. It is well established that maintenance of a quiescent endothelium is vital to prevent coagulation and control vascular permeability as well as regulating vascular tone through production of nitric oxide. In addition, EC contribute to maintenance of the quiescence of circulating leucocytes (reviewed in (1)). Conversely, failure to control vascular permeability and coagulation, an increase in vascular tone and loss of

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leucocyte quiescence can all contribute to EC dysfunction. Thus EC pathology contributes to many conditions such as atherosclerosis, hypertension, thrombosis, stroke, vasospastic disorders and diabetic microangiopathy, as well as the increase in mortality and morbidity associated with chronic inflammation and autoimmune disease (review in, e.g., (2)). There is increasing evidence that endothelial cells are also bona fide antigen presenting cells (APC). In vitro, they present antigen to B7-independent memory T cells inducing proliferation and IL-2 production. In vivo, human endothelium is constitutively positive for major histocompatibility (MHC) class II and vascular structures can be identified by HLA-DR staining in normal tissue sections (3–6). 1.2. Forces on the Endothelium: Shear Stress

Blood vessels are constantly exposed to haemodynamic forces in the form of cyclic stretch, fluid shear stress and hydrostatic pressures. Shear stress is the major haemodynamic force EC respond to, whereas vascular SMC responses are more influenced by cyclic stretch (7, 8). Fluid shear stress acts in parallel to the vessel wall. It results from the friction of blood against the inner lining of the blood vessel wall and is principally sensed by EC (Fig. 3.1; (9)). In “linear”, unbranched areas of the vasculature, blood flows in uniform, laminar patterns and EC experience a mean positive shear stress, around 10–40 dyn/cm2 in the arterial network and 1–20 dyn/cm2 in the venous microcirculation (see Fig. 3.1). In areas A Laminar Flow

R

Blood viscosity, η

Volumetric flow rate, Q

B Disturbed Flow

Fig. 3.1. Diagram showing flow patterns for laminar flow (a) and disturbed flow (b) (adapted from (9)).

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with abrupt curvations or bifurcations, the steady laminar flow pattern is disrupted by regions of separated blood flow creating recirculating sites. Shear stress in these regions varies from negative, zero and positive values (Fig. 3.1; (10, 11)). Parts of the vasculature exposed to steady laminar flow with high shear stress are atheroprotective, whereas areas of turbulent, disturbed flow and low fluid shear stress are prone to develop atherosclerotic lesions (12, 13). 1.3. Atherosclerosis and the Endothelium

It is well established that when EC are subjected to disrupted flow, they take on an activated pro-inflammatory phenotype that supports leucocyte transendothelial migration in vitro and in vivo. Atherosclerotic lesions form at branch points in arteries where flow is not laminar and leucocyte accumulation is observed even in very early lesions, with accumulation of T cells as well as monocytes being well documented in humans (15–17) and in the ApoE–/– or LDL-R–/– mouse models of atherosclerosis (18, 19). The potential importance of T cells for progression of the lesions has been demonstrated using ApoE–/– /Rag-1 or LDL-R–/– /Rag-1 mice which are defective in both T and B cells, but not monocytes. Early lesion development in the Rag-1 mice, compared to wild types, was significantly diminished after 8 weeks on a Western-type diet (WTD), suggesting that lymphocytes play an active role in early lesion development (19). During chronic allograft vasculopathy (CAV), lesions are seen which are not dissimilar to those seen in atherosclerosis, although there are several features that are different (reviewed in detail elsewhere; (20)). As with native atherosclerosis, large accumulations of fibro-fatty deposits have been observed in the subendothelial space as well as proliferating smooth muscle cells that have migrated from the media of blood vessel wall. These VSMC secrete inflammatory cytokines and extracellular matrix proteins, all of which contribute to the progression of the lesion. As with “native” atherosclerosis, elevated numbers of leucocytes have been observed adhering to and transmigrating into the subendothelial space in both human and animal models including increased numbers of CD4 T cells even in the presence of an intact endothelium in non-branching parts of the vasculature where shear stress is high (21, 22).

1.4. Use of Endothelial Cells In Vitro

Much of the information regarding EC function and dysfunction has been derived from cells harvested, grown and studied in culture. EC have been isolated from many different vascular beds and various species including humans. The most common approach for obtaining EC is by enzymatic digestion of cells from large blood vessels, which provides a good yield of high-purity cells. These can be further purified with magnetic bead separation

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and use of selective media (23). Human umbilical vein endothelial cells (HUVEC) are probably the most widely used model for human endothelia (24). 1.5. Use of a Parallel-Plate Flow Chamber

A disadvantage of using cultured cells is the difficulty in recapitulating the forces that EC are exposed to in vivo, in a culture dish. To some extent, this can be overcome by use of twodimensional “flow chambers”. A number of different apparatus have been described including the parallel-plate flow chamber (25) and the cone and plate viscometer (26), both of which have been shown to mimic the flows seen in vivo using a twodimensional/single-cell monolayer setting to enable molecular dissection of the responses due to EC alone. Here, we outline protocols using a parallel-plate flow chamber for long-term exposure of cultured EC to arterial flow conditions, in the presence of purified T-cell populations.

2. Materials 2.1. HUVEC Culture on Glass Slides

1. HUVEC culture medium: Medium 199 with HEPES (PAA) supplemented with 20% foetal bovine serum (BioSera) and L -glutamine (sigma) and penicillin/streptomycin (PAA). 2. HUVEC flow medium: M199 with HEPES supplemented with 10% FCS; L-glutamine, penicillin/streptomycin; amphotericin B (PAA). 3. Sterile 1x PBS (10x PBS; for 1 L add 2 g KCl, 2 g KH2 PO4 , 80 g NaCl, 11.5 g Na2 HPO4, dilute to 1x with ddH2 O and autoclave before use). 4. 1x trypsin/EDTA (PAA). 5. Glass microscope slides (76 × 38 mm; Fisher Life Sciences) [sterilise by autoclaving before use]. 6. Sterile 9 cm Petri dishes. 7. Human fibronectin (Sigma) diluted to 50 μg/ml in 1x PBS. 8. Haemocytometer (e.g. Fisher LifeSciences). 9. Trypan Blue (Sigma).

2.2. T-Cell Purification

1. 15% EDTA for blood collection. 2. 1x PBS supplemented with 2% FBS and 1 mM EDTA. 3. RosetteSep human CD4 T-cell-negative selection cocktail (Stemcell Technologies Inc.). Store at 4◦ C. 4. Histopaque 1,077 cell separation gradient (Sigma). 5. Sterile pastettes (Greiner Bio-One).

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6. RPMI (PAA) supplemented with penicillin/streptomycin, L -glutamine, 10% FCS (T-cell medium). 7. Haemocytometer. 2.3. Flow Loop

1. Flow chamber and apparatus for recirculating flow loop from Cytodyne Inc. (www.cytodyne.net). 2. HUVEC flow medium. 3. For total RNA extraction; TRIzol reagent (Invitrogen) (this will require further reagents including chloroform, isopropanol, 70% ethanol, RNase-free pipette tips and tubes, RNase-free water). 4. For protein extraction; RIPA buffer (20 mM MOPS, pH 7.0; 150 mM NaCl; 1 mM EDTA; 1% NP40; 1% Na deoxycholate; 0.1% SDS), protease and phosphatase inhibitor cocktails (Sigma P2714, P5726), tray containing ice, 1 ml syringes and 21-G needles, microcentrifuge (ideally cooled).

2.4. Immunohistochemistry

1. Ice-cold acetone. (Ensure that acetone is only stored in spark-proof freezers. If this is not available, pre-cool on ice before use.) 2. 100-ml beakers to hold oversized glass slides. (Slides used in the Cytodyne setup described below do not fit in standard Coplin/staining jars.) 3. 1x PBS. 4. Primary antibodies as appropriate (e.g. against CD31; DAKO). 5. Fluorescently conjugated secondary antibodies (e.g. goatanti-mouse-Ig-Alexa 594; Invitrogen). 6. Phalloidin-Alexa 488 (Invitrogen). 7. VectaMount with DAPI (VectorLab). 8. 22 × 50 mm coverslips (e.g. VWR).

2.5. T-Cell Alloproliferation Assay

1. RPMI T-cell medium. 2. Second HUVEC isolate. 3. 5(6)-Carboxyfluorescein diacetate N-succinimidyl ester (CFSE; Sigma) diluted to 1 μM. 4. Flat-bottomed 24-well tissue culture plate.

2.6. PHA Proliferation Assay

1. RPMI T-cell medium (27). 2. Lectin from Phaseolus vulgaris (phytohaemagglutinin; PHA; Sigma) diluted to 2 μg/ml. 3. γ-Irradiation source to prevent division of antigen presenting cells. (It is possible to use 60 μg/ml mitomycin C for 25 min [Sigma] if a suitable radioactive source is not available.)

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4. [3 H] thymidine ([3 H] TdR) (GEC or Sigma). 5. V-bottomed 96-well tissue culture plate. 6. Cell harvester apparatus and β-counter.

3. Methods 3.1. Cell Culture 3.1.1. HUVEC Culture

HUVEC may be obtained from commercial sources (e.g. Promocell, Oxford, UK) or may be isolated from umbilical cords collected with appropriate ethical permission and informed consent from a local maternity unit, according to methods described in detail elsewhere (e.g. (23)) (see Notes 1 and 2). 1. Place 76 × 38 mm glass slides in 90 mm sterile Petri dishes and pretreat with 0.5 ml 50 μg/ml human fibronectin for 45 min at room temperature in a Class II safety cabinet. Then remove excess fibronectin using a sterile pipette. 2. Passage confluent HUVEC cultures following welldescribed protocols using trypsin/EDTA or the supplier’s recommended protocol. 3. Count live cells by Trypan Blue exclusion using a haemocytometer 4. Seed fibronectin-coated slides with approximately 2 × 106 per ml HUVEC in 1 ml of HUVEC medium onto each glass slide (see Note 3). 5. Incubate slides in a 37◦ C/5% CO2 incubator for at least 4 h to allow HUVEC to adhere and then flood slides with 12 ml of flow medium (M199 supplemented with L-glutamine, penicillin (100 units), streptomycin (0.1 mg/ml), 10% FBS and 1/200-dilution amphotericin B) and incubate overnight in 37◦ C/5% CO2 incubator.

3.1.2. T-Cell Purification

There are many protocols for purification of human CD4+ T cells from peripheral blood (28). Protocols employing negative selection are preferred to minimise activation of the T-cell population under examination. 1. Collect peripheral blood by venepuncture into tubes containing 15% EDTA (1 ml for every 50 ml blood collected). 2. Purify CD4+ T cells using method of choice (e.g. RosetteSep negative selection cocktail (Stemcell Technologies Inc.) followed by gradient separation on Histopaque 1,077 and collection of the buffy coat layer using sterile pastettes). 3. Wash purified T cells twice in 1x PBS/2% FBS/1 mM EDTA.

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4. Count purified T cells using haemocytometer and Trypan Blue exclusion to distinguish viable cells. 5. Verify purity by FACS analysis of a sample of cells after staining with fluorochrome-conjugated anti-CD3 and anti-CD4 antibodies. 6. For enumeration of adhered T cells, they may be labelled with CFSE by addition of 1 μl of 1 μM CFSE to 5 × 106 T cells in a volume of 5 ml; incubate in the dark for 5 min, then quench with 5 ml FBS and incubate for a further 5 min. Dilute to 50 ml with 1x PBS; pellet cells and resuspend as appropriate. Uptake of CFSE should be verified by fluorescent microscopy or FACS. 3.2. Use of Parallel-Plate Flow Chamber for Laminar Flow Experiments with HUVEC

A parallel-plate recirculating flow loop system as first described by Frangos (25) may be used for shear stress experiments carried out over a longer period of time to the traditional setups utilising a syringe pump to draw fluid over the parallel-plate flow chamber. The system consists of two reservoirs connected with a flow chamber (Fig. 3.2) to enable recirculation of the flow media and therefore the opportunity to acclimatise EC to flow conditions.

Fig. 3.2. Diagram of flow loop apparatus showing the flow chamber, silicon gasket and the glass slide with the attached confluent monolayer of endothelial cells, which are held together by a vacuum pump at the periphery of the chamber complex. The flow chamber has two slits through which flow medium enters and exits the channel. The (arterial) flow rate is controlled by the peristaltic pump. The medium is recirculated from the reservoir to the inlet tubing onto the flow chamber and back into the reservoir.

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The parallel-plate flow chamber consists of the flow chamber, a gasket and glass slide seeded with EC (Fig. 3.3), all of which are held in place by a vacuum pump. Additionally, the use of a vacuum pump ensures a uniform channel depth (d = 220 μm) across the flow chamber area (a = 16 cm2 ). Flow media is pumped by a peristaltic pump from the lower reservoir to the upper reservoir at a constant rate. Overflow of excess media drains down the glass tube and is collected into the lower reservoir where it can be recirculated. The design of two reservoirs prevents the entry of air bubbles into the primary flow section upstream of the flow chamber and allows the maintenance of a constant hydrostatic pressure head between the upper and the lower reservoir. Flow media enters the flow chamber via the entry port. It passes through the entry slit, over the channel where cells are located, into the exit slit and leaves the flow chamber via the exit port (Fig. 3.3). The flow media is then returned to the lower reservoir for recirculation.

E

C

G

F D H

B

A Fig. 3.3. Cartoon showing parallel flow chamber. When assembling the flow chamber, the gasket (b) is carefully placed onto the flow chamber (c). The glass slide (a), which is coated with HUVEC, is added on top of the gasket with the cells facing towards the flow chamber. A vacuum pump is attached onto the flow chamber (d) to hold glass slides, gasket and flow chamber in place. Media (grey arrows) enters the flow chamber via the entry port (e), runs through the slit (f) over the channel back into the slit (g) and exits the flow chamber through exit port (h). When aligning the gasket, great care is required not to cover the entry (f) and exit slit (g) which would prevent flow of the culture medium.

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The use of a recirculating system allows for longer term culture of EC under laminar flow conditions (up to 96 h in our laboratory) without use of large volumes of cell culture media, and inclusion of a septum port in the lower reservoir enables collection of samples of flow media for analysis of soluble factors released by EC at different time points using appropriate assays. 3.2.1. To Set Up the Apparatus

1. Sterilise parallel-plate flow loop system using ethylene oxide. 2. Pre-warm sterile flow media and warm the 37◦ C chamber and apparatus for at least 1 h before intended use 3. Assemble parallel-plate flow chamber and flow loop according to Fig. 3.2 inside a class II safety cabinet to maintain sterility (see Note 4) 4. Ensure that the connectors on tubing (e.g. Masterflex tubing and connectors from Cole Parmer, London, UK) and on glassware are firmly attached and close the flow loop 5. Add pre-warmed flow media to the bottom reservoir via the three-way tap. 6. Align the sterile gasket (Fig. 3.3b) onto the flow chamber (Fig. 3.3c) in the tissue culture hood being careful not to cover the channel and slits (Fig. 3.3f, g) on the parallelplate flow chamber. 7. The slide (Fig. 3.3a), seeded with a confluent monolayer of HUVEC, can then be mounted onto the gasket on the flow chamber and attached immediately to the vacuum pump (Fig. 3.3d) to hold the flow chamber together. Ensure great care is taken not to move the gasket and glass slide on the flow chamber out of place during the process in order to avoid leakage. 8. After attaching the inlet (Fig. 3.3e) and outlet tubing (Fig. 3.3h) to the flow chamber, carefully move the flow loop apparatus to a pre-warmed 37◦ C incubator (Note 5). 9. Place tubing onto the peristaltic pump and observe system for signs of leakage. During assembly of the flow loop, tubing and/or flow chamber itself should be adjusted to ensure the optimal flow loop conditions, i.e. no hindrance of flow and no air bubbles are trapped in the flow chamber. Air bubbles can be removed using a needle and syringe placed in the septum port located at the exit of the chamber. 10. Level of flow media remaining in the reservoir should be observed to ensure that no leakage has occurred.

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11. After switching on the peristaltic pump, HUVEC on the slide are immediately exposed to a laminar shear stress of >10 dyn/cm2 for 1–96 h. 12. Before co-culture of T lymphocytes with HUVEC, allow HUVEC to become accustomed to the presence of laminar shear stress for at least 18 h (see Note 6). Positioning of a phase contrast microscope within the incubator before commencement of flow will allow visualisation of slides whilst they are being subjected to flow to ensure the presence of an intact monolayer, with cells aligned to the direction of the flow. 13. If desired, carefully wash the HUVEC monolayer whilst maintaining flow by removal of excess flow medium in the lower reservoir and replacement with fresh flow medium (via the septum port). 14. For prolonged co-culture (up to 4 h) of purified CD4+ T cells and HUVEC, resuspend T cells in HUVEC flow media at 1 × 107 per ml and inject 2–5 × 106 into the flow loop via the septum port located in the lower reservoir. 15. At the termination of flow, slides should be quickly removed from the chamber and processed for analysis using immunohistochemistry (whole slides), flow cytometry (intact cells), Western blotting (cell lysates) or PCR (mRNA), and flow media can be collected for measurement of soluble factors. 16. To harvest cells for flow cytometry, rinse slides briefly in 1x PBS and place in a clean dry Petri dish (see Note 7). Add a 1 ml “drop” of Accutase (PAA L11-007) to the top of the slide and incubate for 2–3 min at room temperature. Carefully remove Accutase containing disaggregated cells and place in a 15 ml conical tube containing 0.5 ml FBS. Wash the slide carefully with 2 ml PBS to collect any remaining cells. Pellet and process for flow cytometry. 17. To harvest cells for collection of total RNA, rinse slides briefly in 1x PBS and place in a clean dry Petri dish. Place 0.5 ml TRIzol reagent (Invitrogen) to the top of the slide and incubate 5 min at room temperature. Carefully collect the lysate into a clean 1.5 ml Eppendorf tube and follow manufacturer’s instructions for purification of total RNA. 18. To harvest cells for collection of protein lysates, rinse slides briefly in 1x PBS and place in a clean dry Petri dish. Place the Petri dish in a tray of ice and add 0.5 ml RIPA buffer to the top of the slide. Incubate 10 min on ice, then carefully scrape the lysate to loosen cellular material. Collect the lysate into a clean Eppendorf tube. Push the lysate through a syringe and small bore needle × 10, then centrifuge at

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13,000 rpm for 10 min to pellet nuclear debris. Keep supernatant and store at –80◦ C. 19. Use cells exposed to static culture conditions as controls for all experiments. 3.2.2. Immunohistochemistry

1. At termination of flow, remove glass slides from the flow loop or take out of static culture and quickly rinse in PBS. 2. HUVEC are fixed by submerging slides in ice-cold acetone for 5 min followed by 3 × 5 min washes in 1x PBS. 3. During the washes, prepare the primary antibody solution at the appropriate dilution and aliquot 250 μl onto a clean dry Petri dish. The glass slide is carefully placed on top of the antibody solution ensuring HUVEC on the slide are in contact with the diluted antibody. 4. Incubate for 30 min at room temperature, then remove slides from Petri dishes, rinse and wash three times with 1x PBS. 5. During the washes, prepare the secondary antibody conjugate (e.g. Alexa Fluor 594 goat anti-mouse IgG, Molecular Probe/Invitrogen, UK) at the appropriate dilution together with fluorescently conjugated phalloidin to stain F-actin, if appropriate (e.g. Alexa 488 conjugate, Molecular Probes/Invitrogen, UK). 6. Incubate the slides HUVEC side down in the antibody solution (250 μl) for 30 min at RT. 7. Rinse slides, then wash three times in PBS before carefully adding two drops of mounting medium containing the nuclear counterstain 4,6 diamidino-2-phenylindole (DAPI) for visualisation of cell nuclei. 8. Place long cover slips (Fisher Scientific) on top of the mounting media and store slides at 4◦ C in the dark under humid conditions to avoid drying out before visualisation by fluorescent/confocal microscopy (Supplementary Fig. 3.1).

3.2.3. Analysis of T-Cell Functionality After Co-culture with EC Under Laminar Flow Conditions: Alloproliferation to Third-Party HUVEC

1. Seed a 24-well tissue culture plate with 1 × 105 HUVEC/well in triplicate or quadruplicate using a separate isolate to the one used for co-culture under laminar flow conditions, 24 h before the flow co-culture. 2. During laminar flow co-culture (Section 3.2.1) remove medium from HUVEC in the 96-well plate and treat with 60 μg/ml mitomycin C for 25 min to prevent HUVEC proliferation followed by three washes with 1x PBS to remove all traces of mitomycin C. Replace medium with 250 μl T-cell medium/well.

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3. At termination of flow collect flow media from the lower reservoir and pellet T cells. 4. Count T cells using Trypan Blue to determine viability immediately after cessation of flow. 5. Label with CFSE as described in Section 3.1.2 and resuspend at 8 × 105 cells/ml. Add 250 μl/well (2 × 105 T cells/well). 6. Incubate at 37◦ C for 72–240 h then recover adherent and non-adherent T cells by collection of culture media followed by gentle washing of the monolayer with T-cell media. 7. Analyse by flow cytometry to determine the proliferation of CFSE-labelled T cells (Fig. 3.4).

Fig. 3.4. CD4 T-cell alloproliferation to third-party HUVEC at different times after culture with HUVEC at arterial flow rates. Purified human CD4+ T cells were co-cultured with HUVEC in a recirculating flow loop for times indicated. T cells were recovered from the apparatus, labelled with CFSE and co-cultured with a third-party HUVEC isolate that had been treated with mitomycin C to prevent cell division. T-cell division was estimated by flow cytometry at different time points after commencement of the second co-culture period. n = 2.

3.2.4. Analysis of T-Cell Functionality After Co-culture with EC Under Laminar Flow Conditions: PHA Proliferation Assay

1. During T-cell purification, prepare a small aliquot of peripheral blood mononuclear cells (PBMC; (28)) for use as antigen presenting cells and irradiate using a γ-irradiation source to prevent proliferation. (If no γ-irradiation source is available, it is possible to treat with mitomycin C as above.) 2. At termination of flow, collect flow media from the lower reservoir and pellet T cells. 3. Count T cells using Trypan Blue to determine viability immediately after cessation of flow. 4. Seed 96-well v-bottomed plates ± 2.5 × 104 irradiated PBMC; 2.5 × 104 T cells subjected to co-culture with EC under flow conditions; 2 μg/ml PHA, in triplicate or quadruplicate. 5. Incubate 48 h at 37◦ C, then add 1 μCi [3 H] thymidine and incubate for a further 18 h before harvesting of plates by freezing at –70◦ C followed by thawing and transfer of cell

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160000 140000 120000 cpm

100000 80000 60000 40000 20000 0 CD4 alone CD4 + PHA APC alone APC + PHA

no flow + 1h flow + 4h flow + APC + APC + APC + PHA PHA PHA

15h flow + APC + PHA

Fig. 3.5. CD4 T-cell proliferation to PHA after culture at arterial flow rates for increasing time points. Purified human CD4+ T cells were co-cultured with HUVEC in a recirculating flow loop for times indicated. T cells were recovered from the apparatus and cultured in the presence of PHA and PBMC that had been γ-irradiated to prevent cell division. [3 H] TdR was added for the last 18 h of culture, before harvesting of plates onto filter mats and analysis on a β-counter, n = 2.

lysates to filter mats before analysis using a β-counter (for a detailed protocol of this method, refer to user guide for your cell harvester or (28) (Fig. 3.5).

4. Further Uses of the Apparatus 1. Analysis of leucocyte adhesion T-cell adhesion to the endothelium can be monitored in real time by inclusion of a phase contrast microscope (with cooled CC camera attached) in the 37◦ C incubator housing the flow loop. Alternatively, T cells can be pre-labelled with CFSE prior to inclusion in the flow loop and can be detected either still attached to the endothelium by fluorescence microscopy after fixation of slides or can be quantified by flow cytometry after disaggregation of cells from the slide after flow has ceased. For short-term recirculation experiments the flow can be reduced to maximise T-cell capture. 2. Use of silicone steps to simulate separated flows It is possible to simulate separated flows by inclusion of a silicone (e.g. Sylgard, Sigma) or AralditeTM “step” close to the point of entry of flow medium in the flow chamber (Fig. 3.6). The step is made by painting on liquid silicone or Araldite and allowing at least 24 h to harden followed by sterilisation by autoclaving before seeding of HUVEC. The step should be no more than (height of gasket – 1) micrometres to enable flow to be maintained.

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A

Glassslide

EC monolayer Flow flow chamber

B Glass slide

flow chamber i ii iii

iv

Fig. 3.6. Interposition of a step barrier in the primary flow creates defined areas of disturbed flow downstream. (a) Flow in parallel-plate flow chamber. Laminar flow (black arrows) is created by pumping fluid over an endothelial monolayer plated onto a glass coverslip. (b) Interposition of a step barrier creates areas of disturbed flow downstream: (i) flow recirculation, (ii) flow reattachment, (iii) flow recovery and (iv) recovered laminar shear (adapted from (29)).

5. Notes 1. It is recommended that HUVEC are not used for experiments beyond the third passage after isolation from umbilical cords, as they may lose endothelial phenotype (Section 3.1.1). We recommend routine immunostaining and flow cytometric analysis for surface markers including CD31 (antibodies available from DAKO) to ensure that a pure population of endothelial cells has been obtained (all human endothelial cells should maintain CD31 positivity in culture). 2. Single isolates of HUVEC should be used if co-culture with allogeneic CD4+ T cells will be carried out. 3. Endothelial cells from other tissue beds/species may be cultured under laminar flow conditions. It is important to ensure that the cells are able to adhere to the glass microscope slides and the use of collagen, gelatine or poly-L-lysine may be required in addition to fibronectin to improve adhesion to the slides. 4. It is vital to ensure that the apparatus remains sterile during assembly to ensure that there is no fungal or bacterial contamination during prolonged periods of culture at 37◦ C (Section 3.2.1 Step 3). Therefore, it is recommended that sterile surgical gloves are used whilst assembling the flow apparatus or that non-sterile gloves are sprayed with 70% ethanol before beginning assembly and that all instruments (such as forceps) are sterilised before use.

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5. If the safety cabinet is not situated in very close proximity to the 37◦ C incubator, it is recommended that all parts of the apparatus (slide with monolayer of HUVEC, silicone gasket and chamber; flow loop and tubing) are assembled in the safety cabinet, but connected in the incubator (Section 3.2.1 Step 8). It is also recommended that the flow medium is added once the apparatus is fully assembled in the incubator. Although this may compromise sterility to some degree, it is likely to be achieved more efficiently and with less likelihood of spillage or disconnection of the component parts of the apparatus and therefore overall be less likely to result in contamination of the apparatus. It is recommended that a “dry run” is carried out to determine the most efficient way to transport the assembled chamber and flow loop across the laboratory in the first instance! 6. It is important to acclimatise HUVEC to the flow conditions before starting the experimental protocol as they undergo phenotypic changes during acute exposure to flow compared with longer periods of exposure (Section 3.2.1, Step 12). Within 1 h of commencement of flow, the cells will take on an activated phenotype with expression of enhanced levels of adhesion molecules including VCAM-1 and MHC class I (unpublished observations). However, after several hours’ exposure, HUVEC become quiescent with low MHC class I expression and no expression of VCAM-1. Cells also express the transcription factor KLF-2, which is characteristic of quiescent endothelial cells subjected to laminar flow. 7. When harvesting whole cells, proteins or RNA from microscope slides, it is important to use a dry Petri dish to avoid escape of the medium used to harvest cells (Accutase, RIPA buffer, TRIzol, etc.) onto a wet dish or a piece of tissue paper, which would adversely affect the final yield (Section 3.2.1, Steps 16–18).

Supplementary Fig. 3.1. HUVEC were cultured in static (a) or arterial flow (b) conditions for 24 h before fixation of slides in ice-cold acetone and staining for actin stress fibre formation with phalloidin-Alexa 488 (top right panel, green). Nuclei were stained with DAPI (top left panel, blue), (Bottom left panel, overlay). Representative images from n > 5 experiments.

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References 1. Pober JS, Sessa WC. (2007) Evolving functions of endothelial cells in inflammation. Nat Rev 7, 803–15. 2. Lusis AJ. (2000) Atherosclerosis. Nature 407, 233–41. 3. Choo JK, Seebach JD, Nickeleit V, et al. (1997) Species differences in the expression of major histocompatibility complex class II antigens on coronary artery endothelium: implications for cell-mediated xenoreactivity. Transplantation 64, 1315–22. 4. McDouall RM, Page CS, Hafizi S, Yacoub MH, Rose ML. (1996) Alloproliferation of purified CD4+ T cells to adult human heart endothelial cells, and study of second-signal requirements. Immunology 89, 220–6. 5. McDouall RM, Yacoub M, Rose ML. (1996) Isolation, culture, and characterisation of MHC class II-positive microvascular endothelial cells from the human heart. Microvasc Res 51, 137–52. 6. Muczynski KA, Ekle DM, Coder DM, Anderson SK. (2003) Normal human kidney HLA-DR-expressing renal microvascular endothelial cells: characterization, isolation, and regulation of MHC class II expression. J Am Soc Nephrol 14, 1336–48. 7. Berk BC, Abe JI, Min W, Surapisitchat J, Yan C. (2001) Endothelial atheroprotective and anti-inflammatory mechanisms. Ann N Y Acad Sci 947 , 93–109; discussion -11. 8. Traub O, Berk BC. (1998) Laminar shear stress: mechanisms by which endothelial cells transduce an atheroprotective force. Arterioscler Thromb Vasc Biol 18, 677–85. 9. Matharu NM, Rainger GE, Vohra R, Nash GB. (2006) Effects of disturbed flow on endothelial cell function: pathogenic implications of modified leukocyte recruitment. Biorheology 43, 31–44. 10. Nguyen KT, Clark CD, Chancellor TJ, Papavassiliou DV. (2008) Carotid geometry effects on blood flow and on risk for vascular disease. J Biomech 41, 11–9. 11. Perktold K, Thurner E, Kenner T. (1994) Flow and stress characteristics in rigid walled and compliant carotid artery bifurcation models. Med Biol Eng Comput 32, 19–26. 12. Resnick N, Gimbrone MA, Jr. (1995) Hemodynamic forces are complex regulators of endothelial gene expression. FASEB J 9, 874–82. 13. Resnick N, Yahav H, Schubert S, Wolfovitz E, Shay A. (2000) Signalling pathways in vascular endothelium activated by shear stress: relevance to atherosclerosis. Curr Opin Lipidol 11, 167–77.

14. Chien S. (2007) Mechanotransduction and endothelial cell homeostasis: the wisdom of the cell. Am J Physiol 292, H1209–24. 15. Hansson GK, Libby P. (2006) The immune response in atherosclerosis: a double-edged sword. Nat Rev 6, 508–19. 16. Hansson GK, Robertson AK, SoderbergNaucler C. (2006) Inflammation and atherosclerosis. Annu Rev Pathol 1, 297–329. 17. Jonasson L, Holm J, Skalli O, Bondjers G, Hansson GK. (1986) Regional accumulations of T cells, macrophages, and smooth muscle cells in the human atherosclerotic plaque. Arteriosclerosis (Dallas, Tex) 6, 131–8. 18. Zhou X, Stemme S, Hansson GK. (1996) Evidence for a local immune response in atherosclerosis. CD4+ T cells infiltrate lesions of apolipoprotein-E-deficient mice. Am J pathol 149, 359–66. 19. Song L, Leung C, Schindler C. (2001) Lymphocytes are important in early atherosclerosis. J Clin Invest 108, 251–9. 20. Rahmani M, Cruz RP, Granville DJ, McManus BM. (2006) Allograft vasculopathy versus atherosclerosis. Circ res 99, 801–15. 21. Lai JC, Tranfield EM, Walker DC, et al. (2003) Ultrastructural evidence of early endothelial damage in coronary arteries of rat cardiac allografts. J Heart Lung Transplant 22, 993–1004. 22. Rose ML, Gracie JA, Fraser A, Chisholm PM, Yacoub MH. (1984) Use of monoclonal antibodies to quantitate T lymphocyte subpopulations in human cardiac allografts. Transplantation 38, 230–4. 23. Lawson C.. (2005) Endothelium. In: Freshney RI, ed. Culture of Animal Cells; A manual of Basic Techniques. 5th ed. New Jersey: John Wiley and Sons, 404–8. 24. Jaffe EA, Nachman RL, Becker CG, Minick CR. (1973) Culture of human endothelial cells derived from umbilical veins. Identification by morphologic and immunologic criteria. J Clin Invest 52, 2745–56. 25. Frangos JA, Eskin SG, McIntire LV, Ives CL. (1985) Flow effects on prostacyclin production by cultured human endothelial cells. Science (New York, NY) 227, 1477–9. 26. Malek AM, Gibbons GH, Dzau VJ, Izumo S. (1993) Fluid shear stress differentially modulates expression of genes encoding basic fibroblast growth factor and

Leucocyte Adhesion Under Haemodynamic Flow Conditions platelet-derived growth factor B chain in vascular endothelium. J Clin Invest 92, 2013–21. 27. Lawson C, McCormack AM, Moyes D, et al. (2000) An epithelial cell line that can stimulate alloproliferation of resting CD4+ T cells, but not after IFN-gamma stimulation. J Immunol 165, 734–42.

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28. Strober W. (2006) Immunologic Studies in Humans. In: Current Protocols in Immunology. New Jersey: John Wiley and Sons. 29. Burns MP, DePaola N. (2005) Flowconditioned HUVECs support clustered leukocyte adhesion by coexpressing ICAM1 and E-selectin. Am J physiol 288, H194–204.

Chapter 4 Influence of Stromal Cells on Lymphocyte Adhesion and Migration on Endothelial Cells Helen M. McGettrick, Chris D. Buckley, G. Ed Rainger, and Gerard B. Nash Abstract Methods are described for analysing adhesion and migration of isolated lymphocytes on endothelial cell monolayers which have been co-cultured with different stromal cells, with or without additional cytokine treatment. The different cells types are grown on opposite sides of 3.0- or 0.4-μm pore filters depending on whether migration through the whole construct is to be analysed or adhesion to the endothelial cells alone. Assays may be “static” or filters can be incorporated into flow chambers so that cell behaviour can be directly observed under conditions simulating those in vivo. In general, by choice of method, one can evaluate efficiency of attachment and ability of cells to migrate across the endothelial monolayer, through the filter and through the stromal cell layer. Fluorescence microscopic examination of fixed filters can be used, e.g. to ascertain whether lymphocytes are retained by stromal cells. In general, static assays have the higher throughput and greatest ease of use, while the flow-based assays are more physiologically relevant and allow detailed recording of cell behaviour in real time. Key words: Lymphocyte, endothelial cells, fibroblasts, smooth muscle cells, stromal cells, adhesion, migration, cytokines, cell culture, co-culture.

1. Introduction Leucocyte recruitment is regulated by the local haemodynamic and stromal environments (1). Stromal cells such as fibroblasts or smooth muscle cells (SMC) may influence the normal physiological responses of endothelial cells (EC), while changes in their phenotypes may be associated with chronic inflammatory disorders. For instance, we found that culture of SMC in the secretory state with EC causes marked augmentation of the capture of all types F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_4, © Springer Science+Business Media, LLC 2010

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of flowing leucocytes in response to tumour necrosis factor alpha (TNF-α) (2). In addition, fibroblasts from inflamed tissue (the synovium of patients with rheumatoid arthritis) directly induced adhesion of neutrophils and lymphocytes when cultured with EC (3, 4). Such studies indicate that stromal cells can contribute to tissue- or vessel-specific patterns of leucocyte recruitment, may modulate inflammatory responses in general, or influence the development of disease at specific sites. Thus experimental models in which one can study how lymphocyte adhesion and migration are modified by stromal cells have a variety of potential uses. Here we describe several such models. In general, they rely on culturing EC on one side of a porous filter and the stromal cells on the other, with or without stimulation with cytokines. Subsequent assays of lymphocyte adhesion can be carried out after settling cells onto the endothelial surface for prolonged periods or during perfusion of cells in suspension. If larger pore filters (diameter ∼3 μm) are used, it is possible to follow lymphocyte migration through the two layers of cells. The “static” assays generally quantify how many cells migrate through the co-culture construct, but filters can be cut out and studied microscopically to assess whether cells are retained in the stromal layer. Practically, in flow systems, we have designed chambers for fluorescence microscopy which hold smaller filter inserts (24-well) and used them to quantify the capture process and whether cells become activated and stably adherent or not (5). We have also used larger 6-well inserts, cut the filters out and incorporated them in chambers designed for phase contrast microscopy (6), so that we can follow cells binding and then migrating through the endothelial monolayer, across the filter and into the stromal layer in real time.

2. Materials 2.1. Blood Cell Isolation

1. K2 EDTA in 10-ml tubes (Sarstedt, Numbrecht, Germany). 2. Histopaque 1077 (H1077) (Sigma–Aldrich, Poole, UK). 3. PBSA: Phosphate-buffered saline with 1 mM Ca2+ and 0.5 mM Mg2+ (PBS Gibco, Invitrogen Ltd., Paisley, UK), with 0.15% (w/v) bovine albumin (dilute from 7.5% culture-tested solution; Sigma) and 5 mM glucose. 4. M199-BSA: Medium 199 (M199 – Gibco) supplemented with 0.15% (w/v) bovine albumin (M199-BSA). 5. 2% glutaraldehyde (Cowley, Oxford, UK) diluted in 1/3 strength PBS to be isotonic. 6. Fluorescent nuclear stain, bisbenzimide (stock at 1 mg/ml; Sigma).

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1. M199 supplemented with gentamycin sulphate (35 μg/ml), human epidermal growth factor (10 ng/ml; Sigma E9644) and foetal calf serum (FCS) (20% v/v heat-inactivated) (all from Sigma). Adding hydrocortisone (1 μg/ml, from 10 mg/ml stock in ethanol; Sigma) improves growth if going beyond first passage. 2. Bovine skin gelatin (Type B, 2% solution, culture tested; Sigma). 3. Collagenase (type IA; Sigma) stored at –20◦ C at 10 mg/ml in PBS. Thawed and diluted to 1 mg/ml with M199 for use. 4. Autoclaved cannulae and plastic ties (electrical). 5. EDTA solution (0.02%, culture tested; Sigma). 6. Trypsin (2.5 mg/ml; Sigma) 7. 70% (v/v) ethanol or industrial methylated spirits. 8. Tumour necrosis factor alpha (TNF-α) (Sigma) and interferon-γ (IFN-γ; PeproTech Inc., London, UK) stored in small aliquots at –80◦ C.

2.3. Culture of Stromal Cells

1. Fibroblast complete medium: RPMI 1640 medium (Gibco) supplemented with 1x MEM-non-essential amino acids (stock was at 100x), 1 mM sodium pyruvate, 2 mM L -glutamine, 100 U/ml penicllin, 100 μg/ml streptomycin and FCS (10% v/v heat inactivated) (all from Sigma). 2. Promocell smooth muscle cell (SMC) medium supplemented with gentamycin sulphate (12.5 μg/ml), amphotericin B (12.5 ng/ml), human epidermal growth factor (10 ng/ml), basic fibroblast growth factor (2 ng/ml), dexamethasone (0.4 μg/ml) and FCS (5% v/v heat inactivated) (basal medium and all additional supplements from Promocell, Heidelberg, Germany). 3. Sterile dissecting scissors, scalpel and forceps. 4. EDTA solution (0.02%, culture tested; Sigma). 5. Trypsin (2.5 mg/ml; Sigma). 6. 70% (v/v) ethanol or industrial methylated spirits. 7. Dimethylsulphoxide hybrid-max (DMSO; Sigma).

2.4. Surfaces for Endothelial and Stromal Cell Culture for Assays

1. Cell culture inserts: High-density 0.4-μm or low-density 3.0-μm pore polycarbonate filter inserts in 24-, 12- or 6-well format (referred to as filters in future text) with matching culture plates (BD Pharmingen, Oxford, UK).

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2.5. Flow-Based Adhesion Assay

1. Parallel-plate flow chamber for fluorescence microscopy – for use with intact 24-well Transwell filters (5) (Fig. 4.1): A glass coverslip (5.5 × 2.6 mm). A non-compressible silicon gasket, 250 μm thick, containing a 41 × 6 mm slot which forms the flow channel. Specially designed chamber made up of two parallel plates held together with six screws (Wolfson Applied Technology Laboratory, University of Birmingham, Birmingham, UK). The lower plate has a machined receiving slot of a complementary size for the

Fig. 4.1. Fluorescence parallel-plate chamber. Two parallel perspex plates are separated by a glass coverslip (5.5 × 2.6 mm) and a non-compressible gasket cut from silicon sheet (Esco rubber, 250 μm thick; Bibby Sterilin Ltd., Stone, UK) with a flow channel of 41 × 6 mm and depth of 250 μm cut in it. The plates are held in place by hand-tightened metal screws. Filter inserts are placed into a machined receiving slot of a complementary size for the 24-well insert within the lower parallel plate. The insert forms a sealed base to the flow channel and is held in place by a smaller rubber gasket and perspex plate, held in place with metal screws. The surface of the filter is viewed in an upright microscope. The depth of the back plate holding the filter insert is too great to allow focussing of the transmitted light condenser, so that phase contrast images of high quality cannot be obtained. The surface is thus viewed during experiments using incident light illumination and fluorescence.

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24-well insert, along with inlet and outlet channels. The upper perspex plate has a machined slot to allow objective lens access and a shallow recess milled in it to receive the coverslip. 2. Parallel-plate flow chamber for phase contrast microscopy – for use with cut-out 6-well filters (6) (Fig. 4.2): A glass coverslip (75 × 26 mm; Raymond A. Lamb, Eastbourne, UK). A parafilm gasket (75 × 26 mm) containing a 20 × 4 mm slot. Specially designed chamber made up of two

Fig. 4.2. Phase contrast parallel-plate chamber. Cells are seeded onto 6-well filters, which are cut out onto the glass coverslip (76 × 26 mm). The filter and coverslip are covered with a parafilm gasket of the same size, with a flow channel of 20 × 4 mm and depth of 133 μm cut in it. These are placed on a perspex base-plate with a shallow matching recess milled into it and a viewing slot cut in it. The upper perspex plate has inlet and outlet holes positioned to match the flow channel formed by the gasket slot, allowing liquid to be perfused over the endothelium. The plates have matching holes (threaded in the lower plate) to allow them to be clamped together with hand screws. The parafilm gasket is cut afresh for each coverslip, using a thin aluminium sheet template, 76 × 26 mm, with 20 × 4 mm slot machined in it.

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perspex plates held together with six screws (Wolfson Applied Technology Laboratory, University of Birmingham, Birmingham, UK). The lower plate has a countersunk viewing slot cut in it and a shallow recess milled in it to receive the coverslip, filter and gasket. The upper perspex plate has inlet and outlet holes positioned to match the flow channel formed by the gasket slot, allowing liquid to be perfused over the HUVEC. The depth of the flow channel is defined by the thickness of the gasket, which averages 133 μm. The gasket is cut out from a sheet of parafilm using a rectangular aluminium template (75 × 26 mm) containing a 20 × 4 mm slot. 3. Flow system (Fig. 4.3): Syringe pump with smooth flow (e.g. PHD2000 infusion/withdrawal, Harvard Apparatus, South Natick, MA, USA). Electronic three-way microvalve with minimal dead volume (LFYA1226032H Lee Products Ltd., Gerrards Cross, Buckinghamshire, UK) and 12 V DC power supply. Silicon rubber tubing, internal diameter/external diameter (ID/OD) of 1/3 and 2/4 mm (Fisher Scientific, Loughborough, UK). Three-way stopcocks (BOC Ohmeda AB, Helsingborg, Sweden). Sterile, disposable syringes (2, 5, 10 ml Becton Dickinson, Oxford, UK) and glass 50 ml syringe for pump (Popper MicroMate; Popper and Sons Inc., New York, USA). 4. Video microscope: Microscope with heated stage or preferably with stage and attached flow apparatus enclosed in a temperature-controlled chamber at 37◦ C, with phase con-

Fig. 4.3. Schematic representation of assembled flow system. The parallel-plate flow chamber was incorporated into a perfusion system mounted on the stage of a phase contrast and fluorescence microscope enclosed in a perspex chamber at 37◦ C. It was connected by flexible silicon tubing to a Harvard withdrawal syringe pump at one end or an electronic switching valve at the other. A suspension of purified leucocytes or cell-free wash buffer was perfused through the chamber, typically at a constant wall shear stress of 0.1 Pa. Images from the microscope were captured using CCD video camera and video tape and subsequently digitised for analysis or captured using digital camera straight to computer.

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trast and fluorescence (UV) optics. Video camera (e.g. analogue Cohu 4912 monochrome camera with remote gain control), monitor and video recorder (e.g. time lapse, Panasonic AG-6730) or digital camera (e.g. Olympus UCMAD3 QICAH) directly connected to computer (see below). 5. Image analysis: Computer with video capture card (if using video recordings) or input for digital cameras and specialist software for counting cells, measuring motion, etc. There are a range of commercial packages available, as well as image analysis software (NIH Image http: //rsb.info.nih.gov/nih-image/) available free over the Internet. We currently use Image-Pro software (Media Cybernetics).

3. Methods 3.1. Leucocyte Isolation

1. Draw blood from the ante-cubital vein of normal human volunteers with a minimum of stasis, dispense into K2 EDTA tubes and mix gently but thoroughly (see Note 1). 2. Place 5 ml H1077 in a 10-ml centrifuge tube. 3. Layer whole blood (5 ml) from K2 EDTA tube on top. 4. Centrifuge at 800g for 30 min. 5. Retrieve the mononuclear cells from the top of the gradient at the interface of plasma and H1077. 6. Wash cells twice in PBSA or M199-BSA. 7. To deplete mononuclear cells of monocytes, place in culture dish for 30 min at 37◦ C for monocytes to sediment and adhere. Gently wash off enriched peripheral blood lymphocytes (PBL). 8. Count lymphocytes and dilute to desired concentration in PBSA or M199-BSA or endothelial culture medium (see Note 2). 9. For fluorescence, pre-label cells with 1 μg/ml bisbenzimide 15 min in the dark (see Note 3).

3.2. Isolation and Culture of Endothelial Cells and Stromal Cells

There are various methods for culture of endothelial and stromal cells from different sources, and for the novice, it is probably best to start by buying cells and media from commercial suppliers. A variety of different endothelial cells, fibroblasts and smooth muscle cells are available (e.g. from Asterand, Clonetics,

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ECCAC, Promocell). Our current method for isolating and culturing human umbilical vein endothelial cells is given below, adapted from Cooke et al. (7). 3.2.1. Isolation and Primary Culture of HUVEC

1. Place the cord on paper towelling in a tray and spray liberally with the 70% ethanol. Choose sections of about 3–4 in. that do not have any clamp damage. Each 3–4 in. piece of cord equates to 1 flask of primary cells. 2. Locate the two arteries and one vein at one end of the cord. 3. Cannulate the vein and secure the cannula with an electrical tie. 4. Carefully wash through the vein with PBS using a syringe and blow air through to remove the PBS. 5. Cannulate the opposite end of the vein and secure with electrical tie. 6. Inject collagenase (∼10 ml per 3–4 in.) into vein until both cannulae bulbs have the mixture in them. 7. Place the cord into an incubator for 15 min at 37◦ C. 8. Remove from the incubator and tighten the ties. Massage the cord for ∼1 min. 9. Flush the cord using a syringe and 10 ml PBS into a 50-ml centrifuge tube. 10. Push air through to remove any PBS, repeat this twice more (3 × 10 ml). 11. Centrifuge at 400g for 5 min. Discard supernatant. 12. Resuspend the cells in ∼1 ml of culture medium and mix well with pipette 13. Make up to 4 ml in complete medium. 14. Add cell suspension to a 25 cm2 culture flask. 15. Change medium after 2 h, the next day and every subsequent 2 days. Cells should be confluent in about 3–7 days.

3.2.2. Isolation and Culture of Primary Fibroblasts

Here we give the procedures for isolating dermal fibroblasts and below, for isolating arterial smooth muscle cells from umbilical arteries. The former would require a clinical link through which to obtain skin tissue, e.g. from patients undergoing surgery. 1. Obtain tissue (e.g. ∼1 cm3 ) in a sterile container on ice. 2. If tissue is bloody, wash first with RPMI alone, centrifuge at 300g for 5 min and discard supernatant. 3. Place tissue into a sterile petri dish. Each 1 cm3 piece of tissue will seed four flasks. 4. Using sterile scalpel, remove the fatty (yellow) tissue from the skin.

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5. “Tease” apart the grey skin tissue into fine strand-like remnants less than 1 mm3 . It may be necessary to pipette on a small amount of medium if the tissue starts to dry out and become “sticky”. 6. Add ∼0.25 cm3 of tissue into a 25 cm2 culture flask (T25 flask). 7. Add 7 ml of fibroblast complete medium. 8. Incubate undisturbed at 37◦ C in 5% CO2 for 3 weeks. (Allow time for the fibroblasts to grow out of the tissue.) 9. Change medium by aspirating out two-thirds of old medium and replacing it with fresh medium. During this time only change medium when it becomes yellowish (see Note 4). 10. Initial outgrowth of adherent cells is usually seen after 1–2 weeks. Confluence is normally reached after 3–6 weeks although this depends on tissue type and may vary between donors. 3.2.3. Isolation and Culture of Primary “Secretory” Smooth Muscle Cells from Umbilical Artery

1. Place a 2 in. section of umbilical cord in a sterile petri dish. 2. Locate the two arteries and one vein at one end of the cord. 3. Hold the cord with the sterile forceps. 4. Using sterile dissecting scissors cut along the vein (see Note 5). 5. Open the cord flat and locate the arteries. 6. Cut between the arteries so that they are separated from one another. 7. Cut away all the extraneous tissue surrounding one artery. It is essential to remove all the surrounding tissue to prevent contamination. 8. Cut the artery into 0.5–1 mm pieces. 9. Add 6–10 pieces to a T25 flask. (One artery can be split between three separate T25.) 10. Add 5 ml of Promocell SMC medium. 11. Incubate undisturbed at 37◦ C for 3 weeks. (Allow time for smooth muscle cells to migrate out of the artery.) 12. Visualise under phase contrast microscopy: Smooth muscle cell colonies should have formed. Culture can be continued until confluence is reached (see Note 6).

3.2.4. Dispersal of Endothelial and Stromal Monolayers for Passaging

1. Rinse a flask containing a confluent primary monolayer of cells or smooth muscle cell colonies with 2 ml EDTA solution.

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2. Add 2 ml of trypsin solution and 1 ml of EDTA for 1–2 min at room temperature, until the cells became detached. Tap on bench to loosen. 3. Add 8 ml of culture medium to the flask and transfer the resulting suspension into a 15-ml tube. 4. Centrifuge at 400g for 5 min. 5. Remove supernatant and resuspend the cell pellet in 0.5 ml of culture medium and disperse by sucking them in and out of a pipette tip. 6. Make up to 3 volumes of culture medium and seed three flasks (see Note 7). 7. Repeat steps 1–5 to expand smooth muscle cells and fibroblasts for a minimum of four cycles before use in assays. 3.2.5. Freezing Stromal Cells

1. Repeat steps 1 through 4 from Section 3.2.4. 2. Add 3 ml of ice-cold DMSO:FCS (1:9 ratio) per 75 cm2 culture flask. 3. Add 1 ml into an ice-cold cryovial (Nalgene). 4. Store in –20◦ C for 2 h. 5. Transfer to –80◦ C overnight. 6. Transfer to liquid N2 until future need. 7. To use, thaw the cryovial rapidly at 37◦ C and transfer the 1 ml contents into 5 ml of cold medium (choose appropriate medium for different stromal cells). 8. Centrifuge at 400g for 5 min. 9. Remove supernatant and resuspend the cell pellet in 4 ml of culture medium and transfer to a T25 flask.

3.3. Establishing Endothelial–Stromal Cell Co-cultures on Filters

Depending on the type of assay, endothelial cells will be seeded inside the filter (inner surface) and stromal cells on the outside (outer) surface or vice versa (see Note 8). For static assays and for the parallel-plate flow chamber which takes cut out 6-well filters, seeding of the endothelial cells is on the inner surface, while for the flow chamber which takes intact 24-well inserts, seeding of the endothelial cells is on the outer surface. Whichever assay is employed, the stromal cells are seeded first.

3.3.1. Establishing Stromal Cell Co-cultures

1. Trypsinise a single flask of T75 of stromal cells as in Section 3.2.4 and suspend cells in 8 ml (see Note 9). 2a. For use in static assays or phase contrast chamber, invert the filter in a sterile box and carefully seed stromal cells onto the outer surface of the inverted filter (200 μl on a 24-well filter; 500 μl on a 6-well filter). Incubate at 37◦ C for 1 h, after which the filter is re-inverted and placed into wells containing culture medium.

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2b. For use in the fluorescence chamber, seed 200 μl of stromal cells on the inner surface of the 24-well filters in their wells. 3. After 24 h, trypsinise a single flask of HUVEC as in Section 3.2.4. This will seed four 6-well filters or twenty 24-well filters (see Note 10). 4. Resuspend in 8 ml of medium (see Note 11). 5a. For use in static assays or phase contrast chamber, aspirate the medium from the upper chamber and seed 2 ml of HUVEC to each filter (inner surface) for 6-well format or 200 μl in 24-well format. 5b. For use in the fluorescence chamber, seed HUVEC onto the outer surface as in 2a. 6. Culture endothelial cells with stromal cells for 24 h. 7. Treat with cytokines if desired (see Note 12). 3.4. Adhesion and Migration of Lymphocytes Through Co-cultures on 3.0-µm Pore Filters Under Static Conditions

Below we describe the volumes required when using 24-well filters; to use 12-well or 6-well filters, the medium and cell numbers added must be scaled up accordingly (Fig. 4.4). 1. Remove cytokine-containing medium from the upper and lower chamber. 2. Add 700 μl of fresh M199+BSA to the lower chamber and 200 μl of PBL or chosen lymphocyte sub-type (2 × 106 cells/ml in M199+BSA) to the upper chamber (see Note 13).

Fig. 4.4. Schematic representation of the Transwell assay. Lymphocytes (2 × 106 cells/ml) are added into the upper chamber and allowed to interact with the TNF-stimulated endothelial cells (HUVEC). The lymphocytes either remain nonadherent or become attached to the surface of the HUVEC or migrate through them (2 = white). The lymphocytes may migrate through the filter and either remain adherent to the basal surface among the fibroblasts (3) or fully migrate into the lower chamber of the tissue culture plate (4). Counting of cells retrieved from the upper and lower chambers determines the percentage of lymphocytes that are non-adherent (1) or that fully transmigrate (4). Counting of stained cells below the filter using fluorescence microscopy allows analysis of those that transmigrated but were retained by fibroblasts (3). Total transmigration = (3) + (4).

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3. Allow the PBL to settle, adhere and migrate (see Fig. 4.4) at 37◦ C for desired time (typically 24 h) (see Note 14). 4. Stop the experiment by transferring the filter into a fresh well. 5. Transfer the PBL from the upper chamber (above filter) into a fresh well. 6. Wash the upper chamber twice with 200 μl of M199+BSA and add washouts to the upper chamber samples. These represent the non-adherent PBL. 7. Retrieve cells from the original lower chamber, rinse out with 300 μl of M199+BSA and pool with retrieved cells. Examine well microscopically to ensure that all cells are removed and wash further if necessary (see Note 15). The pooled samples represent those cells that had migrated through both endothelial and stromal layers. 8. Count the “non-adherent” and “transmigrated” samples using a Coulter counter (see Note 16) or haemocytometer. 9. Fix the filter in 2% isotonic glutaraldehyde containing at 1 μg/ml bisbenzimide for 15 min in the dark and then wash four times in PBS. 10. Cut the filter out using a scalpel, directly onto a microscope slide and mount with anti-fade agent (e.g. DABCO; Sigma). 11. Using a fluorescence microscope with UV illumination and 40x objective, focus on the nuclei of the HUVEC. Move the focus down through the filter (a distance of ∼10 μm) until transmigrated lymphocytes adherent to the back of the filter come into view and count these cells. 12. These cells represent those which crossed the endothelium and filter, but were retained by the stromal cells. Their number can be added to the counts from the lower chamber to give the number of lymphocytes that migrated through endothelial cells and filter. 13. All counts should be expressed as a percentage of those originally added. 14. From this data, the percentage of adherent cells, the percentage transmigrated (below the filter and in the lower chamber) and the percentage held by the stromal cells can be determined. 3.5. Flow-Based Assay of Lymphocyte Adhesion and Migration

We have described two different flow chambers. Using 0.4-μm pore filters, the fluorescence chamber allows the analysis of lymphocyte recruitment from flow (capture, rolling and firm adhesion). The phase contrast chamber is suitable for visualising lymphocyte recruitment and additionally allows analysis of

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migration on, through and under the endothelial monolayer. If 3-μm pore filters are used, then migration through the filter into the stromal layer can also be recorded. In our experience, this is a slow process for lymphocytes, but much faster for neutrophils. 3.5.1. Setting Up the Flow Assay

1. Assemble flow system without flow chamber attached (Fig. 4.3). The electronic valve has a common output, and two inputs, from “Wash reservoir” and “Sample reservoir”, which can be selected by turning electronic valve on and off. 2. Fill wash reservoir with PBSA and rinse through all tubing, valves and connectors with PBSA, ensuring bubbles are displaced (e.g. using syringe attached to three-way tap for positive ejection). Fill sample reservoir with PBSA and rinse through valve and attached tubing. Prime downstream syringe and tubing with PBSA and load into syringe pump. All tubing must be liquid-filled to ensure prompt starting and stopping of flow.

3.5.2. Assembling and Connecting the Flow Chamber for Fluorescence Microscopy

1. Align the glass coverslip and large silicon gasket on the top parallel plate, lower the bottom parallel plate onto the gasket and secure with metal screws (Fig. 4.1). 2. Insert the complete 24-well filter into the machined receiving slot in the bottom parallel plate. The endothelial side of the filter aligns with the bottom plate forming a sealed base to the flow channel, butting onto the silicon gasket. 3. Place the small rubber gasket and perspex plate over the base of the filter and secure into place with metal screws. 4. Connect Portex Blue Line Manometer connecting tubing (Portex Ltd, UK) into the inlet and outlet holes in the sides of the bottom plate.

3.5.3. Assembling and Connecting the Phase Contrast Flow Chamber

1. Gently place the 6-well filter onto the centre of the 75 × 26 mm glass coverslip, stromal cells on the outer surface of the filter in direct contact with the coverslip (Fig. 4.2). 2. Using a new scalpel blade (type 10A), carefully cut out the filter. 3. Smooth a section of parafilm on a glass microscope slide and cut round to form a gasket (75 × 26 mm). Cut a slot 20 × 4 mm to form the flow channel using an aluminium template (Fig. 4.2) 4. Place the parafilm gasket over the coverslip, with the flow channel over the filter. 5. Put the glass coverslip into the milled recess in the bottom perspex plate of the flow chamber and place the flow

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channel (top perspex plate) over the endothelial cell surface (Fig. 4.2). 6. Screw the top and bottom perspex plates together. 7. Connect Portex Blue Line Manometer connecting tubing (Portex Ltd., UK) into the inlet and outlet holes in the top perspex plate. 3.5.4. Perfusing Cellular Suspension and Recording Behaviour

1. Place the flow chamber onto microscope stage and start flow by turning on syringe pump in withdrawal mode, with electronic valve and three-way tap in position to allow delivery of PBSA from wash reservoir. 2. Wash out culture medium and locate the endothelial surface using phase contrast (or bright field microscopy if subsequent observations are with fluorescent lymphocytes). 3. Adjust flow rate to that required for assay. To obtain a given wall shear rate or stress, the flow rate (Q) will depend on the flow channel dimensions (see Note 17). 4. Perfusion of cells is typically at a flow rate Q = 0.525 ml/min for the fluorescence flow chamber (where the channel depth and width are 250 μm and 6 mm, respectively) or Q = 0.099 ml/min for the phase contrast flow chamber (where the channel depth and width are 133 μm and 4 mm, respectively). These are equivalent to a wall shear rate of 140s–1 and wall shear stress of 0.1 Pa (= 1 dyne/cm2 ), similar to those found in post-capillary venules. 5. Load isolated cells into sample reservoir and allow to warm for 5 min. 6. Switch the electronic valve so that cell suspension is drawn through microslide. 7. Deliver timed bolus (e.g. 4 min). Typically, flowing cells will be visible after about 30s, the time required to displace dead volume in valve and tubing. 8. Switch electronic valve so that PBSA from wash reservoir is perfused. Again, 30–60 s will be required before all cells have been washed through the flow chamber or microslide. 9. Video recordings can be made as desired during inflow and washout of cells. Typically, a series of fields may be recorded along the centreline of the chamber during inflow (e.g. six fields recorded for 20s each during the last minute of the bolus), for off-line analysis of the behaviour (e.g. rolling or stationary adhesion) of the cells. Another series can be made after 1 min washout (when the bolus is complete) for analysis of the numbers of adherent cells and their behaviour. Fields may be recorded at later times (e.g. after

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a further 5 and 10 min) to assess progress of migration (e.g. through the monolayer or filter), and/or a field can be recorded continuously for 5–10 min to track individual cells and assess velocity of migration. At later time points, recordings can be made of cells beneath 3-μm pore filters (by focussing 10 μm down) and beneath the stromal cell layer. 10. If a defined timing protocol is developed, digital images or sequences of digital images could be recorded instead of video images. The continual recording of the latter gives flexibility in analysis. 11. Data analysis is carried out off-line. 3.5.5. Analysis of Cell Behaviour from Video Recordings

1. Make recordings of a microscope stage-micrometer oriented parallel and perpendicular to the flow. Use this to calibrate the size of video field observed on the monitor during playback and the image analysis software. 2. To quantify the numbers of adherent cells and their behaviour, digitise a sequence of images 20 at 1s intervals from recordings made at the desired times. 3. When played in a loop, cells can be distinguished which are rolling (circular phase-bright cells tumbling slowly at ∼1–10 μm/s over the surface) or stably adherent on the endothelial surface (phase-bright cells typically with distorted outline and migrating slowly on the surface) or transmigrated cells (phase-dark spread cells migrating under the HUVEC). Non-adherent cells will only be visible as blurred streaks. Migrated cells beneath the filter and beneath the stromal cell layer appear phase-bright cells with a distorted shape. When using the fluorescence system, all recruited cells appear bright, with rolling cells being spherical and stably adherent/migrating cells typically being distorted in shape. 4. Count the cells present on a stop-frame video field at the start of a sequence, and then play the loop to assign them as rolling, stationary or transmigrated. Repeat and average counts for the series of sequences recorded at a given time. 5. Convert counts of total adherent cells (rolling + stationary + migrated) to number/mm2 from the known field dimensions. Divide this by the number of cells perfused (in units of 106 cells) to obtain number adherent/mm2 /106 perfused. The number perfused is calculated by multiplying the concentration of the suspension (usually 106 per ml) by the flow rate by the duration of the bolus (e.g. 4 min). This normalisation allows correction for changes in conditions (bolus duration, cell concentration, flow rate)

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between experiments and effectively calculates an efficiency of adhesion. 6. Express the numbers of cells rolling, stationary adherent or transmigrated as percentages of the total adherent cells. When location below the filter is analysed, counts should be added to the total and the percentages in this location calculated. 7. Analysis at different times (e.g. after 1, 5 or 10 min of washout) can be used to quantify the progress of migration through the different layers (endothelium, filter and stromal cells) or any changes in behaviour. 8. To measure rolling velocity, mark the leading edges of a series of cells to be followed and move to second captured frame. Remark the leading edges and record the distance moved. Repeat through the 10s sequence. This will yield data for position versus time. Velocity for each cell can be averaged over the observation time, and estimates of variation in velocity made if desired. 9. To measure migration velocity in extended video sequences, images are digitised at 1 min intervals over 5– 10 min. The cells are outlined and the positions of their centroids recorded at each minute. The changes in positions are used to calculate the distances migrated in each minute. The average velocities can be calculated from the sequence.

4. Notes 1. There are various methods for isolating lymphocytes from blood and Section 3.1 describes a simple one that we use regularly. In the early stages, it is advisable also to test viability of preparations (e.g. ∼99% viable judged with trypan blue) and purity. Lymphocytes prepared in this way will still have some monocyte contamination. Further purification of lymphocyte subsets can be made using immunomagnetic selection (e.g. Dynabeads, Dynal Biotech UK, Bromborough, UK; MACS, Miltenyi Biotec Ltd., Bisley, UK). 2. When added to HUVEC, PBSA is sufficient to maintain viability in short assays for up to about an hour. However, PBSA is unable to maintain an intact HUVEC monolayer following 24 h of culture as judged by visual observations and a decrease in electrical resistance across the monolayer. Lymphocytes can be suspended in endothelial culture

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medium, but in our experience M199+BSA provides a simpler medium without growth factors and FCS which maintains endothelial morphology and electrical resistance for 24 h. 3. We have analysed the effects of various dyes on the capture, adhesion and migration of neutrophils and lymphocytes (8, 9). In our hands, bisbenzimide has the least effect on behaviour. In general, the activatory or damaging effects of fluorescent dyes depend on the duration for which the cells are illuminated. Ideally, illumination should be restricted to the time necessary to capture microscopic images and kept as short as possible. 4. It is important to leave the fibroblasts undisturbed for as long as possible while the cells are growing out of the tissue section. 5. We usually isolate the HUVEC first, which opens and empties the vein, making it easier to cut along. 6. Fast-growing smooth muscle cells in the secretory phenotype are isolated and expanded in this medium. Slowgrowing contractile smooth muscles cells can be generated in vitro by culturing the “secretory” smooth muscles for 72 h in Promocell SMC medium containing only the 5% FCS (omit the growth factors from the complete medium). However, that medium is not compatible with endothelial co-culture (5). 7. From primary smooth muscle cell cultures, we typically expand to passage 4 before freezing in aliquots equivalent to one T25 flask. With primary fibroblasts, of four T25 from a divided tissue sample, one would be split three ways and passaged further and three frozen in liquid nitrogen for later expansion. Experiments would typically be done with cells between passages 4 and 10. 8. Some studies on transendothelial migration have precoated the Transwell filters with collagen or fibronectin (FN) (10–13). It has been suggested that this coating increases the percentage of leucocyte migration. However, comparing uncoated filters with FN-coated filters (either coated before the assay with 20 μg/ml human plasma FN (Sigma) or bought pre-coated with 170–200 μg/ml FN from BD), we found no significant differences in the percentages of neutrophils transmigrating. We have not studied the effects of pre-coating on lymphocyte adhesion and migration. 9. Typically, we seed 2.5 × 104 fibroblasts or 1 × 105 smooth muscle cells in 200 μl on 24-well filters and 30 times as many fibroblasts (7.5 × 105 ) in 500 μl on 6-well filters.

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10. Cells from one confluent 25 cm2 flask of HUVEC, resuspended in 8 ml, will seed the inside of four 6-well filters (2 ml per filter). Alternatively, one 25 cm2 flask can be resuspended in 4 ml and used to seed the inner or outer surfaces of twenty 24-well filters (200 μl per filter). Both produce a confluent monolayer within 24 h. 11. Our standard medium for growing HUVEC contains hydrocortisone. However, this can alter the inflammatory response induced by fibroblasts (3). It is important to consider whether the growth factors and corticosteroids added to medium alter the behaviour of the different cells. In this case, we withdraw hydrocortisone from the culture medium used for co-culture. In some experiments we attempted to co-culture HUVEC with SMC in the contractile phenotype. This required culture in the absence of growth factors and with only 5% FCS in the Promocell SMC medium. However, this medium was unsuitable for endothelial cell culture (2). 12. In studies of lymphocyte adhesion and migration, we have stimulated HUVEC with TNF (100 U/ml), IFN (10 ng/ml) or both for 24 h prior to assay. For endothelial– fibroblast co-cultures, we have tested how different fibroblasts modulate response to the combined cytokines. In studies with endothelial–smooth muscle cell co-cultures, we use TNF over a range of concentrations for 24 h (2). 13. We have used PBL but assessed the content of different sub-populations in the added cells and those that transmigrated using flow cytometry. In this way, for example, migration of CD4+ and CD8+, naive or memory cells can be compared by choice of appropriate fluorescently labelled antibodies. 14. Lymphocytes are slower at migrating through filters than neutrophils (14–16). While optimising this protocol, we analysed lymphocyte migration at 2, 4 and 24 h for unstimulated HUVEC and after stimulation with various cytokine combinations. Transmigration was very low at the early time points and increased significantly with time. We routinely use a 24-h period for lymphocytes and 2-h period for neutrophils. 15. If lymphocytes adhere to the bottom of the well, it can be pre-coated with the non-adhesive substrate polyHEMA (17, 18). 16. There are alternative methods of analysing lymphocyte counts including pre-labelling with fluorescent dyes or radioisotopes. BD Biosciences supplies a Transwell filter which has a patented light-tight PET membrane that

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efficiently blocks the transmission of light within the range of 490–700 nm (www.bdbiosceinces.com). Using this system, the number of transmigrated fluorescent cells beneath the filter can be analysed using a fluorescence plate reader during the assay. 17. The flow rate (Q) required to give a desired wall shear rate (γ w in s–1 ) or wall shear stress (τw in pascal, Pa) is calculated from the internal width (w) and internal depth (h) of the flow channel and the viscosity (n) of the flowing medium using the formulae γw = (6.Q ) / (w.h 2 ) τ = n.γ For the fluorescence flow chamber, w and d are 6 mm and 250 μm, respectively. For the phase contrast flow chamber, w and d are 4 mm and 133 μm, respectively, although the depth varies slightly from parafilm gasket to gasket.

References 1. Nash GB, Buckley CD, Rainger GE. (2004) The local physicochemical environment conditions the proinflammatory response of endothelial cells and thus modulates leukocyte recruitment. FEBS Lett. 569, 13–17. 2. Rainger GE, Nash GB. (2001) Cellular pathology of atherosclerosis: smooth muscle cells prime cocultured endothelial cells for enhanced leukocyte adhesion. Circ Res 88, 615–22. 3. Lally F, Smith E, Filer A, Stone MA, Shaw JS, Buckley C, Nash GB, Rainger GE. (2005) A novel mechanism of neutrophil recruitment in a coculture model of the rheumatoid synovium. Arthritis Rheum 52, 3460–649. 4. McGettrick HM, Filer A, Buckley CD, Rainger GE, Nash GB. (2007) Modulation of endothelial responses by the stromal microenvironment: effects on leukocyte recruitment. Biochem Soc Trans 35, 1161–2. 5. Rainger GE, Stone P, Morland CM, Nash GB. (2001) A novel system for investigating the ability of smooth muscle cells and fibroblasts to regulate adhesion of flowing leukocytes to endothelial cells. J Immunol Met 255, 73–82. 6. Chakravorty S, McGettrick HM, Butler LM, Rainger GE, Nash GB. (2006) Kinetics of neutrophil migration into and away from

7.

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the subendothelial compartment in vitro: effects of flow and of CD31. Biorheology 43, 71–82. Cooke BM, Usami S, Perry I, Nash GB. (1993) A simplified method for culture of endothelial cells and analysis of adhesion of blood cells under conditions of flow. Microvas. Res 45, 33–45. Abbitt KB, Rainger GE, Nash GB. (2000) Effects of fluorescent dyes on selectin and integrin-mediated stages of adhesion and migration of flowing leukocytes. J Immunol Met 239, 109–19. Smith E, Lally F, Stone MA, Shaw JS, Nash GB, Buckley CD, Ed RG. (2006) Phototoxicity and fluorotoxicity combine to alter the behavior of neutrophils in fluorescence microscopy based flow adhesion assays. Microsc Res Tech 69, 875–84. Lampugnani MG, Resnati M, Raiteri M, Pigott R, Pisacane A, Houen G, Ruco LP, Dejana E. (1992) A novel endothelial-specific membrane protein is a marker of cell-cell contacts. J Cell Biol 118, 1511–22. Kuijpers TW, Hakkert BC, Hart MHL, Roos D. (1992) Neutrophil migration across monolayers of cytokine-prestimulated endothelial cells: a role for plateletactivating factor and IL-8. J Cell Biol 117, 565–72.

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12. Cooper D, Lindberg FP, Gamble JR, Brown EJ, Vadas MA. (1995) Transendothelial migration of neutrophils involves integrinassociated protein (CD47). Proc Natl Acad Sci USA 92, 3978–82. 13. Everitt EA, Malik AB, Hendey B. (1996) Fibronectin enhances the migration rate of human neutrophils in vitro. J Leukoc Biol 60, 199–206. 14. Oppenheimer-Marks N, Lipsky PE. (1997) Migration of naive and memory T cells. Immunol Today 18, 456–7. 15. Oppenheimer-Marks N, Ziff M. (1988) Migration of lymphocytes through endothelial cell monolayers: augmentation by interferon-gamma. Cellular Immunol 114, 307–23.

16. Borthwick NJ, Akbar AN, MacCormac LP, Lowdell M, Craigen JL, Hassan I, Grundy JE, Salmon M, Yong KL. (1997) Selective migration of highly differentiated primed T cells, defined by low expression of CD45RB, across human umbilical vein endothelial cells: effects of viral infection on transmigration. Immunol 90, 272–80. 17. Kettritz R, Xu YX, Kerren T, Quass P, Klein JB, Luft FC, Haller H. (1999) Extracellular matrix regulates apoptosis in human neutrophils. Kidney Int 55, 562–71. 18. Folkman J, Moscona A. (1978) Role of cell shape in growth control. Nature 273, 345–9.

Chapter 5 Discriminating Between the Paracellular and Transcellular Routes of Diapedesis Jaime Millán, Eva Cernuda-Morollón, and Severine Gharbi Abstract Leucocyte transendothelial migration (TEM) or diapedesis is pivotal in leucocyte trafficking during the inflammatory and immune responses. The endothelium plays an active role in this process, triggering an array of signalling pathways and reorganizing its cytoskeleton and membrane to facilitate leucocyte TEM. Diapedesis can occur between endothelial cells (paracellular) or through individual endothelial cells (transcellular). This latter route accounts for up to 30% of the total diapedesis in certain endothelial cell types in vitro. Mechanisms underlying both routes of diapedesis have been subjected to intense investigation during recent years. Here we describe a method to discriminate between the paracellular and the transcellular routes of diapedesis in vitro. The method is based on a transmigration assay of human T lymphoblasts through TNF-α-stimulated human primary endothelial monolayers, a triple fluorescence labelling of F-actin, the adhesion receptor ICAM-1 and the junctional protein β-catenin and a subsequent acquisition of z-stacks of high-resolution confocal sections. Key words: Endothelial cells, lymphocyte, transcellular, paracellular, transmigration, diapedesis, junctions, ICAM-1.

1. Introduction Leucocytes continuously traffic between the bloodstream and the surrounding tissue during immune and inflammatory responses. In response to proinflammatory stimuli, endothelial cells express a set of adhesion receptors, such as selectins, ICAM-1 or VCAM-1, that recruit and facilitate the local passage of leucocytes towards the inflammatory focus (1). The process of leucocyte extravasation through the endothelium has been termed leucocyte transendothelial migration (TEM) or diapedesis. F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_5, © Springer Science+Business Media, LLC 2010

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Diapedesis is the key step during leucocyte egression from the circulatory system, which in turn appears altered during the early stages of diseases as important as atherosclerosis or multiple sclerosis (2, 3). During diapedesis numerous endothelial receptors play coordinated responses upon leucocyte engagement. One of these responses is a remarkable three-dimensional remodelling of the luminal endothelial membrane, essential for leucocyte transmigration (4). Therefore, adopting an in vitro system easy to manipulate and to scrutinize by high-resolution microscopy is indispensable in order to investigate mechanisms regulating TEM. It is commonly accepted that diapedesis or TEM occurs through a paracellular route, in which the leucocyte interacts with a set of surface receptors located at intercellular junctions, such as PECAM-1 or CD99, disrupts cell–cell junctions and crosses the endothelial monolayer between two adjacent cells (5–7). However, several in vivo electron microscopy studies have suggested for many years the possibility that leucocytes can transmigrate leaving cell–cell junctions unaltered and following a transcellular route through individual cells (8). Based on these observations, some in vitro systems have been adopted in order to study mechanisms regulating these two routes of diapedesis by confocal microscopy. Using this strategy, it has been shown that trafficking of at least ICAM-1 to non-coated or caveolar vesicles may be involved in this route of transmigration ((9–11), recently reviewed in (12)). The proportion of transcellular versus paracellular routes in vitro varies depending on the leucocyte and endothelial cell types analysed, but transcellular diapedesis can account for up to 30% of total transmigration of T cells across microvascular endothelial monolayers (11). Here we report a detailed protocol to distinguish these two routes of diapedesis in vitro based on the localization by confocal microscopy of filamentous actin (F-actin), ICAM-1 and the junctional βcatenin in co-cultures of T lymphoblasts with primary endothelial cells. We provide specific protocols to perform transmigration assays of T lymphoblast with human umbilical vein endothelial cells (HUVECs) or with human dermal microvascular endothelial cells (HDMVECs), two of the most common human primary endothelial cell types used in vitro.

2. Materials 2.1. Cell Lines and Cell Culture Reagents

1. HUVECs from Lonza (Slough, UK) are cultured in growth medium provided by the manufacturer: EBM-2 (CC-3156), supplemented with 2% heat-inactivated foetal bovine serum

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(FBS), 2 mM glutamine and endothelial cell growth supplement (EGM-2-singlequots; CC-4176) (complete medium). EBM-2 starving medium is made by adding 1% FBS plus glutamine and antibiotics, without growth supplement. 2. Human dermal microvascular endothelial cells (HDMVEC or HDMEC, cat. C-12210) and their medium, Endothelial Cell Growth Medium MV (cat. C-22020) (complete medium), are provided by Promocell (Heidelberg, Germany). 3. Both HUVECs and HDMEC are grown on Nunclon flasks coated with fibronectin from human plasma at 10 μg/ml (Sigma-Aldrich, Gillingham, UK). A stock of 0.5 mg/ml of fibronectin is prepared in PBS, aliquoted and stored at –20◦ C. 4. T lymphocytes are obtained from single-donor buffy coats and cultured in RPMI 1640 (Gibco) supplemented with 10% human AB male serum (BioWest, Nuaille, France). Phytohemagglutinin (PHA) for T-cell stimulation is from Sigma-Aldrich. Interleukin-2 (IL-2) is from Roche Diagnostics (Mannheim, Germany). 5. Phosphate-buffered saline (PBS) and trypsin (0.05%)/ EDTA (0.02%) solutions (Gibco). 6. TNF-α is provided by Insight Biotech. Stock solution is prepared in sterile water. Aliquots can be stored at –80◦ C for 12 months. 2.2. Immunofluorescence

1. Microscope coverslips (13 mm diameter, 1.5 mm thickness) are from VWR International (Lutterworth, UK). Microscopy slides are from Thermo Scientific (Braunschweig, Germany). 2. Paraformaldehyde (PFA) (Sigma-Aldrich) is prepared at 4% (W/V) in PBS, aliquoted and stored at –20◦ C. Thawed aliquots can be stored at 4◦ C protected from light for up to 1 week. 4. Tris-buffered saline (TBS): 25 mM Tris–HCl pH 7.4, 150 mM NaCl. 5. Permeabilization solution: 0.2% Triton-X100 in PBS. 6. Blocking solution: 1% bovine serum albumin (BSA) in PBS. 7. Primary antibodies (working solutions stated): Mouse monoclonal anti-ICAM-1 (1 μg/ml) (clone BBIG-I1, R&D Systems, Abingdon, UK) and rabbit polyclonal antiβ-catenin (1–2 μg/ml) (Sigma-Aldrich). Other antibodies described in Notes or Figures: mouse monoclonal antiVE-Cadherin (1–2 μg/ml) (Pharmingen, Lowley, UK),

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goat polyclonal anti-PECAM-1 and rabbit polyclonal anticaveolin-1 N-20 (2 μg/ml) (Santa Cruz, Santa Cruz, CA). 8. Donkey anti-mouse-FITC, donkey anti-rabbit-Cy5 (used in Section 3), donkey anti-goat-Cy5 (additional stainings described in Section 4 and Fig. 5.2), secondary fluorescent antibodies from Jackson ImmunoResearch (Suffolk, UK) at working dilution of 2 μg/ml. 9. Phalloidin-TRITC (P1971) (Sigma-Aldrich) at working dilution of 4 μg/ml. 10. DAKO fluorescent mounting medium (DAKO Ely, UK).

3. Methods 3.1. Cell Culture 3.1.1. Endothelial Cells

3.1.2. T Lymphoblasts

HUVECs and HDMECs require to be plated on an extracellular matrix (ECM) substrate in order to promote proliferation and survival. Flasks are coated with a solution of 10 μg/ml of fibronectin in PBS for 30 min at 37◦ C. Cells are thawed, grown and passed following manufacturer’s instructions (see Note 1). Cells must be used between passages 2 and 5. In further passages cells grow old age and the ratio between paracellular and transcellular diapedesis is altered. Confluent cells are normally passaged by 1:3 dilutions onto fibronectin-coated flasks. Incubation with the trypsin–EDTA solution should not exceed 4–5 min. 1. T-cell extraction from single-donor buffy coats is carried out using a Phycoll procedure. Typically, 20 ml of blood diluted in PBS (1:1) is slowly laid over 10 ml of Phycoll solution and centrifuged for 20 min at 2,000 rpm (brake off). Leucocytes are segregated in a whitish interface that is carefully collected. Leucocytes are then washed twice in PBS by centrifugation for 5 min at 1,200 rpm. 2. T-lymphocyte activation: T cells are cultured in RPMI medium with 10% heat-inactivated human serum at 37◦ C in a 5% CO2 atmosphere. First, RPMI is supplemented with 0.5 μg/ml PHA for 48 h in order to activate and differentiate T lymphocytes into T lymphoblasts. The medium is then replaced with fresh RPMI containing 10% serum supplemented with 10 U/ml of fresh IL-2 instead of PHA. During the following 2 weeks medium containing fresh IL-2 is changed every 48 h. Transmigration assay is performed between 10 and 15 days after PHA stimulation.

Paracellular and Transcellular Diapedesis

3.2. Transmigration Assay 3.2.1. Day 1: Preparation of Cells for the Transmigration Assay. Seeding Endothelial Cells on Glass Coverslips

3.2.2. Day 2: Mimicking Long-Term Inflammation. Endothelial Stimulation with TNF-α

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1. Fibronectin (10 μg/ml) coating of glass coverslips takes longer than plastic, a minimum of 6 h, although it is preferable to perform the coating over night at 37◦ C prior to cell plating. 2. Cells are trypsinized and plated onto fibronectin-treated coverslips in 24-well plates at 1–1.5 × 105 cells per coverslip in 1 ml of complete medium. The medium is changed 8 h later. TNF-α is a cytokine extensively used to mimic long-term inflammation in HUVECs and HDMEC prior to the transmigration assay (10, 11). Treatment of cells with TNF-α stimulates the expression of receptors involved in leucocyte adhesion and the subsequent diapedesis across the endothelium (Note 2). Depending on the endothelial cell type used for the transmigration assay, the protocol of stimulation can differ slightly (Note 3): 1. Confluent HUVECs are washed once with starving medium (see Section 2) and then incubated with 0.5 ml of starving medium per coverslip for 5 h. 2. Freshly prepared TNF-α solution (10 ng/ml in starving medium) is added to the cell monolayer for 18–22 h.

3.2.2.2. Stimulation of HDMVEC Monolayers

1. HDMVEC are washed and stimulated with TNF-α (10 ng/ml) for 18–22 h in microvascular complete medium.

3.2.3. Day 3: Transmigration Assay

1. Prior to the assay, it is recommended to check briefly in the microscope of the culture room that the inflamed endothelial monolayer is properly stimulated and has changed from a cobblestone to an elongated morphology. Endothelial cells should not be kept out of the incubator for long periods to prevent stress (Note 4). 2. T lymphoblasts are counted and centrifuged; 1.5 × 105 cells are resuspended into 25 μl of endothelial starving medium. These T lymphoblasts are added to the endothelial monolayer and the co-culture is placed in the incubator at 37◦ C, 5% CO2 for 12–13 min. 3. Medium is gently removed and cells fixed with PFA 4% (prewarmed at 37◦ C a few minutes before) for immunostaining. The co-culture should not be washed before PFA fixation in order to prevent cell stress. Non-adhered lymphocytes will be washed away during the immunofluorescence procedure.

3.3. Immunofluorescence Assay (Triple Staining)

In order to distinguish transcellular from paracellular diapedesis at least a double fluorescence staining is required. A first staining with a mouse monoclonal antibody will detect the ICAM-1 receptor, which is involved in the firm adhesion of the T lymphoblast

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and the subsequent diapedesis. ICAM-1 staining will enable the visualization of the endothelial cell perimeter and morphology as well as the membrane remodelling that endothelium undergoes in response to leucocyte adhesion and transmigration. This includes the observation of both paracellular and transcellular endothelial passages that are opened during diapedesis. A second staining with a fluorophore-conjugated phalloidin takes advantage of the remarkable differences between the filamentous actin (F-actin) of T lymphoblasts and endothelial cells that make possible to distinguish T-cell morphology within the cell co-culture. This double staining allows the analysis of the different morphological changes that both endothelium and lymphocyte undergo in each step of the transmigration cascade. Finally, a third staining of β-catenin, a component of cell–cell junctions, is included with the aim of examining the integrity of the contacts between endothelial cells, an important parameter to discriminate paracellular from transcellular. 1. The coverslip containing the fixed cells is incubated with PFA 4% at room temperature for 20 min. 2. The coverslips are then washed three times with TBS and incubated on ice for 15 min in the same solution. Tris buffer from TBS will quench the remaining reactive PFA. 3. Fixed cells are permeabilized with TBS/TX100 0.2% for 5 min at 4◦ C. 4. The coverslip is washed three times with cold PBS and then incubated for 15 min at room temperature with blocking solution. 5. During these 15 min, a cocktail of mouse anti-ICAM-1 and rabbit anti-β catenin primary antibodies is prepared in blocking solution (working dilutions in Section 2). 6. The coverslip is incubated on a drop of 40 μl of primary antibody cocktail for 30 min at 37◦ C in a wet chamber (Note 5). The side of the coverslip containing cells must face the antibody solution. 7. During this incubation, a cocktail of donkey anti-mouseFITC (for anti-ICAM-1) and donkey anti-rabbit-Cy5 (for anti-β-catenin) is prepared at the recommended dilutions (Note 6). 8. Coverslip washes: A piece of dry tissue, tweezers and a beaker containing PBS are required. The coverslip is carefully taken with the tweezers and submerged into PBS for an instant; the border of the coverslip is then put in perpendicular contact with the tissue that by capillarity will absorb the excess of liquid. The coverslip is dipped into PBS again. Repeat washes 15 times.

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9. Just after last wash, the coverslip is placed onto a 40 μl drop of secondary antibody and incubated in the wet chamber for 30 min at 37◦ C. 10. The coverslip is washed again and incubated for 20 min at 37◦ in the wet chamber with a 40 μl drop of phalloidin conjugated to TRITC, which has been previously diluted in PBS. 11. Microscope glass slides are labelled for a proper identification of the stained coverslip. Add a drop of mounting medium on the slide. Coverslips are washed and mounted, so cells remain between the coverslip and the slide surfaces. Excess of mounting medium is removed with a piece of tissue and the mounted coverslip is let dry for 1 h at 37◦ C 3.4. Detection of T-Lymphoblast Transmigration by Confocal Microscopy

1. The stained coverslip is first analysed by epifluorescence. ICAM-1 (FITC) and phalloidin (TRITC) stainings are observed in order to focus and localize an area where endothelial cells form a proper monolayer, ICAM-1 expression is high and sufficient lymphocytes have adhered (Fig. 5.1).

Fig. 5.1. Diapedesis in HUVECs. (a) Single projection of a z-stack of six confocal sections of a transmigration assay between T lymphoblasts and HUVECs. Upper panels show a general view acquired with a ×40 objective in an LSM510 confocal system from Zeiss. Lower panels show a threefold magnification of the squared area and a schematic representation of T lymphoblasts on the endothelial monolayer. Dark grey colour represent areas of T lymphoblasts localized at the basal planes and white colour areas localized at apical planes. Ap.: apical; para.: paracellular; Trans.: transcellular. (b) Selected single confocal sections from (a) displayed from the basal or bottom planes to apical or top planes. Bar: 20 μm.

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2. The microscope is then switched to confocal mode and a ×40 oil immersion objective is used in order to get the resolution required. Top panels in Fig. 5.1a show a typical field that can be observed using this lens. Lasers necessary to excite the different fluorophores are switched on and proper filter sets are selected. An expert’s advice may be required in order to prevent artefacts and crosstalk between fluorophores. The confocal software is programmed for acquisition of z-stacks of confocal sections at different planes, comprising from the bottom of the monolayer that is in close contact with the coverslip, to sections where the apical membrane of the endothelium starts fading and only the lymphocytes apically adhered or in the process of transmigration are detected with the phalloidin staining (Figs. 5.1 and 5.2). The confocal software is able to calculate the number and thickness of the optical sections necessary to acquire all the staining information along the z-axis. Set a resolution of 1,024 × 1,024 pixels per section. Acquisition of images from an area of 30–50 confluent endothelial cells is a good compromise between resolution and number of lymphoblast adhered to the endothelium, so a few transmigration events can be detected and analysed in each field. Changing to ×63 or ×100 objectives will enable further

Fig. 5.2. Detail of paracellular diapedesis in HUVECs. (a) Selected z-stack confocal sections and z-stack projection of a T lymphoblast (arrowhead) undergoing paracellular diapedesis. Medial sections (2.4 μm to the bottom) show the disruption of junctional VE-cadherin and PECAM-1 and the formation of a paracellular gap (arrow). (b) Top: Twofold magnification of the squared area in (a) and a projection of the stack in the z–x axis where it can be observed that the uropod remains apical whereas the cell body has already transmigrated between two endothelial cells. Bottom: Schematic representation of top image. Bar: 20 μm.

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analysis of some of the transmigrating cells previously identified. As examples, micrographs of co-cultures of T lymphoblasts with HUVECs or HDMVECs are shown in the top panels of Figs. 5.1a and 5.3, respectively, which are projections of all the confocal planes of the z-stack in one single image. These images contain a mixture of T lymphoblasts adhered to the apical endothelial membrane or already transmigrated, with a minority of lymphocytes undergoing diapedesis (5–10%), since this process takes seconds. Quantitation of these images can provide a good idea of the rate of transmigration, although, in order to score accurately each cell, a further analysis of individual images in the z-stack is required. In this analysis, T lymphoblasts that are apically adhered to the endothelial monolayer and have not transmigrated are observed in the confocal sections that correspond to the apical endothelial membrane, but do not appear at the bottom or the basal images in proximity to the coverslip (Fig. 5.1, right panels, Fig. 5.3, left panels showing single confocal sections). In contrast, transmigrated cells with spread morphology will appear in these basal planes, whereas the endothelium (ICAM-1 staining) will be detected in apical sections above these set of lymphoblasts (Fig. 5.3, left panels showing single confocal sections). 3.4.1. T-Lymphoblasts Diapedesis

A transmigrating lymphocyte exhibits a very particular shape that is easily identified in confocal sections acquired with a resolution similar to the micrographs of Figs. 5.1 and 5.3. A leading edge appears in the basal sections, in contact with the coverslip and beneath the endothelium. As we screen along the stack towards the top or apical planes, the cell body is detected in the medium optical sections and the rear uropod is localized in the same optical sections as apical lymphocytes (Figs. 5.1 and 5.3, confocal sections). If diapedesis is detected at a late stage, the cell body will appear under the endothelial monolayer (Fig. 5.2). However, in order to score the cell as undergoing diapedesis, the uropod must always be found at the top planes of the stack above any ICAM-1 staining belonging to endothelial cells. ICAM-1 staining reveals the formation of gaps or pores in many of the transmigratory events (Figs. 5.1 and 5.3). Some of these pores will be surrounded by microvilli-like structures that have been called transmigratory cups or docking structures (Fig. 5.3, enlarged area (2)) (10, 13, 14).

3.4.2. Paracellular Versus Transcellular Diapedesis

Most of T lymphoblasts undergoing diapedesis follow a paracellular route between two cells (more than 90% in HUVECs, around 70% in HDMVECs). Cells following a paracellular route

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Fig. 5.3. Transcellular diapedesis in HDMVECs. Upper left panels show a single projection of z-stack of six confocal sections of a transmigration assay between T lymphoblasts and HDMVECs. Arrowhead show remodelling of ICAM-1 and caveolin-1 upon diapedesis. Lower left panels: Selected single confocal sections from the squared area magnified 2.5-fold and displayed from the basal or bottom plane to the apical or top plane. Top right panel is a schematic representation of the squared area (1) where apical and transmigrated T lymphoblasts as well as T lymphoblasts undergoing transcellular diapedesis can be observed. A T lymphoblast that may initiate diapedesis is also pointed. Bottom right panels: Tenfold enlargement of the squared area (2) where another T lymphoblast is following transcellular diapedesis. Note the formation of ICAM-1-enriched microvilli-like structures and the accumulation of caveolin-1 in the pore.

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will disrupt cell–cell junctions in order to transmigrate between two endothelial cells (Figs. 5.1 and 5.2), whereas junctions will remain intact when T lymphoblasts are following transcellular transmigration (Fig. 5.1). So, although diapedesis can be simply observed in a double staining with anti-ICAM-1 antibodies and phalloidin, it is also important to check the integrity of the junctions during transmigration by analysing β-catenin distribution in z-stack projections (Fig. 5.1). Indeed, if the T lymphoblast is transmigrating close to the endothelial cell border but the junctional integrity is not affected, then diapedesis is scored as transcellular. In case of doubt, the event is not quantified. Some other junctional markers involved in paracellular and transcellular diapedesis, such as VE-cadherin, PECAM1 or caveolin-1, can be analysed by immunostaining to further complete the transmigration analysis (Note 7) (Figs. 5.2 and 5.3).

4. Notes 1. Thawing endothelial cells. It is preferable to seed thawed endothelial cells directly from the vial onto the fibronectincoated flask and avoid centrifugation. Typically, the content of one vial containing 5 × 105 cells in 1 ml of freezing solution can be plated on a 75 cm2 flask containing 15 ml of complete medium. Medium containing diluted DMSO can be removed once cells have adhered to the flask. Both Lonza (HUVECs) and Promocell (HDMEC) provide media with growth factors and low serum content. This guarantees more stable growing conditions than media with higher percentage of serum. Importantly, these commercial media are compatible with transfection reagents such as Fugene, lipofectamine or oligofectamine (when antibiotics are removed), enabling the experimental modulation of the expression of molecules using expression vectors or interfering RNA. 2. Leucocytes are able to transmigrate through resting endothelial monolayers in order to conduct immunosurveillance in absence of any proinflammatory stimulation. This trafficking occurs at low rate and, at least in vitro, follows almost exclusively a paracellular route of TEM. In order to mimic inflammation in vitro, several cytokines such as TNF-α, IL-1β or IFN-γ can be used. TNF-α induces signalling pathways that change the morphology of endothelial cells and induce the expression of many receptors involved

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in leucocyte adhesion and transmigration, particularly transcellular diapedesis. ICAM-1 and VCAM-1, which interact with leucocyte β2 and β1 integrins, respectively, and mediate leucocyte firm adhesion, are expressed at the endothelial surface between 4 and 24 h post stimulation. Since signalling mediated by these receptors regulates the subsequent stage of diapedesis, it is important to induce their maximal expression. In addition, TNF-α also induces endothelial cell elongation that starts between 4 and 6 h post stimulation and reaches its maximum at 24 h. This morphological remodelling controls several elements involved in diapedesis, such as preferential areas of leucocyte transmigration or cell-tocell junctions. Therefore, if a scenario of long-term inflammation needs to be reproduced in vitro, confluent endothelial cells on the coverslip must be stimulated between 4 and 24 h in order to promote a proper cell elongation as well as maximal expression of endothelial adhesion receptors. On the other hand, for a proper in vitro leucocyte TEM analysis, it is necessary to generate a confluent monolayer with intact junctional complexes. Time course analysis of AJ in the laboratory has shown that endothelial cells require at least 24 h to generate mature AJ. After 48 h, optimal junctional integrity is obtained. This time frame is compatible with functional experiments of transient transfection of DNA vectors or siRNA. Plating cells at confluence for 36 h followed by stimulation with TNF-α for 20–24 h prior to the transmigration assay is thus a reasonable timing for most experiments. 3. Media for TNF-α stimulation. HUVECs and HDMEC media is provided with different growth factors such as VEGF, FGF or HGF. In the case of HUVECs, rate of total leucocyte transmigration is not significantly altered by the presence or absence of these growth factors. However, increased transcellular diapedesis is observed when HUVECs are stimulated with TNF-α in medium complemented only with 1% foetal bovine serum but no growth factors (starving medium), so we normally use this medium. For HDMEC, a good transcellular diapedesis is observed when cells are stimulated with TNF-α in the complete medium provided by the manufacturer and no starving medium is required. 4. Due to their role in the regulation of the vascular homeostasis, endothelial cells are highly sensitive to mechanical or temperature stress. This has a direct effect on protein complexes involved in cell-to-cell junctions or leucocyte transmigration like filamentous actin, AJ, PECAM-1 or caveolin-1.

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In order to prevent artefactual endothelial cell remodelling before the addition of T lymphoblasts, it is preferable not to wash the endothelial monolayer. The transmigration assay can be carried in the medium containing TNF-α, since this cytokine is unstable and will be degraded at the time of the assay. 5. How to make a wet chamber. Take a plastic culture dish (10 or 15 cm diameter) and lay inside a piece of wet tissue, and onto the tissue a piece of parafilm. Add a drop of 40 μl of antibody solution to the parafilm. Place the coverslip on the drop orientating the side containing cells towards the antibody solution. Close the chamber and incubate at 37◦ C. 6. It is recommended to stain ICAM-1 and F-actin with fluorophores that are excited in the range of 488–550 nm wavelengths, respectively, because their emission wavelength can be detected by the human eye by epifluorescence. The observer will then be able to screen quickly the coverslips through the eyepieces and perform initial localizations and quantitations. It is convenient to perform the third staining of cell–cell endothelial junctions or other molecules of interest with secondary antibodies conjugated to fluorophores in the range of excitation of 633 nm of wavelength, like Cy5. These fluorophores can be properly excited and analysed by the confocal system, but not subjected to a previous epifluorescence screening by the observer, since human eyes cannot detect such long-emission wavelengths (longer than 650 nm). 7. Other proteins involved in cell–cell junctional remodelling during diapedesis that can be included in the third immunostaining are VE-cadherin and PECAM-1 (Fig. 5.2). VE-cadherin forms endothelial AJ that are disrupted upon leucocyte passage during TEM (15). Parajunctional PECAM-1 facilitates diapedesis (16), although it often appears dispersed in response to TNF-α. PECAM-1 has been recently involved in transcellular TEM (9). Alternatively, antibodies that recognize lymphoblast surface proteins playing a role in adhesion and diapedesis, such as β1 or β2 integrins, will label T lymphoblasts exclusively and will facilitate the visualization of transmigrating cells (17). Finally, since transcellular diapedesis occurs preferentially through caveolae-enriched areas, a staining of the scaffolding protein caveolin-1, particularly in microvascular endothelial cells, will surround many of the transcellular pores (Fig. 5.3). References of good antibodies for immunofluorescence are included in Section 2.

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Acknowledgments E.C.M. is supported by a contract from the Spanish Juan de la Cierva program. S.G. is supported by a fellowship from the Spanish Ministry of Science and Education. J.M. was supported by a British Heart Foundation intermediate fellowship (no. FS/04/006), grant SAF2008-1936, and a contract from the Spanish Ramón y Cajal program. References 1. Butcher EC. (1991) Leukocyte-endothelial cell recognition: three (or more) steps to specificity and diversity. Cell 67, 1033–6. 2. Engelhardt B, Ransohoff RM. (2005) The ins and outs of T-lymphocyte trafficking to the CNS: anatomical sites and molecular mechanisms. Trends Immunol 26, 485–95. 3. Libby P. (2002) Inflammation in atherosclerosis. Nature 420, 868–74. 4. Millán J, Ridley AJ. (2005) Rho GTPases and leucocyte-induced endothelial remodelling. Biochem J 385, 329–37. 5. Cho Y, De Bruyn P P. (1986) Internal structure of the postcapillary high-endothelial venules of rodent lymph nodes and Peyer’s patches and the transendothelial lymphocyte passage. Am J Anat 177, 481–90. 6. Faustmann PM, Dermietzel R. (1985) Extravasation of polymorphonuclear leukocytes from the cerebral microvasculature. Inflammatory response induced by alpha-bungarotoxin. Cell Tissue Res 242, 399–407. 7. Muller WA. (2003) Leukocyte-endothelialcell interactions in leukocyte transmigration and the inflammatory response. Trends Immunol 24, 327–34. 8. Feng D, Nagy JA, Pyne K, Dvorak HF, Dvorak AM. (1998) Neutrophils emigrate from venules by a transendothelial cell pathway in response to FMLP. J Exp Med 187, 903–15. 9. Carman CV, Sage PT, Sciuto TE, de la Fuente MA, Geha RS, Ochs HD, Dvorak HF, Dvorak AM, and Springer TA. (2007) Transcellular diapedesis is initiated by invasive podosomes. Immunity 26, 784–97. 10. Carman CV, Springer TA. (2004) A transmigratory cup in leukocyte diapedesis both through individual vascular endothelial cells and between them. J Cell Biol 167, 377–88.

11. Millán J, Hewlett L, Glyn M, Toomre D, Clark P, Ridley AJ. (2006) Lymphocyte transcellular migration occurs through recruitment of endothelial ICAM-1 to caveolaeand F-actin-rich domains. Nat Cell Biol 8, 113–23. 12. Carman CV, Springer TA. (2008) Transcellular migration: cell-cell contacts get intimate. Curr Opin Cell Biol 20, 533–540. 13. Barreiro O, Yanez-Mo M, Serrador JM, Montoya MC, Vicente-Manzanares M, Tejedor R, Furthmayr H, Sanchez-Madrid F. (2002) Dynamic interaction of VCAM-1 and ICAM-1 with moesin and ezrin in a novel endothelial docking structure for adherent leukocytes. J Cell Biol 157, 1233–45. 14. Wojciak-Stothard B, Williams L, Ridley AJ. (1999) Monocyte adhesion and spreading on human endothelial cells is dependent on Rho-regulated receptor clustering. J Cell Biol 145, 1293–307. 15. Allport JR, Ding H, Collins T, Gerritsen ME, Luscinskas FW. (1997) Endothelialdependent mechanisms regulate leukocyte transmigration: a process involving the proteasome and disruption of the vascular endothelial-cadherin complex at endothelial cell-to-cell junctions. J Exp Med 186, 517–27. 16. Mamdouh Z, Chen X, Pierini LM, Maxfield FR, Muller WA. (2003) Targeted recycling of PECAM from endothelial surfaceconnected compartments during diapedesis. Nature 421, 748–53. 17. Shaw SK, Ma S, Kim MB, Rao RM, Hartman CU, Froio RM, Yang L, Jones T, Liu Y, Nusrat A, Parkos CA, Luscinskas FW. (2004) Coordinated redistribution of leukocyte LFA-1 and endothelial cell ICAM-1 accompany neutrophil transmigration. J Exp Med 200, 1571–80.

Chapter 6 Monitoring RhoGTPase Activity in Lymphocytes Marouan Zarrouk, David Killock, and Aleksandar Ivetic Abstract L-selectin is a cell adhesion molecule (CAM) that is essential for the tethering and subsequent rolling of naïve lymphocytes along the luminal wall of postcapillary venules entering lymph nodes. As with many CAMs, L-selectin has the capacity to transduce intracellular signals in response to ligand binding. This implicates CAMs involved in tethering and rolling as contributors to intracellular signals that lead to the transition from rolling to arrest. In addition, studies in L-selectin-null mice have also revealed a role for L-selectin in chemokine-directed cell migration of leucocytes in tissues. The Ras homology (Rho) family of small GTPases are intracellular proteins that respond to signals received from the surrounding environment. The RhoGTPases typically activate downstream targets involved in the remodelling of the actin cytoskeleton, which is essential for continued progression through the multi-step adhesion cascade. This chapter will focus on how to prepare, perform and monitor RhoGTPase activation assays in response to L-selectin stimulation. Although this section focuses on L-selectin stimulation, the methods outlined here can be applied to analysing RhoGTPase activity in response to stimulating other receptors involved in tethering/rolling such as CD44, P-selectin glycoprotein ligand-1 and E-selectin ligand-1. Key words: L-selectin, leucocyte, lymphocyte, cytoskeleton, RhoA, Rac1, Cdc42, pull-down assay.

1. Introduction During the multi-step adhesion cascade lymphocytes undergo highly dynamic changes in cell shape and adhesiveness, turning from spheroid-like blood-borne cells that use low-affinity CAMs (such as L-selectin) into amoeboid-like transmigratory cells that use high-affinity CAMs (such as integrins) (1). The intracellular mechanisms underlying this transition are varied and of immense interest, but are currently poorly understood. Such changes occur in response to extracellular signals from the F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_6, © Springer Science+Business Media, LLC 2010

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microenvironment, which are typically sensed by single- or multipass transmembrane receptors, such as CAMs and chemokine receptors, respectively. L-selectin has been shown to induce intracellular signals that are both distinct and similar to signalling downstream of chemokine receptors (2). Stimulation of both L -selectin and chemokine receptors has been shown to modulate the activity of the RhoGTPase, Rac (3, 4). However, the methods used to study Rac activation in response to Lselectin stimulation are long and indirect, and thus do not conform to a currently accepted practise. Most RhoGTPases shuttle between active and inactive conformations and are dictated by binding to guanosine triphosphate (GTP) or guanosine diphosphate (GDP), respectively (5) (see Fig. 6.1). In their active state, RhoGTPases bind to a multitude of downstream effector targets. The specificity and duration of RhoGTPase binding to downstream targets is modulated by a number of upstream regulators, some of which include GTPase-activating proteins (or GAPs), guanosine nucleotide exchange factors (GEFs) and RhoGTPase dissociation inhibitors (RhoGDIs). The catalytic conversion of GTP to GDP is very slow in most RhoGTPase family members, which is dramatically increased by GAPs and

GTP

GDP

GEF

RhoGDP

GDI

RhoGTP

Out

In

Plasma membrane

Cytosol

P Fig. 6.1. Illustration of a RhoGTPase cycling between active and inactive states, which are influenced by guanine nucleotide exchange factors (GEFs), GTPase dissociation inhibitors (GDIs) and GTPase-activating proteins (GAPs). Active RhoGTPases are often associated with the inner leaflet of the plasma membrane and can be removed from these intracellular domains by the binding of GDIs to the isoprenyl group. Both GAPs and GEFs can be associated with the plasma membrane, but is not shown in this illustration.

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subsequently limits binding to downstream effector targets. A small number do not catalyse GTP, for example RhoE (6), and are constitutively active in cells. In contrast, GEFs facilitate the exchange of GDP for GTP, restoring the activity of RhoGTPases. RhoGDIs are involved in sequestering RhoGTPases in the cytoplasm by binding to an isoprenyl group located at the C terminus of RhoGTPases (7). Here, we will outline approaches to monitor the activities of the canonical RhoGTPases: RhoA, Rac1 and Cdc42 in response to L-selectin stimulation. The experiments outlined in this chapter will allow for the analysis of other cell adhesion molecules involved in leucocyte rolling, such as CD44, PSGL-1 and ESL-1.

2. Materials 2.1. Cell Lines

1. Murine 300.19 pre-B cells are used in these studies, although primary lymphocytes purified from whole blood can also be used. The pre-B cells have been transfected to stably express wild-type (WT) or mutant forms of L-selectin, which has been described previously (8).

2.2. Cell Culture

1. Roswell Park Memorial Institute (RPMI)-1640 (Invitrogen) containing pyruvate supplemented with 10% foetal calf serum (FCS), L-glutamine and 5 mM penicillin/streptomycin. 2. Tissue culture incubator set at 5% CO2 and 37◦ C with humidifying condition. 3. Cell counting equipment; phase contrast light microscope, counter and haemocytometer. 4. Thirty percent (v/v) sterile bovine serum albumin solution (Sigma-Aldrich).

2.3. Protein Expression and Purification of GST-Fused Downstream Effector Targets of RhoA, Rac1 and Cdc42

1. Glycerol stocks of Escherichia coli BL-21 (genotype: B F- dcm+ Hte ompT hsdS(rB - mB -) gal l (DE3) [pLysS Camr ]a endA Tetr ) transformed with expression plasmids containing the open-reading frames of glutathione-Stransferase (GST) fused to downstream effector domains of Rho GTPases (plasmids of PAK-PBD, WASP-CRIB-C and Rhotekin-C21 were kindly provided by John G Collard, the Netherlands Cancer Institute, Amsterdam, the Netherlands). 2. Prokaryotic protein expression plasmids (pGEX – GE Healthcare) containing open-reading frames of the

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downstream effector binding sites for RhoA, Rac1 and Cdc42 (i.e. Rhotekin, PAK and WASP CRIB domain, respectively) fused to GST. 3. Luria-Broth pellets (Sigma-Aldrich) dissolved in an appropriate amount of sterile water and autoclaved for 30 min. 4. Cooled benchtop centrifuge, for example, AllegraTM 6R R centrifuge, Beckman Coulter , UK. 5. Conical flasks (approximately 300 mL size) for growth of bacteria and orbital shaker with variable temperature settings (i.e. 37 and 30◦ C). 6. Isopropyl β-D-1-thiogalactopyranoside (IPTG, SigmaAldrich). 7. STE buffer: 10 mM Tris–HCl, pH 8.0, 150 mM NaCl, 1 mM EDTA. 8. Phenylmethanesulphonylfluoride (PMSF, Sigma-Aldrich). 200 mM stock solution to be stored at –20◦ C. 9. Hyperdermic needle, 19 gauge (Kendall, UK). 5-mL syringe (Sherwood). 10. Lysozyme (Sigma-Aldrich) for lysis of bacteria. 11. Glutathione sepharose 4B beads (GE Healthcare), preequilibrated in STE buffer. 12. Fixed concentration of BSA (1 μg/μL) to use as a standard for establishing approximate concentrations of purified GST-fused protein. 2.4. Cell Stimulation

1. Plastic tissue culture dishes; round triple vent 18-mm diamR eter (Greiner , Germany). 2. Water bath set to 37◦ C (for incubating 1.5 mL tubes). 3. DREG56 IgG1 monoclonal antibody (specifically recognises human L-selectin). This can be purchased from a number of commercial suppliers, for example, Santa Cruz R Biotechnology .

2.5. RhoGTPase Activation Assays

1. Cell lysis buffer: 10 mM MgCl2 , 1 mM EDTA, pH 8.0, 25 mM HEPES pH 7.0, 150 mM NaCl; 2% (v/v) glycerol, 1% (v/v) Triton X-100, 1 mM Na3 VO4 ; 50 nM NaF, 25 nM calyculin A. Make fresh and keep on ice. 2. Glutathione sepharose 4B beads (GE Healthcare), bound to recombinant purified effector binding domain of PAK (for Rac1), WASp CRIB (for Cdc42) and Rhotekin (for RhoA) fused to GST. 3. Tube rotator placed in the cold room.

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4. Cool microcentrifuge tubes (1.5 mL) to 4◦ C. A cooled microcentrifuge that can be set to 4◦ C or alternatively perform the assay in a cold room set to 4◦ C. 2.6. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE)

R 1. NuPAGE Novex pre-cast gradient (4–12%) Bis-Tris gels (Invitrogen): Ten wells, 1.5 mm thick. Manually cast gels can be used, but ensure that the appropriate percentage of polyacrylamide is used to resolve 20 kDa proteins, which is the average molecular weight of RhoGTPases. Thicker gels allow for greater loading of samples (e.g. up to 50 μL of sample for 1.5 mm thick and up to 30 μL of sample for 1.0 mm thick gels). R 2. Xcell SureLock gel electrophoresis tank (Invitrogen). This tank accommodates a maximum of two gels.

3. Electrophoresis buffers are purchased from Invitrogen for running gels and are sold as a 20x concentrated stock solution. R pre-stained molecular weight standards 4. Novex Sharp (Invitrogen).

5. Protein gel loading buffer (4x) according to UK Laemmli: 2.4 mL 1 M Tris pH 6.8, 0.8 g SDS stock, 4 mL 100% glycerol, 0.01% bromophenol blue, 1 mL β-mercaptoethanol (electrophoresis grade), 2.8 mL water. Dilute protein loading buffer 50:50 with purified water to obtain 2x protein loading buffer. 2.7. Western Blotting

1. Xcell II blot module (Invitrogen). Other blotting apparatuses can be used (semi-wet or dry). Ensure that the correct transfer times are used for other methods of protein transfer. Transfer solution is bought from Invitrogen and is supplied as a 20x stock solution. Ensure to add 10% methanol to 1x transfer buffer and increase to 20% when transferring two gels for every blot module. 2. Polyvinylidene fluoride (PVDF) transfer membrane (Millipore). Soak the membrane in neat methanol prior to use and equilibrate back into transfer buffer before layering onto polyacrylamide gel. 3 MM Whatman chromatography paper (Maidstone, UK). 3. Tris-buffered saline (TBS): 20 mM Tris–HCl (pH 7.6), 150 mM NaCl. 4. Supplement TBS with 0.1% (v/v) non-ionic detergent (e.g. Triton-X100 or Nonidet P-40) for washing PVDF membranes (TBST/N). 5. Blocking buffer: 5% (w/v) semi-skimmed powdered milk dissolved in TBS.

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6. Goat anti-mouse secondary antibody conjugated to HRP (Dako). 7. Chemiluminescent solution (Western LightningTM Chemiluminescence reagent, PerkinElmer LAS Inc., USA), Saran R , Japan) for the development of Wrap, X-ray film (Fuji membranes. 2.8. Quantification and Statistical Analysis

R 1. ImageJ downloaded gov/ij/download.html

free

from

http://rsb.info.nih.

R 2. GraphPad Prism used for data handling and statistical analysis (unpaired, two-tailed t-test).

3. Methods Assaying for RhoGTPase activity can be notoriously difficult to master and focussed attention towards minimising a number of variables can help achieve reproducible results. First, one of the major reasons for variable results in RhoGTPase assays is that samples are not kept sufficiently cold after cell lysis. Membrane disruption with non-ionic detergent-based lysis buffers solubilises the RhoGTPases, which can lead to their rapid inactivation by interacting with GAPs. This can increase with rising temperatures. Therefore, after cell lysis, it is critical that all subsequent steps are kept sufficiently cold. Second, the type of cells that will be used for such assays must also be treated uniquely. For example, 300.19 cell line undergoes rapid cell division under optimal growth conditions. Many foetal calf serum components can interfere with RhoGTPase signalling (such as lysophosphatidic acid, which activates RhoA in some cell types (9)). It may, therefore, be necessary to “starve” cells free from serum-derived factors, by incubating cells in very low FCS or completely without FCS for a few hours or overnight. Third, preparing GST-fused RhoGTPase baits on the day of the assay is essential for obtaining maximal binding of the RhoGTPase under study. Finally, the method used to express and prepare recombinant GST-fused bait for GTP-loaded RhoGTPases is of great importance. We have found that isolating and purifying bait protein from E. coli using mild cell disruption techniques (see Section 3.2) increases binding between bait protein and GTP-loaded RhoGTPase. 3.1. Maintenance of Cell Culture

1. Murine 300.19 pre-B cell lines stably expressing WT L -selectin were cultured in RPMI-1640 medium containR ing 5 mM L-glutamine (Gibco Invitrogen, Paisley, UK) supplemented with 10% FCS; 100 U/mL penicillin and

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100 μg/mL; streptomycin and 50 μM β-mercaptoethanol. This cell line has been co-transfected with pBabe vector containing the puromycin resistance marker. Therefore, supplement culture medium with 3 μg/mL puromycin to maintain selection pressure. 2. Dramatic changes in cell density can have a profound effect on L-selectin expression levels, therefore ensure that cells are maintained at a density of 0.1–0.5 × 106 cells/mL R in cell culture flasks T75 or T175 (Corning , UK) in standard mammalian cell culture incubators with 5% (v/v) CO2 /atmospheric air and under humidified conditions (see Note 1). Peripheral blood lymphocytes isolated from whole blood can be used instead of cell lines and are kept under similar conditions (without puromycin selection). 3.2. Protein Expression and Purification of GST-Fused Baits for RhoA, Rac1 and Cdc42

1. Innoculate 30 mL of LB-medium (containing 0.1 mg/mL ampicillin) with glycerol stock of BL21 E. coli transformed with pGEX vector harbouring the RhoGTPase effector domain fused to GST and culture overnight at 37◦ C under aerobic agitation. 2. The next day, dilute overnight culture 1:20 into 100-mL LB-medium containing 0.1 mg/mL ampicillin and incubate at 37◦ C under agitation until an OD600 of approximately 0.8 is achieved. This should take between 2 and 3 h. On average, 30 mL of overnight culture is used to generate 600 mL of culture for protein expression, and 300 mL will be used for a single assay. 3. Induce protein expression by adding 0.5 mM IPTG (final concentration) for 2.5 h at 30◦ C (see Note 2). 4. Aliquot bacterial culture into 12 × 50 mL Falcon tubes and centrifuge for 30–45 min at 3,000 rpm at 4◦ C to harvest bacteria into pellets. 5. Pour off supernatant and vacuum aspirate remaining liquid. Freeze pellets in Falcon tubes overnight at –80◦ C to weaken bacterial cell walls prior to cell lysis step (see Note 3). 6. Remove 6 of the 12 Falcon tubes containing bacterial pellets from –80◦ C and thaw at room temperature. The remaining six tubes can be used for a later assay. Resuspend the pellets in 4.5 mL (total volume) of ice-cold STE buffer supplemented with 1 mM PMSF (make fresh every time) and homogenise by repeated passage through a 19-gauge hyperdermic needle attached to a 5-mL syringe. Ensure that the cell suspension is visibly free from any aggregates whilst swirling tube.

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7. Mix the suspension gently with 100 μg/mL of lysozyme and incubate for 15 min on ice. 8. Add 5 mM DTT, 1% (v/v) Tween-20 and 0.03% (v/v) SDS to the lysate. A change in the viscosity is seen at this point, which is a good indicator of cell lysis. Aliquot lysate into 1.5-mL tubes and centrifuge for 45 min at 14,000 rpm, using a cooled microcentrifuge set at 4◦ C. 9. Mix supernatant with 200 μL of glutathione sepharose R beads that have been pre-equilibrated in STE buffer 4B and incubate for 1 h at 4◦ C under rotation. 10. Wash beads three times with STE buffer at 4◦ C and add 200 μL of STE buffer to generate a final volume of 400 μL 50% bead slurry. Beads should be used within 24 h of preparation. 11. Assess protein yield of GST-fused product using polyacrylamide gel electrophoresis followed by Coomassie Blue staining. Resolve 5 μL of glutathione beads containing the GST-fused product in a single lane. Use the remaining lanes to resolve increasing amounts of BSA standard (generated from crystalline BSA). A range of 3–30 μg of BSA is normally loaded as shown in Fig. 6.2.

Fig. 6.2. Determining protein concentration of GST-fused effector domain bound to glutathione sepharose beads. Increasing amounts of BSA (3, 12, 18, 24, 27 and 30 μg) are resolved in the first six lanes of the polyacrylamide gel. The concentration range normally loaded is between 3- and 30-μg BSA. M = molecular weight standards (from bottom to top: 15, 20, 40, 50, 60, 80 and 110 kDa). B = 5 μL of glutathione beads boiled and loaded onto the last lane of the gel. Coomassie staining of the polyacrylamide gel reveals that 5 μL of beads prepared in this example carries the equivalent of approximately 25 μg of protein.

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1. Harvest cell lines from overnight culture by centrifugation and resuspend in starving medium (RPMI-1640 medium; 3 μg/mL puromycin; 100 U/mL penicillin; 100 μg/mL streptomycin and 50 μM β-ME). Plate cells at a density of 1 × 106 cells/mL into 18 mm (or 10 mm) dishes. Each assay requires 1 × 107 cells. Therefore, as a guide, add 1 × 107 cells per 10 mm dish and 2 × 107 cells per 18 mm dish. 2. Agitate starved cells from round plastic dishes by gentle pipette action using a 10-mL pipette. Harvest cells by mild centrifugation and resuspend in 1% (v/v) BSA/RPMI1640 to a final cell density of 1 × 107 cells/mL. Place 1 mL of cells into a 1.5-mL tube. 3. Add 4 μg of DREG56 or mouse IgG1 isotype control antibody to cells and incubate at room temperature for 1 min and invert tube twice during this period to mix the antibody with cells. 4. Incubate tubes in 37◦ C water bath for the required amount of time, inverting tubes occasionally. The time course employed in our experiment is for 0, 5, 10 and 20 min. Stagger time points so that the longest time point is started first and the shortest last. 5. Place tubes into cooled microcentrifuge set at 4◦ C and spin for 1 min at 5,000 rpm. 6. Vacuum aspirate supernatant from tube and resuspend pellet in 1 mL of ice-cold lysis buffer. Return tubes back to cooled microcentrifuge and spin immediately at 14,000 rpm for 10 min. This step incorporates both cell lysis and centrifugation. 7. Remove 100 μL of clarified lysate and place into a fresh tube containing equal amount of 2x protein loading buffer. This sample will be used for determining total levels of RhoGTPase within a given cell lysate. 8. Place the remaining lysate into fresh tubes containing approximately 100 μg of recombinant GST-fused bait bound to glutathione beads, which equates to approximately 20 μL of beads per tube. 9. Incubate bead/lysate mixture for 1 h under rotation at 4◦ C (in cold room). 10. Wash beads three times in cell lysis buffer and add 20 μL of 2x protein loading buffer. 11. Heat all samples on a heating block at 95◦ C for 5 min. Allow tubes to cool prior to loading and resolving on polyacrylamide gels.

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3.4. SDS Polyacrylamide Gel Electrophoresis

1. The method set out in this section is based on using the Novex (Invitrogen) pre-cast gel system. This system is extremely user friendly and pre-made buffers can be purchased from the same company. 2. Remove 10-well, 1.5 mm thick, 4–12% gradient gel from plastic packaging. Peel off tape at the foot of the gel. Remove 10-well comb and equilibrate wells by filling and emptying with running buffer (using a 5-mL pipette). 3. Mount the pre-cast gel into the electrophoresis tank and, using thin rounded gel loading tips, load 20 μL of whole cell lysate sample into each well. For loading of proteins bound to sepharose beads, spin the tubes once they have been boiled and load the soluble fraction into the wells (leaving beads in the tube). Using gel loading tips will exclude the entry of sepharose beads into pipette tip, making it easier to take up the soluble fraction without contaminating your sample with beads. 4. Close the tank and run on a constant 200 V for approximately 45 min, or when the blue dye front begins to emerge from the foot (bottom) of the pre-cast gel. 5. Remove gel from tank and crack open the plastic casing using an opener provided by the manufacturer.

3.5. Western Blotting

1. Remove wells and foot from gel, and place into a plastic container. Ensure that the plastic container is large enough to hold 1 L of 1x transfer buffer and deep enough to submerge a stack of 6–7 sponges, PVDF transfer membrane and 3 MM Whatman paper that has been cut slightly larger than the gel. 2. Place one soaked sponge on the base of the transfer tank. Then place one sheet of soaked Whatman paper, followed by the gel (make note of the orientation of the gel and exclude any air bubbles) and then by the PVDF membrane. Place one more sheet of Whatman and then add soaked sponges one by one. Seal the stack by pressing lid onto the base and insert the module into the tank. Run for approximately 2 h using a constant power of 25 V. 3. Remove membrane from transfer tank and block in 5% (w/v) powdered milk dissolved in TBS for 1 h at room temperature. 4. Add antibody of interest at 1:1,000 in blocking solution and incubate at 4◦ C with continuous agitation overnight. 5. The next day, remove milk/antibody solution and wash twice with TBS for 5 min each and once with TBSN for 5 min in between the detergent-free washes.

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6. Block the membrane for a further 30 min and subsequently add HRP-conjugated secondary antibody diluted 1:2,000 in blocking solution for 1 h at room temperature. 7. Repeat washes as in Step 5. Remove membrane with tweezers, hold vertically and lightly tap side of membrane onto tissue to remove most of the TBS. Then layer the membrane (face side down) onto a 1-mL mixture of chemiluminescent solutions A and B that have been placed onto a sheet of Saran Wrap and incubate for about 1 min. 8. Lift the membrane with tweezers and remove excess chemiluminescent solution as before and layer the membrane as flat as possible onto a fresh sheet of Saran Wrap and cover the membrane by folding over. 9. Immunodetect RhoGTPase of interest by exposing WestR ern blot to X-ray film (Fuji , Japan) and develop automatically (Xograph, UK). 10. Ensure that the signal is not overexposed, so that bands can be quantified more easily. A good example of band intensity is shown in Fig. 6.3a.

Fig. 6.3. Cdc42 activity decreases in response to stimulation of WT L-selectin. (a) Representative Western blot reveals the relative levels of GTP-bound Cdc42 (upper panel) and total levels of Cdc42 detected from a fraction of the whole cell lysate (lower panel). Both sets of bands were subjected to densitometry using ImageJ software. (b) Histogram depicting the relative changes in fold activity of Cdc42 in response to L-selectin stimulation over time.

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3.6. Quantification and Analysis

1. Scan X-ray film using a standard image scanner (for examR ple, Canon , UK), save image as JPEG or TIFF file and open the scanned image file through the ImageJ application. R Other applications can be used, such as Adobe Photoshop, to obtain densitometric data. 2. Draw square/rectangular objects around each band to be analysed. Select the largest band first as each subsequent band will be monitored using the same-sized square/rectangle. When selecting a band, ensure that all sides of the band do not meet the edges of the square/rectangle. This ensures that the background signal is also incorporated into the quantification. 3. Obtain pixel values for each band derived from scans of GTP-bound and total GTPase. 4. For every time point, calculate the ratio of GTP-bound GTPase over total GTPase. 5. Assign the value “1” to the ratio obtained for 0 min and normalise all ratios from subsequent time points against 0 min. Arrange data in the form of a histogram as shown in Fig. 6.3b.

4. Notes 1. Reducing the temperature from 37◦ C to 30◦ C during IPTG induction results in higher soluble protein yield. Protein yield may be slightly reduced at lower temperatures, but this is outweighed by decreasing the chances of obtaining inclusion bodies in your preparation. 2. Bacterial cell pellets can be left frozen for up to 3 months at –80◦ C. 3. Avoid using tissue culture flasks for the starvation step. Cells seem to adhere more avidly to the plastic of flasks than tissue culture dishes. References 1. Ley K, Laudanna C, Cybulsky MI, Nourshargh S. (2007) Getting to the site of inflammation: the leukocyte adhesion cascade updated. Nat Rev Immunol 7, 678–89. 2. Zarbock A, Ley K. (2008) Mechanisms and consequences of neutrophil interaction with the endothelium. Am J Pathol 172, 1–7.

3. Brenner B, Weinmann S, Grassme H, Lang F, Linderkamp O, Gulbins E. (1997) L-selectin activates JNK via src-like tyrosine kinases and the small G-protein Rac. Immunology 92, 214–9. 4. Nijhara R, van Hennik PB, Gignac ML, Kruhlak MJ, Hordijk PL, Delon J, Shaw S. (2004) Rac1 mediates collapse of microvilli

Monitoring RhoGTPase Activity in Lymphocytes on chemokine-activated T lymphocytes. J Immunol 173, 4985–93. 5. Ivetic A, Ridley AJ. (2004) Ezrin/radixin/ moesin proteins and Rho GTPase signalling in leucocytes. Immunology 112, 165–76. 6. Riento K, Guasch RM, Garg R, Jin B, Ridley AJ. (2003) RhoE binds to ROCK I and inhibits downstream signaling. Mol Cell Biol 23, 4219–29. 7. Olofsson B. (1999) Rho guanine dissociation inhibitors: pivotal molecules in cellular signalling. Cell Signal 11, 545–54.

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8. Ivetic A, Florey O, Deka J, Haskard DO, Ager A, Ridley AJ. (2004) Mutagenesis of the ezrin-radixin-moesin binding domain of L-selectin tail affects shedding, microvillar positioning, and leukocyte tethering. J Biol Chem 279, 33263–72. 9. Ridley AJ, Hall A. (1992) The small GTPbinding protein rho regulates the assembly of focal adhesions and actin stress fibers in response to growth factors. Cell 70, 389–99.

Chapter 7 Visualisation of Signalling in Immune Cells Leo M. Carlin, Konstantina Makrogianneli, Melanie Keppler, Gilbert O. Fruhwirth, and Tony Ng Abstract Currently, a great number of approaches are employed in investigation of the immune system. These range from experiments in live animals and biochemical techniques to investigate whole organs or cell populations down to single cell and molecular techniques to look at dynamics in specific cell–cell interactions. It is the latter approach that this chapter focusses on. The use of Förster resonance energy transfer (FRET) techniques to probe protein–protein interactions that are involved in receptor signalling to the cytoskeleton in intact cells is now well established. Various FRET biosensors are available to visualise several critical cell processes, giving information about activity and the location of key signalling molecules. As a specific set of examples in this chapter, we have generated variants of the original Rho, Rac and Cdc42 “Raichu” probes and improved their fluorophore combination to make them suitable for FLIM. These were employed in a number of assays to determine signal dynamics in T and NK cells. Specific protocols of how to use these probes and technical notes are described. Key words: Förster resonance energy transfer (FRET), fluorescence lifetime imaging microscopy (FLIM), Rho GTPases: Rho, Rac and Cdc42, signalling, protein interactions.

1. Introduction Genetically encoded fluorescent molecules allow tracking of the location of molecules of interest in cells, but they only give spatio-temporal information. This has allowed scientists to elucidate a lot about immune cell signalling where the constituents of a cascade translocate to different cellular compartments. However, if the distribution of a component stays the same, but is post-translationally modified or changes its affinity for a downstream molecule, then spatio-temporal information alone cannot F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_7, © Springer Science+Business Media, LLC 2010

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elucidate this. Various Förster resonance energy transfer (FRET) techniques have been successfully employed to detect these posttranslational modification events in intact cells. When coupled to multi-photon-based, high-resolution fluorescence lifetime imaging microscopy (FLIM), these techniques allow determination of populations of interacting protein species on a point-bypoint basis at each resolved voxel in the cell. FLIM-based FRET assays have been successfully applied to monitor signalling events in live cancer cells, in archival pathological tissues and, more recently, in fluorescently labelled cells in live animals, i.e. intravital fluorescence lifetime imaging in deep tissues (see Section 5). These assays provide important spatio-temporal information about post-translational modifications (e.g. protein phosphorylation (1–3) or ubiquitination (4)/sumoylation (5)), and interactions between signalling receptors (integrins, CD44, chemokine receptors and receptor tyrosine kinases (RTK)), protein kinases (PKC, Src kinases) and many cytoskeletal remodelling proteins including ezrin, fascin, RhoGTPases, WASP and the Arp2/3 complex (6–15). The term “biosensor” has been applied to a group of technologies that give a read-out of the activity of a molecule. A wealth of genetically encoded biosensors is now available which can be used to look at, for example, calcium concentration (16), small GTPase activity (17–21) or the activity of certain cell surface receptors (22). An early example of this approach is a cyclic-AMP monitor which consists of cAMP-dependent protein kinase where the catalytic and regulatory domains are labelled, respectively, with a pair of fluorescent molecules between which Förster resonance energy transfer (FRET) can occur. When cAMP binds to the regulatory subunits, the catalytic subunits dissociate and the FRET is lost (23). Another early application of this idea is the “cameleon” calcium probe (16). The cameleon probe consists of cyan or blue fluorescent protein, calmodulin, the calmodulin-binding peptide M13 and yellow or green fluorescent protein (16). When cameleon encounters Ca++ the calmodulin wraps around M13 and brings the two fluorescent proteins into close enough proximity for FRET to occur. Thus, FRET between the two fluorophores becomes a measure of calcium concentration. Other FRET biosensors similar in concept have since been produced and exploited in a number of different laboratories, notably, the phosphorylation and guanine-nucleotide exchange monitor (PHOGEMON) probes by Prof. Matsuda (17–20). The PHOGEMON series of probes use a layout of yellow fluorescent protein (YFP) bound to a sensor region (e.g. Ras) linked by a flexible peptide linker to an effector region (e.g. Raf) which is bound to Cyan fluorescent protein (CFP) and finally a membrane anchor (e.g. farnesyl moiety). When GTP binds Ras, for example, the sensor region binds the effector region bringing the CFP

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and YFP into close proximity allowing FRET. A RhoA probe has also been developed where the sensor region sits outside of the fluorescent proteins so the whole length molecule with its wildtype tail can be used (21). There are now FRET biosensors for ERK (24), EGF receptor (22, 25), N-WASP (26), myosin light chain kinase (27), PKC (28, 29), Akt (30), phosphatidylinositol3,4-bisphosphate [PtdIns (3, 4)P (2)] and phosphatidylinositol3,4,5-trisphosphate [PtdIns (3–5)P (3)] (31). Immune surveillance requires immune cells to accurately differentiate between healthy and diseased cells. When interacting with potential target cells or antigen-presenting cells (APC), natural killer (NK) and T cells must dynamically reorganise their cytoskeleton to bring receptors to the cell–cell interface and transport vesicles containing effector molecules (32). This complex network of intracellular signalling components and associated regulatory mechanisms would be best studied by direct visualisation in cells in real time by imaging techniques. Recently, within the immunological field, there has been a major emphasis on developing advanced optical cell imaging methods to study lymphocyte signalling. In addition to complementary “population biochemistry”-based approaches (33), the novel optical imaging methods can provide valuable additional insight into the mechanisms of, e.g. immune cell–cell contact, termed the immunological synapse (IS), and its dynamics (formation and maintenance). A great deal has been learnt about supramolecular chemistry at the IS by using fluorescently labelled receptors, ligands and key signalling molecules and monitoring them in either fixed samples or by time-lapse microscopy (34–38). However, most of the fluorescence studies to date have focussed on the distribution of proteins in and around the IS (39, 40). The dynamic changes of the signalling function of these proteins have not hitherto been adequately addressed. Rho family GTPases regulate actin cytoskeletal remodelling, cell polarisation and adhesion in a number of cell types including T lymphocytes (41). In their active GTP-bound form, Rho GTPases bind to downstream effector proteins. The Raichu-Rac and Cdc42 probes mentioned earlier (18–20) have been modified in our laboratory to be suitable for FRET determination by FLIM (Fig. 7.1) by replacing the original fluorescent proteins with monomeric red fluorescent protein 1 (mRFP1) and enhanced green fluorescent protein (eGFP). By imaging these biosensors in live and fixed T and NK cells, Cdc42 and Rac activity may be measured on a cell-by-cell basis over time in response to ligands and antibodies immobilised on a substrate, or in response to antigen-bearing target cells/APC. In the case of coverslip-bound ligands or antibodies one gains total selectivity over which receptors are activated on the immune cell without a background of co-receptors and adhesion molecules

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Fig. 7.1. Cdc42 “Raichu” biosensor. When Cdc42 is in its active GTP-bound state, it binds PAK1 bringing GFP into close proximity with mRFP1. FRET reduces the GFP lifetime (τ ), which is measured by FLIM. τ is a measure of the average time that a molecule remains in the excited state. Practically, it is taken as the time required for the fluorescence intensity to fall to 37% (1/e) of its initial value. The loss of energy from the excited state via emission is best described by first-order reaction kinetics, resulting in an exponential fluorescence decay function: I(t) = I0 exp (-t/τ ). I(t) = fluorescence intensity as a function of time, I0 = fluorescence intensity at time zero.

beyond our control. However, this model can sense activating or inhibitory signals per se but not in the full context required for a complete physiological response. A model consisting of Jurkat cells adhering onto a glass surface (stimulatory substrate) precoated with stimulatory antibodies has previously shown a direct correlation between the polarisation of the microtubule organizing centre (MTOC) towards the APC-mimicking (42) stimulatory substrate and a redistribution of the T-cell receptor to the bottom of the cell attached to the substrate below. Here we present protocols not only for cell-stimulatory substrate-based applications but also for the imaging of cell–cell conjugates, the combination of which allows us to look at immune cell signalling in detail.

2. Materials 2.1. Cells

1. Jurkat human T-cell line (ATCC) and β1-integrin-deficient mutants A1 (43) 2. Raji line of B-lymphoblast-like cells established from a Burkitt’s lymphoma (ATCC)

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3. YTS human NK cell line (A subclone of YT; (44)) 4. The 721.221 human EBV transformed B-cell lymphoma that does not express classical class I MHC molecules (HLAA, B, C; from laboratory of Prof. L. Lanier; (45)) 2.2. Cell Culture and Transfection by Electroporation

1. Complete RPMI medium. RPMI 1640 medium (SigmaAldrich) supplemented with 10% foetal bovine serum (Sera Laboratories International Ltd.), 1% L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin (Life technologies Ltd., UK). 2. HEPES Buffer Solution (Invitrogen Gibco). 3. Sterile electroporation cuvettes (4 mm).

2.3. Ligand-Dependent Activation Assay for Jurkat T Cells

1. Fibronectin (Sigma-Aldrich) – Aliquots in sterile PBS, store at –20◦ C. 2. Laminin (Sigma-Aldrich) – Aliquots in sterile PBS, store at –20◦ C. 3. Vitronectin (Sigma-Aldrich) – Aliquots in sterile PBS, store at –20◦ C. 4. Anti-CD3 mAb (UCHT1; Cancer Research UK Monoclonal antibody service). Store at 4◦ C. 5. Anti-β1-integrin mAb (12G10; a kind gift from Prof. M. Humphries, University of Manchester). Store at 4◦ C. 6. Goat anti-mouse IgG (Fc-specific) F(ab )2 fragments (Sigma-Aldrich). Stock solution aliquots in PBS, store at –20◦ C. Make 10 μg/ml solution in bicarbonate buffer for use. 7. Anti-CD43 mAb (BD Pharmingen). Aliquots in sterile PBS store at –20◦ C. 8. Paraformaldehyde (Sigma-Aldrich): Prepare 4% (w/v) solution in PBS. The solution may need to be carefully heated on a stirring hot-plate in a fume hood to dissolve and is best stored, aliquoted, at –20◦ C. 9. Mowiol mounting medium (ICN) containing 2.5% (w/v) 1,4-diacabicylco[2.2.2]octane (DABCO; Sigma-Aldrich).

2.4. Conjugate Formation

1. CellTracker Orange CMTMR (5-(and-6)-(((4-chloromethyl)benzoyl)amino)tetramethylrhodamine; Invitrogen) dissolves in DMSO (Sigma-Aldrich); store aliquots at –20◦ C. 2. Superantigen enterotoxin E (SEE) (Toxin Technology). Resuspend in sterile PBS and store at –80◦ C. Special licence is required for acquiring this toxin and it is necessary to

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liaise with your institution’s safety officer for its appropriate disposal. 3. Poly-L-lysine (Sigma-Aldrich). Store 1 mg/ml aliquots at –20◦ C. Dilute stock with sterile PBS to make 10 μg/ml to use on the chamber slides. Store the final solution at 4◦ C for no longer than 4 weeks.

3. Methods 3.1. Cell Culture

3.2. Electroporation

Wild-type Jurkat, YTS, 721.221, β1-deficient Jurkat (A1) and Raji B cells are cultured in selection-free complete RPMI medium. A1wtβ1, A1β1-NPKY, A1β1-NPIY, A1β1-793 clones are cultured in complete RPMI medium containing 1 mg/ml G418 (Life Technologies Ltd.). All the cell lines double every 24–36 h and are passaged at 3–8 × 105 cells/ml (see Note 1). 1. Split cells the day before electroporation at 1:2 dilution. 2. The same electroporation protocol can be used for wild-type Jurkat, YTS, A1 Jurkat and any A1-derived cell lines expressing different β1 cDNAs. 3. The Raichu-Rac and Raichu-Cdc42 DNA used for YTS and Jurkat electroporation is prepared using endotoxin-free kits (Qiagen) and stored in distilled water. For co-transfection experiments the two different plasmids are used at 1:2 ratio. 4. Splitting the cells 1:2 for two consecutive days can improve transfection efficiency. 5. Use 106 cells for each sample and wash with serum-free media containing 25 mM HEPES before mixing with the plasmid DNA. Microcentrifuge tubes and 4 mm cuvettes are labelled in advance for each sample. Plasmid DNA of each sample is transferred in the respective labelled tube. Cell suspension and DNA should be mixed thoroughly and transferred carefully in the cuvette to avoid bubbles. The electroporation medium does not need to be pre-warmed at 37◦ C. 6. Electroporate in 250 μl serum-free RPMI medium containing 25 mM HEPES using 40 μg of plasmid DNA at 260 V/960 μF using the Gene pulser II electroporation system (Biorad). 7. Leave cuvettes on ice for 10 min before electroporation. 8. After electroporation resuspend samples in pre-warmed complete RPMI media and wash twice before transferring cells to a flask. During your washes, remove all debris thoroughly to ensure good post-electroporation survival.

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9. Allow cell lines to express the construct for 24 h before imaging. 3.3. Integrin Ligand and Antibody Stimulation Assays

1. Use fibronectin, vitronectin and laminin at 10 μg/ml. Coat sterilised coverslips with each of the ligand for 1 h at room temperature before allowing cells to adhere on them (see Note 2). 2. For the antibody assays, coat coverslips with a goat antimouse (Fc-specific) F(ab )2 fragment (10 μg/ml in bicarbonate buffer) at 4◦ C overnight (see Note 2). 3. Wash excess F(ab )2 with PBS and apply antibody (e.g. antiCD3 or β1-integrin mAb) solution (10 μg/ml) on the glass for 2 h at 37◦ C. 4. Wash antibody-coated coverslips with warm serum-free media to remove excess antibody before allowing the cells to attach for the required period of time. 5. For serum starvation experiments, electroporated cells are washed with serum-free media and incubated on antibodies in serum-free conditions for the required time. 6. See Note 3.

3.4. Ligand-Dependent Activation Assay for Live Imaging

1. For live imaging, coat chambered cover glasses (LabTek; Nunc) with goat anti-mouse (Fc-specific)F(ab )2 fragment (10 μg/ml) (Sigma) at 4◦ C overnight. 2. Incubate the chamber slides with the stimulatory monoclonal antibodies, at a final concentration of 10 μg/ml in PBS, for 2 h at 37◦ C. 3. Remove excess antibody by washing with serum-free media. 4. Wash cells and resuspend them in pre-warmed RPMI without phenol red, containing 10% FCS and 25 mM HEPES. 5. Plate at a density of 1 × 106 cells/ml during imaging.

3.4.1. Jurkat–Raji Conjugate Formation

1. Pre-incubate Raji APCs for 20 min at 37◦ C with CMTMR (10 μM) fluorescent marker. 2. After three washes in warm media, incubate cells for 45 min at 37◦ C with 1 μg/ml SEE. 3. Mix Jurkat (2 × 105 ) with equal number of Raji and incubate at 37◦ C for the required time period. Centrifuge the combined cell suspension at 1,000 rpm for 5 min. 4. Resuspend gently and treat with Cytofix/Cytoperm (BD Pharmingen) for 5 min at 4◦ C to fix and permeabilise the conjugates. 5. Upon fixation, allow them to settle on poly-L-lysine (10 μg/ml)-coated chamber slides (LabTek Chamber slides; Nunc) at 4◦ C overnight before mounting. Do not incubate

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live cells on poly-L-lysine to avoid cytoskeleton rearrangements and adhesion on the substratum. 3.4.2. YTS:721.221 Conjugate Formation

1. Pre-incubate 721.221 for 20 min at 37◦ C with CMTMR (10 μM) fluorescent marker. 2. Wash 3× with RPMI 1640. 3. Mix YTS (1 × 106 ) with an equal number of 721.221 cells and incubate at 37◦ C for the required time periods in <500 μl of medium so that centrifugation is not required to pellet the cells for fixation. Resuspend gently in 4% paraformaldehyde or Cytofix/Cytoperm (if cells are going to be stained later; BD Pharmingen) for 15 min at 4◦ C to fix and permeabilise the conjugates. 4. Wash in PBS-TWEEN-20 (0.1% v/v) or PBS depending on whether the cells are going to be stained with antibodies/phalloidin or not, respectively. 5. To stain F-actin with phalloidin-Alexa647 (Invitrogen), resuspend the cells in 16.5 μM phalloidin in PBS 1% BSA (w/v) for 1 h at 4◦ C and wash 2× with PBS– TWEEN-20 and once with PBS before mounting as in Section 3.5.1.

3.5. Preparation of Samples for FLIM and Confocal Microscopy 3.5.1. Preparation of Samples That Do Not Require Antibody Staining

3.5.2. Preparation of Samples for Antibody Staining

1. Cells or conjugates that are not stained with antibodies are fixed in 4% paraformaldehyde for 15 min at room temperature. 2. Treat with NaBH4 (1 mg/ml) for 5 min to quench free aldehyde groups. 3. After rinsing twice in PBS, and once with deionised water, the coverslips can be mounted using mounting medium (ICN, CA, USA) containing 2.5% (w/v) 1,4diacabicylco[2.2.2]octane (DABCO; Sigma-Aldrich, UK). 1. For antibody staining, block samples with 10% horse serum v/v (Sigma-Aldrich) 3% (w/v) BSA in PBS containing 0.1% (v/v) TWEEN-20 (replace PBS for TBS when staining for phosphorylation groups) for 1 h at 4◦ C. 2. Incubate with primary antibody (1:50 to 1:500 depending on Ab used in blocking buffer used in step 1.) for 2 h at 4◦ C. 3. Wash with appropriate buffer and incubate with secondary antibody (1:500 to 1:1,000 in blocking buffer used in step 1) for 1 h at 4◦ C. 4. Mount coverslips as described in Section 3.5.1.

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1. Fluorescence lifetime imaging in our laboratory is achieved on a custom-built fluorescence lifetime imaging microscope with bespoke acquisition and analysis software. However, general details and advice on image acquisition and analysis are given below. 2. Figure 7.2 is a simplified schematic of our system. 3. Epifluorescence images of all fluorophores are collected before FLIM. GFP:FITC filter-set, mRFP and CMTMR:Cy3 filter-set and Alexa 647:Cy5 filter-set (all Chroma). 4. For imaging GFP in YTS and Jurkat cells a 40× oilimmersion objective with a numerical aperture of 1.4 was used. 5. GFP was excited by two-photon excitation with a titanium:sapphire pulsed laser at 900 nm (two-photon wavelength 450 nm).

Fig. 7.2. A schematic of a typical confocal or multi-photon excitation TCSPC FLIM set-up. A pulsed laser (black arrows) excites the sample at a MHz repetition rate, and the fluorescence (grey arrows) is detected with a descanned (for a confocal) or non-descanned (for multi-photon excitation) detector. The electronic signals for the detected photons and for the beam scan synchronisation are fed into a TCSPC card in a PC. The time delay between the laser excitation pulse and the arrival of the fluorescence photon is measured many times, and a histogram of photon arrival times is built up in each pixel, which represents the fluorescence decay in each pixel. Subsequent data analysis allows the extraction of the fluorescence lifetime τ from the decays and its visualisation in a FLIM map. τ is visualised on a greyscale.

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6. The fluorescence lifetime is determined by time-correlated single photon counting (TCSPC), using a Becker & Hickl card (see Section 4 and Note 4 for details). 7. Images must be collected with care so that enough photon data are acquired to allow an accurate fit of fluorescence lifetime. However, if the excitation power is too high for a given sample brightness photons can arrive at the detector too fast to measure (termed pile-up). Therefore to get the best resolution a compromise must be found between acquisition time and excitation power.

4. Discussion on How to Improve the Accuracy and Specificity of the Assays

The Raichu biosensors for Cdc42 and Rac1 are known to exhibit a baseline level of sensitised emission due to FRET, as a result of the finite separation between the two fluorescent proteins within the same construct even when the biosensor is in the open conformation (18). A number of negative controls are therefore required to establish the baseline for non-specific activities of the biosensors. As an example, for un-stimulated controls, we measured the GFP fluorescence lifetime of Raichu-Rac-expressing Jurkat T cells that had been fixed with paraformaldehyde while in suspension, then cytospun onto a slide before mounting (Fig. 7.3). Additional controls include (i) cells that express Raichu probes which contain the T17N mutation which is known to have a much reduced affinity for GTP (Fig. 7.3) and (ii) cells that express wild-type Raichu probes but are adhered onto glass surface pre-coated with irrelevant antibodies (e.g. anti-CD43 mAb). In the example shown in Fig. 7.3, the presence/absence of FRET was determined by fitting the experimental data to a single exponential decay. Sufficient reduction in the measured lifetime indicates FRET. As a more accurate assessment, one can apply a bi-exponential fluorescence decay model to the data to determine the fluorescence lifetimes of non-interacting and interacting subpopulations, respectively. This can be further coupled to various global analysis techniques that simultaneously fit the fractional contributions in all pixels and the spatially invariant lifetimes. The latter are outside the scope of this chapter but have been described elsewhere (46–48). In another example shown in Fig. 7.4, YTS NK cells are surveying potential target cells in the presence or absence of an integrin blocking antibody. When the integrin is blocked, F-actin does not cluster at the IS and there is a corresponding drop in Cdc42 activity. In this example, a bi-exponential fluorescence

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Fig. 7.3. “Raichu” Rac biosensor imaging in Jurkat T cells on antibody-coated cover slips. Jurkat T cells were transfected with Raichu-Rac or Raichu-RacN17. Cells were either fixed in solution (cytospin) or allowed to adhere on anti-β1-integrin antibody (12G10) for 30 min at 37◦ C prior to fixation. Multi-photon FLIM was undertaken to determine the extent of FRET between the donor GFP and the acceptor mRFP1 upon binding of PAK CRIB to the Rac1 region of the probe. FRET results in shortening of the GFP lifetime (τ ); mean FRET efficiency was calculated based on the normalised pixel count of each cell. (a) Left, greyscale images display multi-photon GFP intensity. Right, GFP lifetime (τ ) is shown in greyscale using the standard inverse look-up table. (b) Mean FRET efficiency graph presents cumulative data of three independent experiments (n > 10 cells). ∗∗ p-value < 0.01 in comparison with cytospin control.

decay model has been used to fit the Raichu-Cdc42 FLIM data. This allows us to calculate the average fraction of pixels where the probe is undergoing FRET (i.e. those which fit a shorter lifetime). Cumulative data from several experiments are displayed in Fig. 7.4b.

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Fig. 7.4. YTS NK cell surveying a potential target cell. (a) Left panels, YTS NK cells transfected with eGFP/mRFP RaichuCdc42 were mixed with 721.221 target cells labelled with CMTMR, and after 15 min of co-incubation fixed and stained with phalloidin (Right panels). Actin is known to accumulate at the activating NK IS. When CD11a was blocked with an antibody (Clone 38, CRUK Ab collection) phalloidin clustering was abrogated at 15 min (b) The mean fractional intensity of the shorter lifetime in a bi-exponential fit (F2) was reduced to baseline levels (data averaged from five cells).

5. Outlook and Future Development

Advances in light microscopy, particularly in two-photon excitation, are starting to allow us to describe events in immune cell signalling not only in terms of in vitro cell–cell interactions but also in the physiologically more relevant in vivo situation, i.e. in living organs or animals. The first studies to take advantage of this have focussed on tracking immune cells in the periphery and within lymph nodes (49, for a review see 50). One elegant example uses multicolour intravital two-photon microscopy, where several different cell types may be tracked simultaneously, revealing that T cells travel down a network of follicular reticular cells to navigate within the lymph node (51). These studies reveal a lot about the geography of the immune cell interactions. However, switching from spatio-temporal cell tracking to activity tracking of key signalling components during physiological interactions will lead to newly derived image parameters that report on the functionality of these immune cell–cell interactions and can quantitatively predict biological decisions that are made subsequent to the contact process and are critical to immune cell/target cell fate. To acquire these kinds of data, a number of challenges must be overcome by combined interdisciplinary approaches, some aspects of which are briefly described in the following paragraphs. Nonlinear multi-photon microscopy has several advantages over single-photon excitation, which allow imaging at

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significantly higher depths within a tissue (52). First the penetration depth of near infra-red (NIR; as used in two-photon excitation) is deeper in scattering tissues, and second NIR is generally less phototoxic due to a lack of significant endogenous (singlephoton) absorbers (53). Furthermore, confocal set-ups are disadvantageous for deep-tissue imaging since there is a lot of light loss due to the confocal pinhole used. Consequently, wide-field set-ups are preferable and accurate spatio-temporal fluorescence information is accessible via synchronisation with the excitation light scanning unit. Another technological improvement required for deep-tissue imaging is efficient wave-front correction. Several methods are currently explored in the field of adaptive optics and “deformable mirrors” are a promising route to efficient wavefront correction. The principle has already been shown to work in optical microscopy (54); however, the technology needs to be improved in order to reduce (online)-reaction times to aberrations induced by variations in tissues (55) in live specimens. In addition to the advance of the microscope, there is also improvement in the field of genetically expressible fluorophores. Single-photon and two-photon excitabilities of fluorophores differ. Luckily, fluorescent proteins have rather large two-photon cross-sections and consequently are easy to excite. Normally, for deep-tissue imaging such fluorescent proteins are desirable that can be excited simultaneously, but emit at different wavelengths that can be spectrally easily separated. A very recent example is the combination of eGFP with the red fluorescent protein mKeima (56). This is important since it omits the sometimes tedious process of laser tuning during an experiment. However, for FRET applications these combinations are obviously not suitable. In our laboratory we are currently exploring the suitability of a variety of RFPs in order to find the best FRET pair for in vivo two-photon microscopy. Although there are considerable technological improvements facilitating deep-tissue imaging and leading the way to in vivo FRET by FLIM, there will always be a limitation of specimens that are easily accessible in a living organism. To a certain degree surgical procedures extend the applicability of optical microscopy techniques to “surgically accessible” targets. We are currently exploiting this opportunity for breast cancer xenograft models and have achieved intravital FLIM/FRET imaging by exposing the tumour and draining lymph nodes by cutting a skin flap, in order to facilitate their optical accessibility. Implantation of window chambers is a less physiological in vivo model; however, they are vastly advantageous over all other currently available methods if repeated imaging over long periods of time is required. Taken together, interdisciplinary progress in advanced optical microscopy techniques currently overcomes several challenges of in vivo deep-tissue imaging techniques and, more importantly, in

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the near future these techniques are likely to expand their applicability to in vivo nano-scale measurements by FRET, and thus probing direct molecular interactions and their dynamics rather than “just” spatio-temporal distribution.

6. Notes 1. After 12 passages alterations in cell morphology and size of wild-type Jurkat were noticed, so to ensure reliable and reproducible results, none of the T-cell lines described should be maintained for more than ten passages. 2. Before applying any ligand/antibody solution onto glass, put coverslips on hydrophobic film to ensure the ligand/ antibody solution stays on the glass and is applied evenly over the coverslip surface. 3. For effective comparison between conditions and to reduce variation between different transfection experiments, cumulative FRET data should be obtained from independent sets of experiments done in parallel. Take special care to generate and compare results between independent sets of experiments only. 4. Please refer to (57, 58) and web sites for Becker & Hickl (http://www.becker-hickl.de) or PicoQuant (http:// www.picoquant.com) for advances in technology for timecorrelated single photon counting (TCSPC), a subject which is beyond the scope of this chapter.

Acknowledgments Konstantina Makrogianneli was supported by a joint research studentship from MRC/King’s College London. Leo Carlin is supported by a UK EPSRC grant (EP/C546105/1). G. Fruhwirth is supported by the King’s College London and University College London Comprehensive Cancer Imaging Center CR-UK & EPSRC in association with MRC and DOH (England) C1519/A10331 M. Keppler and T. Ng are supported by an endowment fund from the Dimbleby Cancer Care to King’s College London. The multi-photon FLIM system was built with support from both the Medical Research Council Co-Operative Group grant (G0100152 ID 56891) and a UK Research Councils Basic Technology Research Programme grant (GR/R87901/01).

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Chapter 8 Methods for Quantitation of Leukocyte Chemotaxis and Fugetaxis Fabrizio Vianello, Elda Righi, and Mark C. Poznansky Abstract Chemoattraction and chemorepulsion are complex directional responses of a cell to external chemotactic stimuli. The decision of a cell to move towards or away from a chemokinetic source includes detection and quantitation of the gradient of the chemotactic agent, biochemical transmission of the stimulus, and translation into a directional migration. This chapter describes a number of in vitro and in vivo assays that can be used to generate and measure both chemoattraction and chemorepulsion of leucocytes. These tools may eventually allow the further characterisation of the mechanism of this complex and physiologically and pathologically important phenomenon. Key words: Chemotaxis, fugetaxis, in vivo imaging, lymphocyte trafficking.

1. Introduction Active movement of leucocytes towards a site of antigen challenge, infection, or tissue injury is a central component of the establishment of both inflammatory and immune responses (1–4). The term chemotaxis describes all directional migration of leucocytes up a gradient and towards a peak of concentration of a chemoattractant or a chemotactic factor, which includes chemokines, a superfamily of 8- to 10-kDa proteins (4, 5). Up until recently, it was thought that eukaryotic cells could undergo active movement only towards an agent, although cellular structures including eukaryotic neuronal growth cones were shown to move both towards and away from a chemokinetic agent such as semaphorin and netrin-1 (6–9). Convincing evidence has F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_8, © Springer Science+Business Media, LLC 2010

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subsequently been provided that immune cells including lymphocytes are capable of bidirectional movement. T cells, mature single-positive (SP) thymocytes, monocytes, neutrophils, and dendritic cells can undergo active movement away from a chemokinetic agent and that this mechanism may be biologically relevant to T lymphopoiesis and leucocyte trafficking, the generation of immune-privileged sites, immune evasion by viruses and cancer, and a diverse range of other physiological and pathological processes (10–16). The term “fugetaxis” has been introduced into the scientific literature and is used interchangeably with the term chemorepulsion and refers to an agent that can repel cells (15, 17). Previous studies have shown that T cells can migrate away from a high concentration of the chemokine SDF-1 or CXCL12 when the same chemokine is a known chemoattractant for T cells at lower concentrations. CXCL12-induced chemorepulsion is Gαi protein mediated, suggesting that the concentration of ligand and receptor occupancy can influence the directional decision (17). The local and temporal coexistence of chemotaxis and fugetaxis may play a role in physiological and pathological aspects of lymphocyte migration. For example, CXCL12 was shown to be highly expressed in the thymus and in the bone marrow (18). The thymus represents an interesting model in which to study chemoattraction and chemorepulsion, particularly in the involvement of these directional decisions in the process of thymocyte emigration or egress from the medulla and into the vasculature. The finding that thymocyte egress requires the presence of an extrathymic chemoattractive gradient of sphingosine1-phosphate (S1P) from the thymus to the plasma has been augmented by the finding that this mechanism may act in concert with intrathymic chemorepellent gradients of CXCL12 (14, 19). Chemoattraction and chemorepulsion are relevant in many fields of human pathology as cancer and transplantation. An important consideration in the development of T-cell-based cancer immunotherapy is that effector T cells must efficiently traffic to the tumour microenvironment in order to control malignant progression. There is evidence that tumours may be able to evade an efficacious immune response by secreting chemorepellent levels of chemokines such as CXCL12 (16). On the other hand, the same mechanism can be positively exploited in the transplantation setting by inducing high levels of CXCL12 to avoid T-cell infiltration and thereby, graft rejection (20). All these evidences of a role of chemotaxis and fugetaxis in lymphocyte trafficking have been consistently demonstrated by accurate and reproducible techniques.

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The aim of this chapter is to discuss and give details on the range of methods that are currently available for identifying and quantitating chemoattraction and chemorepulsion, or fugetaxis of leucocytes.

2. Materials 2.1. Transmigration Assay

1. C57Bl6 mice (4–6-week old). 2. Chicken ovalbumin (Sigma) containing less than 0.01% lipopolysaccharide by the limulus assay and dissolved in 100 μl complete Freund’s adjuvant (Sigma). 3. Chicken ovalbumin 100 mg dissolved in 250 ml doubledistilled H2 O. 4. SDF-1 (PeproTech). 5. Anti-CD3-conjugated antibody. 6. PBS. 7. Iscove’s modified Dulbecco’s medium supplemented with 0.5% foetal calf serum, 50 units/ml penicillin, 50 μg/ml streptomycin, and 292 μg/ml L-glutamine (Mediatech, Herndon, VA). 8. Transwell system 6.5 mm diameter and 5 mm pore size with polycarbonate membrane (Corning). 9. Microscope or FACS.

2.2. Assays for Detection of the Migrational Response of T Cells to Cell-Secreted CXCL12

1. DMEM supplemented with 10% foetal bovine serum, penicillin, and streptomycin. Cells producing different levels of CXCL12. 2. Fibronectin- or laminin-coated 24-well or 48-well plates (BD Biosciences). 3. Plastic cylinder for seeding cells into the plate (Costar). 4. Microscope and digital camera.

2.3. Leucocyte Chemotaxis in Linear and Complex Gradients of Chemokines Formed in a Microfabricated Device

1. Human IL-8 (PeproTech, Rocky Hill, NJ). 2. Neutrophils are isolated from human whole blood by density-gradient centrifugation and purified by hypotonic lysis. 3. Nikon Eclipse TE2000-S microscope (Nikon, Japan). 4. IPLab 3.6.1 software (Scanalytics, Fairfax, VA). 5. MetaMorph 4.5 software (Universal Imaging, Downington, PA). 6. MATLAB 13 software (The Mathworks, Inc., Newton, MA).

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2.4. Abrogation of Intraperitoneal Inflammatory Infiltration of Mice

1. C57Bl6 mice (4–6-week old). 2. Chicken ovalbumin (Sigma) containing less than 0.01% lipopolysaccharide by the limulus assay and dissolved in 100 μl complete Freund’s adjuvant (Sigma). 3. Chicken ovalbumin 100 mg dissolved in 250 ml doubledistilled H2 O. 4. SDF-1 (PeproTech). 5. Anti-CD3-conjugated antibody. 6. PBS. Details on materials for MRI tracking of adoptively transferred antigen-specific T cells and for intravital microscopy of rat mesentery have been included in Sections 3.5 and 3.6.

3. Methods 3.1. Transmigration Assay

A variety of transmigration assays exist which exploit the ability of migratory cells to polarize and migrate in response to a gradient of a chemotactic agent. The in vivo correlate of in vitro transmigration is thought to be the migration of cells towards sites of tissue injury, pathogen invasion, and/or immune challenge. Cells to be used in the transmigration assays can be derived from a number of sources, including the diseased tissue itself or purified subpopulations of inflammatory and immune cells isolated from the peripheral blood. Chemokine-dependent chemotaxis is assayed on various leucocytes by an in vitro two-chamber migration assay. Quantitation can be performed by cell count or by flow cytometry on the cells collected from the chambers. The following protocol has been established in our laboratory to assess chemotaxis and fugetaxis in the presence of different concentrations of CXCL12. 1. Fivex 104 cells resuspended in 100 μl of medium are added to the upper chamber of Costar Transwells (6.5 mm diameter, 5 μm pore size, polycarbonate membrane). 2. SDF-1 at concentrations of 10 μg/ml, 1 μg/ml, 100 ng/ml, 10 ng/ml, and 0 ng/ml is added in the lower, the upper, or both lower and upper chambers of the Transwell to generate a standard “checkerboard” analysis matrix of positive, negative, and absent gradients of SDF-1, respectively. 3. Cells are incubated for 3 h at 37◦ C in an atmosphere of 5% CO2 . 4. After incubation, cells are collected from the lower chamber and counted.

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This method allows the visualization of T-cell responses in noncontrolled CXCL12 gradients generated by cells over time. It is suitable for assessing the migratory response of leucocytes to a gradient of chemokine generated by a definite cell population (20). 1. Control cells and cells producing the chemokinetic/fugetactic molecules are seeded in a 24- or 48-well plate using a plastic cylinder and grown as round 6-mm patches. 2. Cells are incubated at 37◦ C in 5% CO2 overnight in medium. 3. Murine T cells are then added to the plate spreading them all around the cylinder. 4. The plate is incubated for 1 h. 5. The cylinder is carefully removed while checking for adherence of cells. 6. Cells are incubated for 15 h at 37◦ C. 7. Images are then taken every 30 s for 60 min using a digital camera controlled by IPLab software. 8. Migratory paths of randomly selected T cells towards and/or away from cells producing the chemokine can be tracked using MetaMorph software. 9. The mean chemotropic indices (MCIs) can be determined as a quantitative measurement of directional movement using MATLAB software.

3.3. Leucocyte Chemotaxis in Linear and Complex Gradients of Chemokines Formed in a Microfabricated Device

Microfabrication has great promise for creating and controlling microenvironments in which cell behaviour can be observed in real time. Soft lithography with polydimethylsiloxane (PDMS) is customarily used as a stamping technique to prepare surfaces that are in a quasi-two-dimensional array. The chemotaxis devices can be used to generate stable gradients, manipulate the microenvironment of cells, and study the basic mechanisms of cell migration in real time. This technology has been used to generate stable, soluble chemoattractant gradients. These gradients are produced by controlled diffusive mixing of species in solution that flow inside a network of microchannels under conditions of low Reynolds number. The described method has been successfully applied to neutrophil chemotaxis by generating soluble linear gradients of IL-8. (13, 21) 1. Neutrophils at a concentration of 5 × 106 cells/ml are loaded uniformly across the migration channel (dimensions 4,000 μm × 450 μm × 130 μm) and allowed to adhere within a microfluidic device for up to 30 min (on average, 1,200 neutrophils adhere within the migration channel of the device).

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2. Adherent neutrophils within the device are then exposed to the absence or the presence of a linear gradient of human IL-8. 3. Migration is observed in a Nikon Eclipse TE2000-S microscope. Bright field images (10×) are taken every 30 s for 40 min using a digital camera. 4. Cell movement (50 cells) can be tracked using MetaMorph 4.5 object-tracking application, which generated tables of Cartesian coordinate data for each migratory step during the migration of a cell. 5. Tracking data are analysed in Excel and MATLAB 13 to determine angular frequencies, mean-squared displacements, migration velocities, persistence times, random motility coefficients, and random walk path lengths. 3.4. Abrogation of Intraperitoneal Inflammatory Infiltration of Mice

Several systems have been developed to assess leucocyte infiltration into specific anatomical sites in vivo. Chemokines can be injected intradermally, subcutaneously, or into the intraperitoneal space and leucocyte infiltration thus quantitated. Recruitment of endogenous leucocytes or adoptively transferred leucocytes can be assessed in this manner. In our experience, the assessment of an infiltrate in response to direct injection with a chemokine in the peritoneal space is consistently reliable and reproducible (13, 22). 1. Prime C57 BL/6 J mice (Jackson Laboratories, Bar Harbor, Maine) subcutaneously with 100 μg chicken ovalbumin dissolved in 100 μl complete Freund’s adjuvant. 2. After 3 days, ovalbumin-primed mice are challenged by intraperitoneal injection of 100 μg ovalbumin dissolved in 250 μl double-distilled H2 O (control group). 3. The following day, a second intraperitoneal injection of 250 μl mouse SDF-1α is given at a concentration of 100 ng/ml (experimental group 1), 1 μg/ml (experimental group 2), or PBS (experimental group 3). 4. Mice are sacrificed after 24 h. 5. Peritoneal lavage with 5 ml PBS is performed by directly visualizing the peritoneal sac and its contents to avoid peripheral blood contamination. Peritoneal fluid samples obtained in this way contain less than 0.1% red blood cells. 6. Pellets from IP samples are prepared. 7. Samples are stained with anti-CD3 and other T-cell antibodies for flow cytometry quantitation. The proportion of each T-cell subpopulation is determined as a percent of the total nucleated cell fraction in the peritoneal fluid.

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Visualization of adoptively transferred T cells in vivo depends on efficient labelling methods and on the detection system. MRI is non-invasive and provides high spatial resolution images in vivo. By using nanoparticles as oxide particles to efficiently label lymphocytes, it is possible to detect adoptively transferred T cells in vivo via MRI over time. The following technique is based on labelling of T cells with a magnetic nanoparticle. We have experience with tatderivatized CLIO (cross-linked superparamagnetic iron oxide) which consists of dextran-coated superparamagnetic iron oxide nanoparticles which are vehicolated into cells by the HIV-1 tat peptide (16, 23). This technique is particularly suitable for studying tumour masses producing chemokines that may affect T-cell recruitment by chemoattraction or chemorepulsion (16, 23). 1. Tumour-specific CD8+ T cells are incubated with CLIOHD (2–300 μg Fe/ml/10 × 106 cells) for 4 h at 37◦ C and washed three times by centrifugation through 40% Histopaque-1077. 2. Mice are injected subcutaneously with tumour cells expressing the chemokine (in one flank) and with control tumour cells (in the opposite flank). 3. When subcutaneous tumours become palpable, 3 × 107 CLIO-HD-labelled tumour-specific CD8+ T cells are adoptively transferred via i.p. injection. Alternatively, T cells can be transferred by i.v. injection. 4. Distribution of CLIO-HD-labelled cells over time is then assessed via repetitive MRI after 12, 24, and 48 h from adoptive transfer. 5. Tumours are subsequently excised and used for histological analysis. 6. MRI quantitation of T-cell recruitment is based on different parameters, which allow to correlate T2 map to number of T cells per voxel (23).

3.6. Intravital Microscopy of Rat Mesentery

An established technique for investigating the in vivo events within a microvascular bed is by the use of intravital microscopy. This procedure allows the direct visualization of the microcirculation within translucent tissues such as the mesentery in anaesthetized animals. Migration of leucocytes from the microvascular lumen to the extravascular tissue and the effect of chemokine gradient in the bidirectional movement can be visualized using this approach (13, 22). 1. Mice are sedated (i.m. Hypnorm). 2. Anaesthesia is then performed with i.v. sodium pentobarbitone (30 mg/kg loading dose followed by 30 mg/kg/h). Animals are kept at 37◦ C on a heated microscope stage.

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3. Following midline abdominal incision, the mesentery is arranged adjoining the terminal ileum over a glass window on the microscope stage and pinned in position. 4. The mesentery should be kept warm and moist by continuous application of Tyrode’s balanced salt solution. 5. To assess neutrophil migration, the mesentery can be stained with acridine orange (Sigma-Aldrich), a nuclear dye, and scanned with a 488-nm laser line generated from an Argon laser. Visualization can be performed by confocal microscopy. 6. Neutrophil recruitment and migration can be recorded by time-lapse video microscopy and migrating neutrophils positively identified and counted following acridine orange staining of the mesentery. Extravasated leucocytes are defined as those in the perivenular tissue adjacent to, but remaining within a distance of 50 μm of, a 100-μm vessel segment under study.

4. Notes 1. About 50–60% of T cells are expected to migrate towards a chemotactic concentration of SDF-1 in a typical transmigration assay (11). 2. In the assay for detection of the T-cell migration towards cells secreting CXCL12, the total number of cells seeded in the plastic cylinder depends on the amount of chemokine secreted. In our experience, 1–2 × 105 cells producing the chemokine were seeded and 1 × 106 T cells/well were then added using a 48-well plate. Incubation time of chemokineproducing cells should be tested as it may vary depending on the adherence of cells. 3. The majority of neutrophils adhere within the device after their initial introduction and remain adherent within the device throughout the 40-min video-recording period. When neutrophils are exposed to a steeper gradient (0–2.4-μM) of IL-8, a significant reduction in cell adherence can be expected (about 10% adherent cells). 4. It is important that the mice from different experimental groups be of appropriate genetic background, age, and have similar weights. Differences in genetic background, age, and weight can result in differences in cell yield. T-cell recovery from the peritoneal cavity of ovalbumin-challenged mice typically ranges from 0.5 to 1.5 × 105 per millilitre of peritoneal fluid. If it is obvious that bleeding occurred, be

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mindful about errors in cell counts and red blood cells in differentials, or if necessary, discard the sample. 5. T-cell infiltration of tumours as detected by MRI may show some variability. We found that T-cell recruitment reaches the highest level after 24–48 h. As CLIO-HD is released in the tissue following T-cell death, a signal can be detected even after 48 h from adoptive transfer. 6. Expected viability of CD8+ T cells is >95% when CLIO-HD concentration is 50–300 μg Fe/ml/106 cells.

5. Conclusions Eukariotic cell chemoattraction and chemorepulsion play fundamental roles in physiological and pathological processes. Migration is a balance among chemoattraction, chemokinesis (random movement), and chemorepulsion and this regulates the movement of leucocytes into and from anatomic sites, including the bone marrow, the thymus, and lymph nodes. Over the past 5 years there has been a considerable expansion in the number of methodological approaches available to scientists for quantitation of cell migration. These techniques vary widely from phase-contrast fluorescent digital video microscopy to the examination of individual cell movement in response to chemokines to the use of knockout mice to delineate the effects of specific chemokines and chemokine receptors in vivo. These diverse methodological approaches have revealed an intricate world of factors influencing and inflammatory cell localization, which appear critical to the pathophysiology of a wide variety of diseases ranging from allergic lung inflammation, through atherosclerosis to the way in which the immune system handles infectious agents such as Toxoplasma and Leishmania. During the next 10 years, we should expect to see a further expansion in the methodological approaches to studying the roles of chemokines and chemokine receptors, which should itself lead to a greater understanding of human diseases and the development of novel therapeutic approaches in order to combat them. References 1. Baggiolini, M. (1998) Chemokines and leukocyte traffic. Nature 392: 565–568. 2. Luster AD (1998) Chemokines–chemotactic cytokines that mediate inflammation. N Engl J Med 338: 436–445. 3. O’Neil D, Steidler L. (2003) Cytokines, chemokines and growth factors in the pathogenesis and treatment of inflamma-

tory bowel disease. Adv Exp Med Biol 520: 252–285. 4. Zlotnik A, Yoshie O (2000) Chemokines: a new classification system and their role in immunity. Immunity 12: 121–127. 5. Rossi D, Zlotnik A (2000) The biology of chemokines and their receptors. Annu Rev Immunol 18: 21–242.

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6. Goshima Y, Sasaki Y, Nakayama T, Ito T, Kimura T (2000) Functions of semaphorins in axon guidance and neuronal regeneration. Jpn J Pharmacol 82: 273–279. 7. Kikutani H, Suzuki K, Kumanogoh A (2007) Immune semaphorins: increasing members and their diverse roles. Adv Immunol 93: 121–143. 8. Ly NP, Komatsuzaki K, Fraser IP, Tseng AA, Prodhan P, et al. (2005) Netrin-1 inhibits leukocyte migration in vitro and in vivo. Proc Natl Acad Sci U S A 102: 14729–14734. 9. Suzuki K, Kumanogoh A, Kikutani H (2008) Semaphorins and their receptors in immune cell interactions. Nat Immunol 9: 17–23. 10. Brainard DM, Tharp WG, Granado E, Miller N, Trocha AK, et al. (2004) Migration of antigen-specific T cells away from CXCR4binding human immunodeficiency virus type 1 gp120. J Virol 78: 5184–5193. 11. Poznansky MC, Olszak IT, Foxall R, Evans RH, Luster AD, et al. (2000) Active movement of T cells away from a chemokine. Nat Med 6: 543–548. 12. Stevceva L, Yoon V, Anastasiades D, Poznansky MC (2007) Immune responses to HIV Gp120 that facilitate viral escape. Curr HIV Res 5: 47–54. 13. Tharp WG, Yadav R, Irimia D, Upadhyaya A, Samadani A, et al. (2006) Neutrophil chemorepulsion in defined interleukin-8 gradients in vitro and in vivo. J Leukoc Biol 79: 539–554. 14. Vianello F, Kraft P, Mok YT, Hart WK, White N, et al. (2005) A CXCR4-dependent chemorepellent signal contributes to the emigration of mature single-positive CD4 cells from the fetal thymus. J Immunol 175: 5115–5125. 15. Vianello F, Olszak IT, Poznansky MC (2005) Fugetaxis: active movement of leukocytes

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away from a chemokinetic agent. J Mol Med 83: 752–763. Vianello F, Papeta N, Chen T, Kraft P, White N, et al. (2006) Murine B16 melanomas expressing high levels of the chemokine stromal-derived factor-1/CXCL12 induce tumor-specific T cell chemorepulsion and escape from immune control. J Immunol 176: 2902–2914. Huttenlocher A, Poznansky MC (2008) Reverse leukocyte migration can be attractive or repulsive. Trends Cell Biol 18: 298–306. Shirozu M, Nakano T, Inazawa J, Tashiro K, Tada H, et al. (1995) Structure and chromosomal localization of the human stromal cellderived factor 1 (SDF1) gene. Genomics 28: 495–500. Schwab SR, Cyster JG (2007) Finding a way out: lymphocyte egress from lymphoid organs. Nat Immunol 8: 1295–1301. Papeta N, Chen T, Vianello F, Gererty L, Malik A, et al. (2007) Long-term survival of transplanted allogeneic cells engineered to express a T cell chemorepellent. Transplantation 83: 174–183. Irimia D, Liu SY, Tharp WG, Samadani A, Toner M, et al. (2006) Microfluidic system for measuring neutrophil migratory responses to fast switches of chemical gradients. Lab Chip 6: 191–198. Thompson RD, Wakelin MW, Larbi KY, Dewar A, Asimakopoulos G, et al. (2000) Divergent effects of platelet-endothelial cell adhesion molecule-1 and beta 3 integrin blockade on leukocyte transmigration in vivo. J Immunol 165: 426–434. Kircher MF, Allport JR, Graves EE, Love V, Josephson L, et al. (2003) In vivo high resolution three-dimensional imaging of antigenspecific cytotoxic Tlymphocyte trafficking to tumors. Cancer Res 63: 6838–6846.

Chapter 9 Analysis of CXCR3 and Atypical Variant Expression and Signalling in Human T Lymphocytes Anna Korniejewska, Malcolm Watson, and Stephen Ward Abstract Members of the chemokine (Chemotactic cytokines) superfamily and their receptors play a major role in trafficking of immune cells under homeostatic and inflammatory conditions. The chemokine receptor CXCR3 is expressed mainly on activated T lymphocytes and binds three pro-inflammatory, interferonγ-inducible chemokines: monokine induced by IFN-γ (Mig/CXCL9), IFN-γ-induced protein-10 (IP-10/CXCL10) and IFN-γ-inducible T-cell α-chemoattractant (I-TAC/CXCL11). CXCR3 and its agonists are involved in a variety of inflammatory pathologies, making this receptor an attractive target for the design of new anti-inflammatory drugs. Interestingly, a growing body of evidence suggests the existence of at least two novel variants of CXCR3, namely CXCR3-B and CXCR3-alt, which present challenges in the design of new anti-inflammatory drugs targeting CXCR3. In this chapter, we describe the collection, isolation and activation of human peripheral blood-derived T lymphocytes and methods to examine the expression of CXCR3 and its atypical variants at both mRNA and protein levels, as well as protocols for exploring the biochemical and functional responses of T lymphocytes to all known CXCR3 agonists. Key words: T lymphocytes, chemokines, chemokine receptors, CXCR3, chemotaxis, PI3K.

1. Introduction Chemokines are small molecules of approximately 7–10 kDa that form a large cytokine family composed of ˜50 members in the human system. These proteins influence an array of cellular activities such as chemotaxis, adhesion, angiogenesis and proliferation. Approximately 20 different chemokine receptors have been identified as mediators of chemokine activities. All chemokine receptors belong to the large family of seven-transmembrane F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_9, © Springer Science+Business Media, LLC 2010

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domain, G-protein-coupled receptors (GPCR). Chemokines are functionally important for progression of many pathological phenomena including several inflammatory/autoimmune diseases as well as cancer. So-called inflammatory chemokines constitute the vast majority and are inducible and control cell recruitment to sites of infection and inflammation. Certain chemokines are also involved in developmental processes including lymphopoiesis, cardiogenesis and development of the nervous system (1). The transition from bone marrow-resident haematopoietic stem cells through development of T-cell precursors in the thymus, migration into secondary lymphoid organs for immune response initiation and maturation into circulating memory and effector T cells involves sequentially co-ordinated changes in the profiles of chemokine receptor expression to guide cells into the appropriate microenvironment. Characterisation of expression profiles of chemokine receptors has been instrumental in defining subsets of human memory T cells with distinct migratory capacity and effector functions. For example, CCR7 expression discriminates between lymph node-homing central memory T cells and tissue-homing effector memory T cells (2). In addition, CXCR3, CXCR6 and CCR5 are preferentially expressed on Th1 cells (3), while CCR3, CCR4 and CCR8 (along with the PGD2 receptor CRTH2) are expressed on Th2 cells (4, 5) More recently, CCR2, CCR6 and CCR9 have been reported to be expressed on Th17 cells (6–9). The binding of chemokines to their GPCRs stimulates a complex network of intracellular signalling including calcium signalling and activation of phosphoinositide 3-kinase (PI3K) and Akt protein kinase, as well as phospholipase C, MAP kinases (ERK1/2, p38 and JNK) and several small GTPases including Ras and the Rho-family GTPases, Ras and Rac (10, 11). Agonist stimulation also leads to receptor internalisation, providing a regulatory mechanism for intracellular responses by reducing the number of surface-expressed receptors. Following ligand binding, there are two major routes whereby GPCRs are internalised into cells. The first and most well-defined route involves the binding of arrestin to the phosphorylated receptor, which leads to clathrin binding. The receptor–arrestin complex is then sequestered in clathrin-coated pits. This pathway is often considered a default system for degradation and recycling of receptors (12, 13). The second pathway involves invaginations of the cell membrane known as caveolae and functions independently of clathrin-coated pits (14, 15). The chemokine receptor CXCR3 is expressed on a wide variety of cells including activated T lymphocytes, NK cells, malignant B lymphocytes, endothelial cells and thymocytes (16–21). Three major CXCR3 ligands, CXCL9, CXCL10 and CXCL11, have been identified, all of which are induced by IFNγ and are therefore thought to promote Th1 immune responses

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(22–24). Recent studies have shown that different CXCR3 ligands exhibit unique temporal and spatial expression patterns, suggesting that they have non-redundant functions in vivo. Moreover, the CXCR3 ligands share low sequence homology (around 40% amino acid identity) and exhibit differences in their potencies and efficacies at CXCR3, with CXCL11 being the dominant ligand in several assays (23, 25). CXCR3 and its agonists have been implicated in the induction and perpetuation of several human inflammatory disorders (26) including atherosclerosis (27), autoimmune diseases (28), transplant rejection (29, 30) and viral infections (31). These findings have made CXCR3 a popular target for development of new potential anti-inflammatory strategies. In recent years, however, two main variants of CXCR3 receptor have been identified, namely CXCR3-B (32) and CXCR3-alt (33). Both variants are generated via alternative splicing of mRNA encoding the original CXCR3 receptor (henceforth referred to as CXCR3-A). In the case of CXCR3-B, alternative splicing resulted in extension of NH3 terminus by 52 amino acids and this form of receptor has been shown to bind platelet factor 4 (PF4/CXCL4) in addition to the three classical CXCR3 agonists. In contrast CXCR3-alt is a truncated version of CXCR3 (lacking 101 amino acids), which consequently exhibits a dramatically altered C terminus and with a predicted four to five transmembrane domain structure. Despite this drastically modified structure, CXCR3-alt has been shown to bind and respond to CXCL11 (33). In this chapter we describe methods for the isolation and ex vivo activation and expansion of human T lymphocytes in order to characterise the expression of CXCR3-A and its atypical variants as well as protocols for exploring the biochemical and functional responses of T lymphocytes to all known CXCR3 agonists.

2. Materials 2.1. T-Cell Isolation, Activation and Ex Vivo Expansion

R 1. RPMI-1640 tissue culture medium (Gibco /Invitrogen, UK) (see Note 1).

2. Phosphate-buffered saline (PBS without Ca2+ and Mg2+ ) R /Invitrogen, UK). (Gibco 3. Lymphoprep (Ficoll-Paque 1.077 g/mL density) (AxisShield, Cambridgeshire, UK). 4. Heparin (500 U/mL in H2 O) (Sigma-Aldrich, Gillingham, UK). 5. Staphylococcal enterotoxin B (SEB) (Sigma-Aldrich, Gillingham, UK). Stock solutions (1 mg/mL) were prepared in sterile Milli-Q water and stored at –20◦ C.

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6. Phytohaemagglutinin (PHA) (Sigma-Aldrich, Gillingham, UK). Stock solutions (1 mg/mL) were prepared in PBS and stored at –20◦ C. R CD3/CD28 T-cell expander (Invitrogen, 7. Dynabeads Dynal AS, Oslo, Norway).

8. Interleukin-2 (IL-2) (PeproTech, UK). Aliquots of IL-2 (3,600 U/mL) were prepared in RPMI-1640 medium and stored at –80◦ C. 9. Human T-lymphocyte isolation kits: Pan T Cell Isolation Kit, CD8+ T-Cell Isolation Kit II, CD4+ T-Cell Isolation Kit II (Miltenyi Biotec GmbH). 10. 175 cm3 tissue culture flasks (NuncTM , UK). 11. 50 mL transparent polypropylene centrifuge tubes (Greiner Bio-One, UK). 12. MACS magnetic cell separator or Dynal magnetic particle concentrator. 2.2. RNA Extraction, Reverse Transcription and PCR Analysis of CXCR3 Expression

R 1. TRIzol reagent (Invitrogen, UK).

2. Chloroform, propan-2-ol (isopropyl alcohol), ethanol (Fisher Scientific). 3. Omniscript Reverse Transcriptase Kit (Qiagen, UK). 4. Oligo (dT) – a homo-oligomeric deoxyribonucleotide (poly dT) – primers used in the reverse transcription of polyadenylated mRNA (Promega, Madison, WI, USA). 5. RNase inhibitor – RNasin Plus (Promega, Madison, WI, USA). 6. Easy-A high-fidelity PCR master mix (2x, 0.1 U/μL) (Strategene). 7. Distilled water (UltraPureTM DNase/RNase-free, Invitrogen, UK). 8. Forward and reverse primers were designed to amplify the gene of interest. Nucleotide sequences of primers used in the current study are detailed in Table 9.1. 9. Routine electrophoresis-grade agarose (Sigma-Aldrich, UK). 10. 50X Tris-acetate-EDTA (TAE) buffer (242 g Tris base (Sigma-Aldrich, UK), 57.1 mL acetic acid, 100 mL 0.5 M EDTA (both BDH Chemicals Ltd, UK), add deionised water to 1 L and adjust pH to 8.5. TAE buffer is stored at room temperature). 11. Ethidium bromide (Bio-Rad, UK). 12. Gel loading solution (6x concentrated) (Sigma-Aldrich, UK). 13. 1 kb DNA ladder (New England BioLabs, UK).

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Table 9.1 PCR primers to identify CXCR3 and its atypical variants Sequence (5 to 3 )

Product size (bp)

Reference/ source

CXCR3-A sense

GCAAGAGCAGCATCCACATC

770

33

CXCR3-A antisense

GCAAGAGCAGCATCCACATC

CXCR3-alt sense

CCAAGTGCTAAATGACGCCG

622

33

CXCR3-alt antisense

CTCCCGGAACTTGACCCCTGTG

CXCR3-B sense CXCR3-B antisense

ATGGAGTTGAGGAAGTACGGCCCTGGAAG

440

Dr A.J. Mcknight UCB Celltech

Primer’s name

AAGTTGATGTTGAAGAGGGCACCTGCCAC

2.3. Flow Cytometry Analysis for Expression of T-Cell Markers and Receptors of Interest

1. Fluorescein isothiocyanate (FITC)-conjugated mouse antihuman CD3 antibody (isotype IgG1κ, clone WT31) and FITC-conjugated mouse anti-human CD4 (isotype IgG1κ, clone 11830) (BD Biosciences, Oxford, UK); FITCconjugated mouse anti-human CD8 (isotype IgG2B, clone 37006) and phycoerythrin (PE)-conjugated mouse antihuman CXCR3 (isotype IgG1, clone 49801) all at concentrations of 25 μg/mL (R&D Systems, Abingdon, UK). 2. Isotype-matched controls: FITC-conjugated mouse IgG1κ (BD Biosciences, Oxford, UK), FITC-conjugated mouse IgG2B and PE-conjugated mouse IgG1 (R&D Systems, Abingdon, UK). 3. FACS buffer; phosphate-buffered saline (PBS) containing 5% FBS and 0.05% sodium azide (BDH Chemicals Ltd, UK) stored at 4◦ C. 4. 5 mL polystyrene round bottom tubes (BD Falcon, UK).

2.4. Determining Phosphorylation Levels of Intracellular Proteins in Response to Stimulation with Chemokines by Flow Cytometry

1. Human recombinant chemokines CXCL9, CXCL10 and CXCL11 (PeproTech, UK). 2. Polyclonal anti-phospho-S6 ribosomal protein (Ser235/ 236) antibody (cat. no. 2211) produced in rabbit (Cell Signaling Technology). 3. FITC-conjugated anti-rabbit IgG (whole molecule) antibody produced in sheep (Sigma-Aldrich, UK).

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4. Rabbit IgG (Sigma-Aldrich, UK). 5. Formaldehyde, methanol (Fisher Scientific, Loughborough, UK). 1. For antibodies and other reagents see Section 2.3.

2.5. Analysis of Receptor Internalisation by Flow Cytometry

2. Human recombinant CXCL9, CXCL10, CXCL11, CXCL4 (see Section 2.4).

2.6. In Vitro Cell Migration Assay

1. RPMI-1640 medium supplemented with 0.1% bovine serum albumin (BSA) (Sigma-Aldrich, Gillingham, UK). 2. PBS (see Section 2.1). 3. Human recombinant chemokines: CXCL9, CXCL10, CXCL11, CXCL4 (see Section 2.4). R 4. ChemoTx System 96-well chemotaxis plates (Neuroprobe, Gaithersburg, USA).

5. 96-well filter plate funnels (Neuroprobe, Gaithersburg, USA). 6. 96-well opaque white plates (e.g. OptiPlate PerkinElmer, USA). 7. Calcein AM (Molecular Probes, Eugene, OR). 8. CellTiter-Glo reagent (Promega, Southampton, UK).

3. Methods 3.1. Isolation and Ex Vivo Activation and Expansion of Human T Lymphocytes

T-lymphocyte isolation from freshly donated human peripheral blood and their ex vivo expansion provide a useful protocol for studying biochemical and functional events in T lymphocytes. After separation, the peripheral blood-derived mononuclear cells (PBMCs; a mixture of monocytes and lymphocytes) are activated and kept in culture up to 12 days under conditions (see Note 1) which promote T-lymphocyte proliferation, activation and upregulation of CXCR3. 1. Collect whole blood donated by healthy human volunteers in heparinised syringe (500 U per 50 mL of blood) (see Note 2). 2. Dilute collected blood 1:1 in a sterile 175 cm3 tissue culture flask with RPMI-1640 medium and mix gently (see Note 3). 3. Carefully overlay 35 mL of blood–medium mix on 15 mL of Lymphoprep in 50 mL transparent conical centrifuge tubes (e.g. Falcon tubes) and centrifuge at 400g at 20◦ C with the brake off for 30 min (see Note 4). 4. Following centrifugation, the PBMCs fraction containing lymphocytes and monocytes is seen as a “milky” layer on top

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Fig. 9.1. Schematic illustration of PBMC separation with Lymphoprep after centrifugation.

of higher density Lymphoprep (Fig. 9.1). Carefully remove that layer and transfer to fresh 50 mL tube (see Note 4). 5. Following removal, wash removed cells (representing PBMCs) three times in 50 mL of RPMI-1640 medium (see Note 5) and re-suspend in RPMI-1640 medium containing 10% FBS and 1% penicillin–streptomycin (see Note 6). 6. Cells are stimulated with either 5 μg/mL of PHA or 1 μg/mL SEB (see Notes 7 and 8). 7. After 3 days, the non-adherent cells are removed, washed three times in RPMI-1640 medium (50 mL) and resuspended in complete RPMI-1640 medium (containing 10% FBS, 50 U/mL penicillin and 50 μg/mL streptomycin) and supplemented with IL-2 (20 U/mL) (see Note 8). 8. T lymphocytes are then kept in culture for up to 12 days, cells being washed and re-suspended in fresh medium supplemented with 20 U/mL of IL-2 every 2–3 days (see Note 9). 3.2. RNA Extraction, Reverse Transcription and PCR Analysis of CXCR3 Expression

The commercially available anti-CXCR3 antibodies are unable to distinguish between CXCR3-A, CXCR3-B and CXCR3-alt, while reported CXCR3-B antibodies either are not widely available or have limited specificity. In addition, there are currently no reported antibodies to CXCR3-alt. However, expression of individual CXCR3 isoform mRNA in human T lymphocytes can be monitored as described below: 1. Approximately 9 days post-isolation and initial activation, activated human T lymphocytes (5–10 × 106 cells) are

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removed from culture, pelleted and lysed in 1 mL of TRIzol reagent and incubated for 5 min at room temperature (also see Notes 10 and 11). This step allows the complete dissociation of nucleoprotein complexes. 2. Chloroform is added (0.2 mL per 1 mL of TRIzol used) and samples are agitated by hand for 15 s and incubated for 2–3 min at room temperature followed by centrifugation for 15 min at 4◦ C and at no more than 12,000g. After centrifugation, the mixture separates into different phases: lower red phenol–chloroform phase, an interphase and upper aqueous phase containing RNA. 3. The aqueous phase is carefully collected from each sample and transferred to a fresh tube (see Note 12) and RNA was precipitated by mixing with 0.5 mL of isopropyl alcohol. Samples were then incubated for 10 min at room temperature and centrifuged for 10 min at 4◦ C at no more than 12,000g. Precipitated RNA may be seen as a gel-like pellet on the side/bottom of the tube. 4. Supernatants are carefully discarded and RNA is washed in at least 1 mL of 75% ethanol. Samples are mixed by vortexing and centrifuged for 5 min at 4◦ C at no more than 7,500g. 5. Washed RNA pellets are briefly dried by air or vacuum dried for approximately 10 min (see Note 13) and dissolved in RNase-free water containing 0.5% SDS by passing a few times through a 1 mL pipette tip, followed by incubation for 10 min at 55–60◦ C. 6. The concentration of RNA is determined by measuring the absorbance at 260 nm (A260) in a spectrophotometer. 7. RNA dilutions (e.g. 1:50) are prepared in RNase-free water. The same water in which the RNA is diluted is used to calibrate the spectrophotometer (see Note 14). 8. Purity of RNA is estimated by the ratio of the readings at 260 and 280 nm (A260 /A280 ). Partially dissolved RNA solutions have an A260/280 <1.6 if excluded of contaminates such as protein and phenol. 9. The cDNA is prepared by reverse transcription of cellular RNA with oligo (dT) using the Omniscript RT Kit (Qiagen, UK) (see Notes 15 and 16). 10. RNA, oligo (dT) and RNAsin Plus are mixed and incubated for 5 min at 65◦ C. This step helps opening secondary structures of RNA and binding primers. The rest of the mix containing reverse transcriptase (RT) enzyme is added and incubated for 60 min at 40◦ C and 10 min at 70◦ C.

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11. Polymerase chain reaction (PCR) is performed using primers binding to CXCR3 and its variants CXCR3-B and CXCR3-alt described in Section 2.2 (see Notes 17 and 18). 12. PCR products were analysed on 1.2% agarose gels (see Notes 19–23). A 10 μL of PCR reaction was transferred to the fresh Eppendorf tubes and 2 μL of 6x loading buffer was added. If PCR product will not be used in other applications (e.g. cloning), then 6x loading buffer can be added straight to whole PCR reaction. 13. Samples are mixed and loaded into the agarose gel. DNA standard was loaded into the first well (see Note 24). 14. The gel is run at 5 V/cm of agarose gel. The running gel was monitored and power source switched off when the bromophenol blue (from the loading buffer) has run threefourths the length of the gel (see Note 25). 15. Carefully transfer the gel to a UV transilluminator (preferably within its holder) to visualise PCR products (Fig. 9.2, see Notes 26 and 27).

Fig. 9.2. PCR analysis of mRNA expression of CXCR3 and its spliced variants in human lymphocytes. T lymphocytes (9–12 days post-isolation and activation) were lysed in TRIzol reagent and RNA was extracted and reverse transcribed followed by protocols described in Section 2.3. PCR was performed using primers designed against CXCR3A, CXCR3-B and CXCR3-alt. Product sizes obtained were as follows: 770 bp – CXCR3-A, 622 bp – CXCR3-alt and 440 bp for CXCR3-B.

3.3. Flow Cytometry Analysis of CXCR3 Expression

Once T lymphocytes have been isolated and expanded ex vivo in the presence of IL-2, they can be used for the appropriate biochemical and functional studies as required. Here we investigate surface expression of CXCR3 on CD3+ T lymphocytes as well as on the CD4+ and CD8+ subpopulations. Expression of T-cell surface markers such as CD3, CD4 and CD8 as well as characterisation of CXCR3 expression can be performed as follows: 1. Wash T lymphocytes (9–12 days post-isolation and activation) three times in RPMI-1640 medium and re-suspend at

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1 × 106 cells per sample in 90 μL of ice-cold FACS buffer (see Section 2.2). 2. Add 10 μL of anti-CXCR3-PE and either anti-CD3FITC, anti-CD4-FITC, CD8-FITC or appropriate isotypematched immunoglobulin controls (typical concentration 2.5–5 μ/mL) to cell suspensions and incubate for 30 min at 4◦ C in the darkness (see Notes 28–32 and Section 2.3). A sample should be left untreated to serve as a useful negative control. 3. After incubation time, wash cells twice, re-suspend in 300 μL of FACS buffer and transfer to 5 mL polystyrene FACS tubes. Analyse samples using a flow cytometry instrument and appropriate software. In the present study, samples were analysed on a Becton Dickinson FACSCanto instrument using BD FACSDiva software. Gate cells of interest (from unstained sample) using forward and side scatter. Analyse tested samples with at least 10,000 gated events counted per sample (see Note 33). 3.4. Determining Intracellular S6 Ribosomal Phosphorylation in Response to CXCR3 Agonists by Flow Cytometry

Activation of the PI3K-dependent signalling pathway is elicited by most chemokine receptors (10, 11). This can provide readout to determine whether chemokine receptors elicit intracellular signals. Direct measurement of PI3K products in primary cells is technically formidable, so phosphorylation of proteins downstream of PI3K, such as Akt or S6 ribosomal protein, is often used as surrogate readout of PI3K signalling output (34). We find that the antibodies directed against phospho-S6 ribosomal protein (Ser235/236) are much more reliable in FACS-based assays than phospho-Thr308/Ser473-Akt phosphoprotein-specific antibodies and use the following protocol: 1. Wash T lymphocytes (9–12 days post-isolation/activation) three times and re-suspend in serum-free RPMI-1640 medium at concentrations of 2 × 106 cells/mL. Aliquot 500 μL of cell suspension into Eppendorf tubes (see Note 34). 2. After washing steps, incubate cells for 60 min in water bath at 37◦ C (see Note 35). 3. Cells are treated with appropriate vehicle or concentrations of appropriate CXCR3 agonist (e.g. CXCL11) for the time required (usually 1–10 min) (see Note 36). 4. Terminate stimulations by quick centrifugation for 15–30 s at no more than 6,000g at room temperature, aspiration of supernatant and addition of fixing solution (2% formaldehyde in PBS) for 10 min at 37◦ C followed by 1 min on ice. 5. Cells are washed twice in 1 mL of PBS and permeabilised by the addition of 0.5 mL of 90% methanol at –20◦ C, followed by gentle vortexing and incubation at 4◦ C for 30 min.

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6. Wash T cells again in PBS and re-suspend in 100 μL of blocking solution (e.g. PBS containing 0.1% BSA). Incubate for 10 min at room temperature (alternative blocking solution can be used, see Note 28). 7. After blocking step, wash once and re-suspend cells in 100 μL of PBS and add the anti-phospho-S6-specific antibody or matching isotype control (200 ng antibody in PBS). Incubate for 30 min at room temperature. 8. Wash cells twice and incubate in 100 μL of PBS containing FITC-conjugated anti-rabbit IgG secondary antibody (at the optimal dilution according to manufacturer’s instructions, typically a 1 in 200 dilution) for 30 min at room temperature (see Note 31–33). 9. After incubation with appropriate primary and secondary antibodies, wash and re-suspend cells in 500 μL of FACS buffer (see Section 2.3) for flow cytometric analysis using FACSCanto flow cytometer. Analyse data using appropriate software, e.g. FACSDiva or CellQuest software (Fig. 9.3a). 3.5. Analysis of Agonist-Induced Receptor Internalisation by Flow Cytometry

Agonist stimulation of GPCRs including chemokine receptors often leads to receptor internalisation. This process provides an essential regulatory mechanism for intracellular biochemical responses and functional outcomes. If cell numbers are limiting and/or phosphoprotein-specific antibodies that recognise activated components of signalling pathways are of poor quality or sensitivity, another indirect way of assessing agonist-induced receptor activation is to monitor the loss of surface-expressed receptors. Often antibodies against chemokine receptors are of higher quality and specificity and better suited to use in flow cytometry than antibodies directed against intracellular signalling molecules. Antibodies must be directed against extracellular epitopes of the receptor of interest for internalisation assays. In contrast to the use of phosphoprotein-specific antibodies directed against activated components of signalling pathways (described in Section 3.4), monitoring loss of receptor surface expression as a readout of agonist action has the added advantage that a permeabilisation step (as in Section 3.4) is not required for the antibody to encounter its target. 1. Wash and stimulate T cells as described in Section 3.4 (Steps 1–3). 2. Terminate stimulation by adding 90 μL of ice-cold FACS buffer. 3. Add 10 μL of mouse anti-human CXCR3-PE receptor antibody as detailed in Section 3.3 (see Notes 30 and 31). 4. After incubation with antibody, wash samples twice and resuspend in 500 μL of ice-cold FACS buffer (see Section 2.3)

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Fig. 9.3. CXCR3 agonists stimulate phosphorylation of S6 ribosomal protein and internalisation of CXCR3. A total of 1 × 106 mL–1 T lymphocytes (9–12 days post-isolation and activation) were prepared as described in the text and stimulated with CXCL11 (at concentrations indicated) for 2 min. T cells were then either (a) fixed and permeabilised followed by incubation with anti-phospho-S6 ribosomal protein antibody and anti-rabbit Ig-FITC-conjugated secondary antibody and analysis by flow cytometry or (b) incubated with anti-CXCR3-FITC antibody at 4◦ C and analysed by flow cytometry for loss of CXCR3 surface expression. Data is representative of at least three independent experiments.

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for flow cytometric analysis using appropriate FACS instrument as outlined in Section 3.3. Analyse data using appropriate software, e.g. FACSDiva or CellQuest software (Fig. 9.3b). 3.6. In Vitro Directional Cell Migration Assays

T-lymphocyte migration is essential for lymphocyte accumulation at sites of inflammation and homeostatic trafficking into lymph nodes and tissue. To determine the functional responsiveness of CXCR3-A and its variants expressed on T lymphocytes, analysis of the effect of CXCR3 agonists on cell migration is required. Techniques that have been used to study directional leucocyte migration (chemotaxis) in vitro include methods in which chemotaxis is measured by direct observation of cellular migration and methods in which chemotaxis is measured by quantifying leucocyte migration into or across membranes. Several different techniques have been described to measure leucocyte migration in 96-well chemotaxis chambers (35), but we favour the NeuroR probe ChemoTx System 96-well chemotaxis chamber as it permits rapid, sensitive and consistent measurement of T-lymphocyte migrational responses to chemoattractants. It is also relatively simple and high throughput and is easily adapted to study migrational responses of other leucocytes. 1. Before performing cell migration assays, wash T lymphocytes twice in RPMI-1640 medium and re-suspend at 3.2 × 106 cells/mL in RPMI-1640 medium containing 0.1% BSA (see Note 34). R 2. Prepare ChemoTx System 96-well chemotaxis plates (see Note 37). Peel the cover off the container as indicated on the label. Work on a clean flat surface, and touching only the edges lift the clear plastic lid and microplate out of the container. Leave the framed filter in the container.

3. Place the microplate on the surface well-side up; the lettering should be right reading, with the well-labelled A1 in the upper left corner. Do not touch the wells. 4. Leave cells to rest for 30 min in water bath set up at 37◦ C and during that time add inhibitors if required. Prepare appropriate concentrations of chemokines in RPMI-1640 medium containing 0.1% BSA. 5. Set your pipette to dispense 29 μL if your ChemoTx microplate has 30 μL wells (set to 299 μL if using 300 μL wells). The ChemoTx system is designed to allow for variations in delivered volume of ±2 μL (±4 μL for 300-μL wells). Fill lower chambers (wells) of chemotaxis plate with 29 μL of chemokine dilutions or media (avoid air bubbles). 6. Touching only the edges of the frame take the framed 5 μm filter from the container and position it over the

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filled microplate printed side up, with the A1 corner at the upper left. Set the frame down on the microplate so that the four pins in the microplate enter the corresponding holes in the filter frame. Press the frame down firmly at each corner until the frame makes contact with the perimeter of the microplate. Examine the sites to be sure the fluid makes contact with the filter at each completely filled site (Fig. 9.4a). Prepare each point in triplicate.

Fig. 9.4. Migratory responses of human T lymphocytes to CXCR3 agonists. (a) Image showing schematic illustration of Neuroprobe 5 μm pore filter with hydrophobic surface surrounding area above each well. (b) Schematic illustration of single well loaded with chemokine solution, overlaid with filter and cell suspension loaded on the top. (c) Activated T lymphocytes were washed and re-suspended at 3.2 × 106 mL–1 and stimulated with CXCL9, CXCL10 and CXCL11 at the concentrations indicated for 3 h as described in Section 3.6. Data represents mean ± SEM (n = 3).

7. Load 25 μL of previously prepared cell suspension on the top of the filter, as explained in Fig. 9.4b (see Notes 38 and 39). 8. Place the chemotaxis plate in the humidified incubator set up at 37◦ C, 5% CO2 for 3 h (see Note 40). 9. After incubation remove non-migrated cells from the top of the filter by carefully wiping the surface twice with a piece of Whatman paper and centrifuge plate for 10 min at 400g with brake off (see Note 41).

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10. Carefully remove filter and transfer cells that had migrated into wells (bottom chamber) to polystyrene tubes containing 300 μL of ice-cold PBS. 11. Count migrated cells using FACSCanto system for 30 min (see Notes 42–45).

4. Notes 1. All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise noted. Cell culture medium (e.g. RPMI-1640) is supplemented with 10% heat-inactivated foetal bovine serum (FBS) as well as antibiotics streptomycin (50 mg/mL) and penicillin (50 U/mL). 2. Other anti-coagulants such as citrate, acid citrate dextrose or citrate phosphate dextrose may also be used. 3. Alternatively, blood may be diluted with PBS which is cheaper. 4. It is important to avoid shaking or any disturbance (such as brake-on deceleration) to the tubes after centrifugation. Precise visual identification of the mononuclear interphase is necessary for optimum isolation and depends on transparent conical centrifuge tubes. The buoyant mononuclear cell fraction should be opaque, off-white in colour and located between 15 and 20 mL tube marks. A representative diagram of the separated layers after centrifugation with Lymphoprep, as they appear ideally undisturbed in a transparent 50 mL conical tube after purification, is shown in Fig. 9.1. 5. Prolonged exposure to Lymphoprep can be deleterious to T cells in culture, so extensive cell washing is required. 6. The PBMCs are usually re-suspended in a volume of media equivalent to the volume of blood from which they were isolated. 7. PHA is a lectin isolated from plants and acts as a mitogen which induces activation of T cells by cross-linking to glycoproteins on the cell surface. PHA will yield activated T lymphocytes that are predominantly CD8+ . Alternatively, SEB is one of the best known superantigens, which function by binding to MHC class II molecules expressed on the surface of professional antigen presenting cells (APC) present within the PBMC population. SEB then acts as cross-linker subsequently binding to α-chain of T-cell receptor (TCR), stimulating robust activation of

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T lymphocytes. SEB will yield activated T lymphocytes that are predominantly CD4+ . 8. For the rapid expansion of freshly isolated T cells, it is advised to stimulate the isolated PBMCs with antiCD3/CD28 mAb-coated Dynabeads (approximately one bead per T cell). This method of stimulation more accurately mimics the in vivo presentation of antigen to T cells, but avoids engagement/activation of the co-inhibitory receptors such as CTLA-4 (36). Furthermore, this method does not require large numbers of autologous/MHCmatched APC and antigen or allogenic mononuclear cells and mitogen. The T-lymphocyte population should first be purified by negative selection using Pan T-Cell Isolation Kits (as described in Section 2.1). T lymphocytes are not contaminated with monocyte/macrophages and the antibody-coated beads are simply removed using a magnet. T lymphocytes can be sustained and expanded in culture for several weeks. 9. T cells in culture will grow better if they are kept at high density (up to around 1 × 106 cells/mL). Fresh medium containing 20 U/mL IL-2 is added and should be added every 2–3 days. Usually volume of medium is doubled every 2 days. 10. Cell samples lysed in TRIzol can be placed and stored at –80◦ C. 11. Washing cells before adding TRIzol should be avoided because of increased risk of mRNA degradation. 12. The lower organic phase can be collected if isolation of DNA or protein is required. 13. RNA pellet should not be completely dried, because it significantly decreases its solubility. SDS solution should be avoided when RNA will be used in subsequent enzymatic reactions. 14. An absorbance value of 1 at 260 nm corresponds to 40 μg of RNA/mL (for measurements in water). Therefore, concentration of RNA sample = 40 × A260 × dilution factor and total yield = concentration × volume of sample in millilitres. 15. Control sample should be prepared which contains water instead of reverse transcriptase enzyme. This will be needed to exclude a contamination with genomic DNA. 16. The RT-PCR reaction mix is as follows: 10x buffer (2 μL), 5 mM dNTP mix (2 μL), oligo (dT) (0.5 μL), RNAsin Plus (0.5 μL), Omniscript RT (1 μL, 4 units per reaction), 50 ng–2 μg RNA template (1 μg), dH2 O (up to 20 μL volume).

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17. Reaction mix for PCR reactions is as follows: 2x Easy-A high-fidelity PCR master mix (12.5 μL), 10 μM 5 primer (2 μL), 10 μM 3 primer (2 μL), approximately 1 μg cDNA (prepared in Section 3.2, Step 9) and distilled water (up to 25 μL volume). 18. Reaction conditions are as follows: (i) initial denaturation, 5 min, 94◦ C; (ii) denaturation, 1 min, 94◦ C; (iii) annealing, 0.5 min, 60◦ C; (iv) extension, 2 min, 72◦ C; (v) final extension, 10 min, 72◦ C; (vi) final hold, ∞, 4◦ C. 19. EDTA (ethylenediaminetetraacetic acid) solution should be prepared in advance. EDTA will not dissolve completely until the pH is adjusted to about 8.0. For a 500 mL stock solution of 0.5 M EDTA, weigh out 93.05 g EDTA disodium salt (FW = 372.2). Dissolve in 400 mL deionised water and adjust the pH using NaOH. Add water to a final volume of 500 mL. 20. 1.2% of agarose is prepared in 100 mL of 1x TAE buffer and agarose melted by heating in a microwave for 2 min, followed by gentle mixing until completely dissolved. 2 μL ethidium bromide (10 mg/mL) is added to 100 mL of cooled agarose solution (in order to minimise production of ethidium bromide vapour), mixed and slowly poured in an appropriate tank. Required combs are placed into the gel. Any air bubbles should be carefully removed by pushing them away using a pipette tip. Gel is left to set for 30–60 min. 21. The gel may appear set earlier than 30–60 min after pouring. However, running DNA into an agarose gel that is not completely set can lead to poor resolution and give rise to smeary diffuse bands. 22. 1x TAE buffer (running buffer) is added to an appropriate gel running tank. The gel is then placed in the tank and the comb removed. Note that the running buffer should completely cover the gel surface. 23. Ethidium bromide is highly mutagenic and should be handled with extreme caution. Disposable gloves should be worn at all times. Dispose of the contaminated tip into a dedicated ethidium bromide waste container. 24. To make sure that your samples run evenly, take care to avoid using the end wells of gels as edges of the gel may break while transferring from running tank to transluminator. 25. Also ensure the end with the wells is in line with the cathode, which is usually black. The DNA is negatively charged and will migrate towards the positively charged (red) anode.

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26. UV light is carcinogenic and no naked skin must be exposed to this light, therefore eye and face protection, gloves and long sleeves should be worn while working with UV light. 27. Some agarose gel holders are not UV transparent, so in that case gel should be placed onto the surface of UV light box. 28. Flow cytometry antibodies used in the present study did not show any non-specific binding (as shown by lack of isotype control binding). In some cases, where antibody shows high level of non-specific binding cells can be resuspended and incubated for 60 min at room temperature in blocking buffer (PBS with 10% of FBS and 5% of BSA). This treatment (prior to adding the primary antibody) should significantly decrease non-specific binding. 29. All incubations with antibodies/isotype-matched controls should be performed at 4◦ C to minimise internalisation of receptors and loss of surface-bound antibodies. Additionally, presence of sodium azide in the FACS buffer helps prevent capping and shedding or internalisation of the antibody–receptor complexes. Samples must be protected from light while working with fluorochrome-conjugated antibodies. 30. Concentration of antibody may vary and it is usually suggested by the manufacturer. However, in some instances, it is useful to perform the titration curve in order to determine a saturating concentration of antibody for the particular cell type. This can be done by serial dilutions of antibody in the concentration range recommended by the manufacturer. Multicolour analysis can be performed depending on availability of suitable range of fluorochrome-conjugated antibodies and specification of flow cytometry instruments available to researcher. 31. Some antibodies may require longer incubation times. These conditions should be determined in the preliminary experiments. 32. If no fluorochrome-conjugated antibodies are available, un-conjugated primary antibodies can be used followed by incubation with appropriate species-directed fluorochrome-conjugated secondary antibody. Secondary antibodies are typically used at 1:200 dilution (final concentration range 0.1–10 μg/mL), but should be determined by researcher individually. When using fluorochrome-conjugated secondary antibodies, one sample should be treated with secondary antibody alone in the absence of primary antibody, to act as a control for nonselective secondary antibody binding.

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33. It is important to prepare appropriate cell concentrations in your sample. During FACS analysis, the flow rate of the cell suspension should be at the rate of 100–1,000 cells/s. 34. Always make sure that 24 h prior to agonist stimulation, T lymphocytes are washed free of IL-2. This will minimise basal responses. 35. During this incubation time, pharmacological inhibitors can be added if required. 36. Chemokine stocks are prepared in sterile, filtrated PBS containing 0.1% BSA, at 10 μM, aliquoted and stored at –80◦ C (short-term storage at 4◦ C). Further, chemokine dilutions required for cell stimulations are made up in serum-free RPMI-1640 medium. Prepare 100x concentrated chemokine dilutions. R 37. The ChemoTx System is a disposable 96-well cell migration system (Fig. 9.4a and b). Chemotaxis assays and other cell activity assays that take less than 6 h normally do not R require sterile instruments, and the ChemoTx System is not sterilised. Based on the standard 96-well microplate format, it is compatible with most 96-well fluorescent plate readers, as well as most liquid dispensing robots. The system consists of a microplate, a polycarbonate membrane filter and lid. Plates are made of moulded tissue culture-grade transparent polystyrene, with 30 or 300 μL well capacity. The filters for this system are available in a wide range of pore sizes. For T-lymphocyte migration, the 5 micron filter is recommended. It is bonded to a frame for ease of handling and selectively coated with a hydrophobic mask around each of the test sites. The hydrophobic mask allows pipetting of cell suspension directly onto the sites on the top side of the filter, where it stays as hemispherical drops. Additional hydrophobic coatings provide the seal between the bottom of the filter and the rims of the plate, so there is no need for gaskets and clamping hardware. Bubble entrapment is eliminated in both the top and bottom fluids by this design. The lid aids in reducing evaporation and facilitates stacking. There are several possible combinations of plate well volume and exposed filter area. For most fluorescence assays, the 101 series, with 30 μL plate well volume and 8 mm2 filter areas, works well. If the number of migrated cells does not create sufficient signal strength relative to the background, the 106 series, with 25 mm2 area, is recommended. For longer incubations, a unit with 300 μL plate volume will allow for more media to sustain the cells. When using rare or expensive reagents, an alternative is to use the 106 series with 30 μL well volume for the migration.

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38. Pipette a drop of cell suspension onto each site on the filter top. If the sites are 3.2 mm in diameter (with 8 mm2 of exposed filter area), put 20–25 μL of cell suspension on each site. If the sites are 5.65 mm in diameter (with 25 mm2 of exposed filter area), put 50–60 μL of cell suspension on each site. 39. Hold the pipette vertical and express the cell suspension slowly and evenly. Electronic multi-channel pipettes are ideal for this application. A hydrophobic mask on the filter’s top side causes each drop of cell suspension to stay within the perimeter of its site on the filter; no top plate with top wells is needed. 40. Cover the microplate and filter with the clear plastic lid and incubate. For most chemotaxis assays, the filled instrument is incubated at 37◦ C in humidified air with 5% CO2 . Incubation time depends on the type of cells and the chemoattractants being used. 41. Centrifugation will allow the cells which are still attached to the filter to go into the solution and will prevent losing migrated cells while removing the filter. 42. Flow-Count Fluorospheres (Beckman Coulter) are added to each sample prior to flow cytometry to quantify the number of cells that migrate into the lower chamber compared to input cells. Flow-Count Fluorospheres are a suspension of fluorospheres used to determine absolute counts on the flow cytometer. Each fluorosphere contains a dye which has a fluorescent emission range of 525–700 nm when excited at 488 nm. They have uniform size and fluorescence intensity and an assayed concentration allowing a direct determination of absolute counts. 43. Random migration of unstimulated cells will account for some of the cells that pass through the filter. Migrated cells at the negative control sites show the extent of unstimulated random migration (see Note 38), which can then be differentiated from directional migration or chemotaxis. The assay design will attempt to maximise the differential between the number of migrated cells between the negative controls and the treated samples. Data is expressed as number of migrated cells (average from triplicates ± standard deviation, Fig. 9.4c) or chemotaxis index (this can be obtained by dividing the number of cells migrated to chemokine solution by number of cells randomly migrating). 44. As an alternative to flow cytometric analysis of migration, the cells can be loaded with an appropriate fluorescent dye (e.g., Calcein AM). Typical incubation times would

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be for 30 min at 37◦ C. To count the cells that have migrated through the filter, place the microplate in a fluorescence plate reader and count (35). Pipette into one row or column of wells a known serial dilution of Calcein AM-loaded cell suspension to be used. On the filter sites above these wells, do not pipette cell suspension. When the plate is read, this serial dilution serves as a standard with which to compare the readings from the experimental wells. 45. Another alternative to flow cytometry analysis of cell migration is usage of the CellTiter-Glo dye (Promega, UK) and luminescence detection. Incubate your chemotaxis plate as described in Section 3.6, remove filter and place 96well filter plate funnel (Neuroprobe, Gaithersburg, USA) on the plate. On top of this, place a 96-well opaque white plate (e.g. OptiPlate PerkinElmer, USA), invert plate/funnel/white plate “sandwich” and centrifuge for 5 min at 500g at room temperature. After centrifugation you should get around 30 μL (if you are using 30 μL capacity chemotaxis plate) of cell suspension in each of the white plate wells. Add 30 μL of reconstituted CellTiterGlo dye to each well, seal the OptiPlate and place on a gentle shaking platform for 10 min. Read the plate on a Packard TopCount (5 s per well) using a luminescence setting. Run a standard curve alongside each assay to relate the readings to number of migrated cells per well or alternatively express your data as a chemotactic index relative to unstimulated/vehicle control (as explained in Note 43). References 1. O’Hayre M, Salanga CL, Handel TM, Allen SJ. (2008) Chemokines and cancer: migration, intracellular signalling and intercellular communication in the microenvironment. Biochem J 409, 635–49. 2. Sallusto F, Lenig D, Forster R, Lipp M, Lanzavecchia A. (1999) Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature 401, 708–12. 3. Qin S, Rottman JB, Myers P, Kassam N, Weinblatt M, Loetscher M, Koch AE, Moser B, Mackay CR. (1998) The chemokine receptors CXCR3 and CCR5 mark subsets of T cells associated with certain inflammatory reactions. J Clin Invest 101, 746–54. 4. Nagata K, Tanaka K, Ogawa K, Kemmotsu K, Imai T, Yoshie O, Abe H, Tada K, Nakamura M, Sugamura K, et al. (1999) Selective expression of a novel surface molecule

5.

6.

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Chapter 10 Transfection of Indoleamine 2,3 Dioxygenase in Primary Endothelial Cells Petros XE Mouratidis and Andrew JT George Abstract The endothelial cell plays a central role in the control of inflammatory processes. The recruitment of inflammatory leucocytes into the blood vessels is controlled by the expression of adhesion molecules on the endothelium as well as the secretion and presentation of chemokines. Indoleamine 2,3 dioxygenase (IDO) is an enzyme that metabolises tryptophan and is known to be central in the regulation of immune responses. IDO can be expressed on endothelial cells and can alter T-cell responses to the endothelium. We show that IDO can be readily transfected in primary endothelial cells and detail a method to determine the activity of the transfected protein. Key words: IDO, endothelium, inflammation, transfection, T cells, gene transfer, transplantation.

1. Introduction Endothelium can be involved in inflammation-related processes such as graft rejection, atherosclerosis and vasculitides (1, 2). In addition to the effect of endothelial cells in the recruitment of leucocytes into the blood vessels, endothelial cells can also present antigens to white blood cells influencing their activation (3–5) and transmigration (6, 7). Indoleamine 2,3 dioxygenase (IDO) is an intracellular haeme-containing enzyme that catalyses the oxidative cleavage of the indole ring of regulatory molecules such as L-tryptophan via the kynurenine pathway (8, 9). Local depletion of L-tryptophan together with the production of kynurenine metabolites can modulate inflammation responses (10, 11). IDO activation can result in inhibition of the F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_10, © Springer Science+Business Media, LLC 2010

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growth of bacteria, viruses and parasites (12–15) as well as modulation of T-cell responses partly by enhancement of proliferation of regulatory T cells (16–19). In addition increased expression of IDO is related to a favourable environment for tumour growth and graft survival (20–22). Although post-translational modification and differential degradation rate may affect the enzyme activity (23, 24) it is generally accepted that IDO activity depends on its transcription rate and protein levels. An approach that will help determine the role of IDO in inflammation is using gene transfer of an IDO-containing plasmid in primary endothelial cells. Using this approach, enhanced IDO activity can be investigated for its effects on bacterial growth, T-cell responses and tolerance, angiogenesis and tumour growth.

2. Materials 2.1. Isolation and Purification of Endothelial Cells from Saphenous Veins

1. Collagenase II (GIBCO, Paisley, UK): store at 4◦ C. Prepare fresh 1 mg/ml solution of collagenase II in PBS or HBSS. Filter sterilise and warm up to 37◦ C. After use it can be stored at 4◦ C for up to a week without significant loss of activity. 2. PBS or HBSS without calcium and magnesium. 3. Trypsin containing 0.05% EDTA (GIBCO). Store at 4◦ C. Pre-warm at 37◦ C before use. 4. Syringes and needles. 5. CD105 microbeads (Miltenyi Biotech, Auburn, CA, USA). Store at 4◦ C. Keep on ice during use. 6. Buffer for microbeads: It is prepared using 1× PBS, pH 7.4, 2 mM EDTA and 0.5% FCS. Keep on ice during use. 7. MS Columns (Miltenyi Biotech).

2.2. Cell Culture

1. Endothelial SFM media (Invitrogen, Paisley, UK) and endothelial basal media EGM2 (Clonetics, Walkersville, MD, USA). Store at 4◦ C. Prepare medium by combining equal volumes of endothelial SFM and endothelial EGM2 medium. Add 15–20% sterile filtered FBS. Store at 4◦ C. 2. Penicillin/streptomycin (BioWhittaker, Walkersville, MD, USA). Freeze stock solutions. Add 5 ml of stock solution to 500 ml of endothelial medium. Glutamine (BioWhittaker). Freeze stock solutions. Add 5 ml stock solution to 500 ml of endothelial medium.

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3. Endothelial cell growth supplement (ECGS) (BD Biosciences, NJ, USA). Store at 4◦ C. Dissolve ECGS in endothelial medium with no additives in a concentration of 5 mg/ml. Filter sterilise and keep at –20◦ C until use. Add at least 5 ml of stock solution for each 500 ml endothelial medium. 4. Heparin (SIGMA, Poole, UK). Dissolve heparin with endothelial medium containing no additives to make up a 50 mg/ml stock solution. Filter sterilise and add to 500 ml of endothelial medium to achieve a 100 μg/ml final concentration. 2.3. Transfection of Endothelial Cells

1. Microporator MP-100 (Labtech International, Ringmer, UK) 2. Microporation tips and buffers MP-1096 kit (Labtech International). Keep tips and buffers provided in the kit at room temperature until opened. After use keep suspension and electrolytic buffers at 4◦ C.

2.4. IDO Activity Assay

1. Trichloroacetic acid (TCA) (SIGMA). Prepare a 30% solution of TCA in distilled water. Solution can be stored at 4◦ C for a week without significant degradation. 2. Ehrlich reagent prepared using 6.2 ml 1-propanol, 1.5 ml distilled water, 2.6 ml of 70% perchloric acid and 1.5 g of 4-dimethylbenzinamide. 3. Colorimetric plate reader with capability of measuring absorbance at 492 nm.

3. Methods Saphenous veins with a typical length of 4–5 cm that have not been used during the operation are obtained from patients who undergo bypass surgery (see Note 1). Appropriate ethical approval needs to be obtained first in addition to informed consent forms signed by the patients themselves, in accordance with local regulations and guidelines. Veins can be kept in saline for up to 3 days at 2–8◦ C or used immediately so that endothelial cells are isolated (25). Human saphenous vein endothelial cells (HSVEC) are difficult to grow, replicate slowly (doubling times range between 5 and 7 days) and die after 3–4 weeks. Detection of IDO may be performed at the mRNA, protein and activity level. Real-time RT-PCR can be used to determine IDO mRNA levels. Immunoblotting can also be used to determine cytoplasmic IDO protein levels. A protocol to determine

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IDO activity based on detection of L-kynurenine in culture supernatants is described herein. 3.1. Isolation of Endothelial Cells from Saphenous Veins

1. Using a syringe and a needle (if necessary) the lumen of the vein is washed with 10 ml PBS (see also Note 2). 2. Using a syringe and a needle the lumen of the vein is then washed with 10 ml pre-warmed 1 mg/ml collagenase II solution (see Note 3). 3. Using appropriate thread or a tubing clamp, close one part of the vein. 4. Fill the lumen of the vein with collagenase II. 5. Close the other end of the vein using a tubing clamp or appropriate thread so that collagenase II does not leak. 6. Incubate vein at 37◦ C for 10–15 min. 7. After incubation massage vein gently. 8. Cut two sides of the vein open and collect collagenase II solution in a universal. 9. Flush through the lumen of the vein 10 ml of pre-warmed trypsin–EDTA and collect the effluent in the same universal that the collagenase II solution was collected. 10. Add to the mixture 5 ml endothelial medium containing FBS to neutralise trypsin action. 11. Centrifuge the mixture at 500×g for 5 min. 12. Resuspend the pellet in 8 ml endothelial medium and plate cells in a T25 flask. 13. Renew endothelial medium in the flask twice per week until flask is fully confluent (see Note 4). 14. When flask is fully confluent, remove medium and wash cells twice with 5 ml ice-cold PBS. 15. Remove PBS and trypsinise cells with 3 ml trypsin–EDTA for 2–3 min. 16. Add 3 ml endothelial medium. 17. Transfer medium–cell mixture to a universal and centrifuge at 500×g for 5 min. 18. Discard supernatant and wash pellet twice with 5 ml icecold microbead buffer. 19. Count total number of cells. 20. Resuspend 1,000,000 cells in 80 μl microbead buffer together with 20 μl of anti-CD105 microbeads. 21. Incubate on ice for 15 min. To check purity add FITCconjugated human CD105 antibodies and incubate for further 5 min (See Fig. 10.1).

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Fig. 10.1. Saphenous vein endothelial cells. Saphenous vein endothelial cells have been isolated using the protocol described herein and transfected with EGFP using the microporation method. Fluorescent microscopy has been used to obtain bright field (A) and fluorescent (B) images of these cells.

22. After incubation, add 2 ml ice-cold microbead buffer to the mixture. 23. Centrifuge sample at 200×g for 10 min. 24. Discard supernatant and resuspend sample in 500 μl microbead buffer. 25. Place an MS column to a magnet and apply onto it 500 μl microbead buffer containing no cells. 26. After microbead buffer runs through the column, apply sample onto the MS column and allow running through. 27. After sample has run, the column is washed three times with 1 ml ice-cold microbead buffer. 28. Take the MS column off the magnet and flush through 2 ml ice-cold microbead buffer. 29. Collect effluent in a universal. 30. Centrifuge sample at 500×g for 5 min. 31. Resuspend pellet in 6 ml endothelial medium and plate in a T25 flask. 32. Incubate cells in a humidified atmosphere at 37◦ C, 5% CO2 or proceed to FACS analysis to determine purity of cells based on CD105 antibody fluorescence (see Note 5). 33. To continue culture, wash cells with 5 ml sterile ice-cold PBS and renew endothelial medium every 2–3 days. 34. When flask is ∼80% confluent wash cells twice with 5 ml sterile PBS. 35. Remove PBS and add 3 ml pre-warmed 1× trypsin/EDTA solution to the flask. 36. Incubate flask in a 37◦ C incubator for 2–3 min. 37. Make sure that all cells have detached from surface of the flask using visual inspection under the microscope.

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38. Add 5 ml endothelial medium to the flask. 39. Transfer the cell/medium/trypsin solution to a universal. 40. Centrifuge at 500×g for 3 min. 41. Remove supernatant and resuspend in 10 ml endothelial medium. 42. Add 3 ml cell/medium solution in a T25 flask and supplement using another 3 ml endothelial medium to make it a total 6 ml endothelial medium/cell solution. 43. Treat flask as detailed at step 32 for up to 5–6 passages. 44. Centrifuge remaining 7 ml endothelial medium/cell mixture in the universal at 500×g for 3 min and treat according to experimental set-up. 3.2. Transfection of Endothelial Cells Using Electroporation

1. Wash cells in flasks with 5 ml PBS (see Fig. 10.2). 2. Discard PBS and add 3 ml trypsin–EDTA in each T25 flask and incubate at 37◦ C for 2–3 min. 3. Add 3 ml endothelial medium to the flask and transfer solution with cells to a universal.

HUVEC Control Transfected

HSVEC Control Transfected

45 kDa 42 kDa

IDO

Actin

Fig. 10.2. Transfection of pcDNA3.1-IDO in endothelial cells using microporation. Human umbilical vein endothelial cells (HUVEC) and human saphenous vein endothelial cells (HSVEC) were transfected with pcDNA3.1-IDO construct using the microporation method. Whole cell lysates were collected from these cells after 2 days. Immunoblotting was carried out to assess IDO. IDO is present in transfected cells but not in control cells.

4. Centrifuge sample at 500×g for 5 min. 5. Discard supernatant and resuspend cells in 10 ml PBS. 6. Count total number of cells. 7. Centrifuge cells at 500×g for 5 min at 4◦ C and resuspend cells in 1 ml PBS. 8. Transfer 1 ml aliquot to Eppendorf tube and centrifuge at 500×g for 5 min at 4◦ C. 9. Discard supernatant and resuspend cell pellet in 24 μl suspension buffer for every 100,000 cells (see Notes 6 and 7). Keep at 4◦ C for the duration of the experiment.

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10. Add 1 μg DNA for every 24 μl sample. 11. Electroporate cells using the microporation apparatus. Set voltage at 1,350 mV, pulse number at 1, duration at 30 ms and use p10 tips (see Note 8). 12. After electroporation put cell aliquots in endothelial medium in 24-well plates at 37◦ C and leave them incubate overnight. 13. If IDO activity assay is required, then next day replace medium with 0.5 ml endothelial medium per well of a 24-well plate. Also add exogenous L-tryptophan to make a final concentration of 100 μM in endothelial medium. 3.3. Indoleamine 2,3 Dioxygenase Activity Assay

To determine IDO activity a colorimetric activity assay based on detection of L-kynurenine is used (see Note 9). Tryptophan added in the endothelial medium is catabolised by IDO to N-formylkynurenine which is then converted to kynurenine. Kynurenine levels can be quantified and directly correlate to functional IDO activity. Because kynurenine can be further metabolised to downstream products, it is not entirely quantitative, but can provide a relative indication of IDO activity. 1. Prepare a series of L-kynurenine standards in endothelial medium from 0 μM up to 200 μM in a 96-well plate (see Note 10). 2. From the medium of transfected and control cells transfer 100 μl of supernatant to the 96-well plate. Use duplicates for each condition. 3. Prepare fresh 30% trichloroacetic acid (TCA). 4. Add 100 μl of 30% TCA to each well so that a final volume of 15% TCA is used. 5. Incubate the 96-well plate at 50◦ C for 30 min. 6. Centrifuge the plate at 2,000 rpm for 10 min. 7. Transfer 100 μl supernatant from each well to another 96well plate. 8. To each well add 100 μl freshly made Ehrlich reagent. 9. Incubate plate at 65◦ C for 15 min. 10. Allow plate to cool down. 11. Measure absorbance at 492 nm in a colorimetric plate reader. 12. Determine the amount of kynurenine in the supernatant by comparing values from transfected and mock-transfected conditions with values from standards (see Note 11).

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4. Notes 1. A vein longer than 5 cm is necessary to obtain a sufficient number of primary endothelial cells. 2. The protocol can also be applied for endothelial cells from different sources such as umbilical veins and arteries. 3. Take care not to pierce the vein with a needle. If vein is pierced then cut the appropriate part with a scissor and use the part of the vein that remains intact. 4. To get a fully confluent flask with endothelial cells normally a period of 2 weeks is needed. However, depending on the success of the extraction technique it is possible that a shorter or longer period may be required. 5. Purity of endothelial cells can be assayed using anti-CD105 antibody conjugated to FITC using FACS analysis. 6. In our experience the density of cells in microporation suspension buffer is crucial to the success of the technique. Also transfection rates higher than 50% are achieved when cell confluency after microporation in 24-well plates is higher than 70%. 7. Make sure you eliminate bubbles from the microporation suspension buffer. 8. Microporation of endothelial cells can be achieved using different values for voltages and duration and amount of DNA. This has to be optimised by the end user according to the type of endothelial cell in use. 9. IDO activity assay has a sensitivity threshold as high as 5 μM kynurenine. 10. If IDO activity is not detected then reduce amount of endothelial medium as low as 0.4 ml per well of a 24-well plate and repeat the experiment. 11. To compare the number of cells between conditions several assays can be used such as the trypan blue method by simple counting of cells using a cytometer and microscopy. Our preferable method is the sulforhodamine B protein assay (26) for determining the approximate number of adherent cells. References 1. Nourshargh S, Marelli-Berg FM. (2005) Transmigration through venular walls: a key regulator of leukocyte phenotype and function. Trends Immunol 26(3), 157–65.

2. Choi J, Enis DR, Koh KP, Shiao SL, Pober JS. (2004) T lymphocyte-endothelial cell interactions. Annu Rev Immunol 22, 683–709.

IDO in Endothelium 3. Savage CO, Hughes CC, McIntyre BW, Picard JK, Pober JS. (1993) Human CD4+ T cells proliferate to HLA-DR+ allogeneic vascular endothelium. Identification of accessory interactions. Transplantation 56(1), 128–34. 4. Page CS, Holloway N, Smith H, Yacoub M, Rose ML. (1994) Alloproliferative responses of purified CD4+ and CD8+ T cells to endothelial cells in the absence of contaminating accessory cells. Transplantation 57(11), 1628–37. 5. Ma W, Pober JS. (1998) Human endothelial cells effectively costimulate cytokine production by, but not differentiation of, naive CD4+ T cells. J Immunol 161(5), 2158–67. 6. Berg LP, James MJ, Alvarez-Iglesias M, Glennie S, Lechler RI, Marelli-Berg FM. (2002) Functional consequences of noncognate interactions between CD4+ memory T lymphocytes and the endothelium. J Immunol 168(7), 3227–34. 7. Marelli-Berg FM, Frasca L, Weng L, Lombardi G, Lechler RI. (1999) Antigen recognition influences transendothelial migration of CD4+ T cells. J Immunol 162(2), 696–703. 8. Mellor AL, Munn DH. (2004) IDO expression by dendritic cells: tolerance and tryptophan catabolism. Nat Rev Immunol 4(10), 762–74. 9. Grohmann U, Fallarino F, Puccetti P. (2003) Tolerance, DCs and tryptophan: much ado about IDO. Trends Immunol 24(5), 242–8. 10. Lee GK, Park HJ, Macleod M, Chandler P, Munn DH, Mellor AL. (2002) Tryptophan deprivation sensitizes activated T cells to apoptosis prior to cell division. Immunology 107(4), 452–60. 11. Terness P, Bauer TM, Röse L, Dufter C, Watzlik A, Simon H, Opelz G. (2002) Inhibition of allogeneic T cell proliferation by indoleamine 2,3-dioxygenase-expressing dendritic cells: mediation of suppression by tryptophan metabolites. J Exp Med 196(4), 447–57. 12. Däubener W, Spors B, Hucke C, Adam R, Stins M, Kim KS, Schroten H. (2001) Restriction of Toxoplasma gondii growth in human brain microvascular endothelial cells by activation of indoleamine 2,3-dioxygenase. Infect Immun 69(10), 6527–31. 13. Hansen AM, Ball HJ, Mitchell AJ, Miu J, Takikawa O, Hunt NH. (2004) Increased expression of indoleamine 2,3-dioxygenase in murine malaria infection is predominantly localised to the vascular endothelium. Int J Parasitol 34(12), 1309–19.

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14. Schroten H, Spors B, Hucke C, Stins M, Kim KS, Adam R, Däubener W. (2001) Potential role of human brain microvascular endothelial cells in the pathogenesis of brain abscess: inhibition of Staphylococcus aureus by activation of indoleamine 2,3-dioxygenase. Neuropediatrics 32(4), 206–10. 15. Adam R, Rüssing D, Adams O, Ailyati A, Sik Kim K, Schroten H, Däubener W. (2005) Role of human brain microvascular endothelial cells during central nervous system infection. Significance of indoleamine 2,3-dioxygenase in antimicrobial defence and immunoregulation. Thromb Haemost 94(2), 341–6. 16. Frumento G, Rotondo R, Tonetti M, Damonte G, Benatti U, Ferrara GB. (2002) Tryptophan-derived catabolites are responsible for inhibition of T and natural killer cell proliferation induced by indoleamine 2,3-dioxygenase. J Exp Med 196(4), 459–68. 17. Beutelspacher SC, Tan PH, McClure MO, Larkin DF, Lechler RI, George AJ. (2006) Expression of indoleamine 2,3-dioxygenase (IDO) by endothelial cells: implications for the control of alloresponses. Am J Transplant 6(6), 1320–30. 18. Fallarino F, Grohmann U, Hwang KW, Orabona C, Vacca C, Bianchi R, Belladonna ML, Fioretti MC, Alegre ML, Puccetti P. (2003) Modulation of tryptophan catabolism by regulatory T cells. Nat Immunol 4(12), 1206–12. 19. Puccetti P, Grohmann U. (2007) IDO and regulatory T cells: a role for reverse signalling and non-canonical NF-kappaB activation. Nat Rev Immunol 7(10), 817–23. 20. Beutelspacher SC, Pillai R, Watson MP, Tan PH, Tsang J, McClure MO, George AJ, Larkin DF. (2006) Function of indoleamine 2,3-dioxygenase in corneal allograft rejection and prolongation of allograft survival by over-expression. Eur J Immunol 36(3), 690–700. 21. Witkiewicz A, Williams TK, Cozzitorto J, Durkan B, Showalter SL, Yeo CJ, Brody JR. (2008) Expression of indoleamine 2,3dioxygenase in metastatic pancreatic ductal adenocarcinoma recruits regulatory T cells to avoid immune detection. J Am Coll Surg 206(5), 849–54. 22. Munn DH, Mellor AL. (2007) Indoleamine 2,3-dioxygenase and tumor-induced tolerance. J Clin Invest 117(5), 1147–54. 23. Fujigaki S, Saito K, Takemura M, Maekawa N, Yamada Y, Wada H, Seishima M. (2002) L-tryptophan-L-kynurenine pathway metabolism accelerated by Toxoplasma

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gondii infection is abolished in gamma interferon-gene-deficient mice: crossregulation between inducible nitric oxide synthase and indoleamine-2,3-dioxygenase. Infect Immun 70(2), 779–86. 24. Oh GS, Pae HO, Choi BM, Chae SC, Lee HS, Ryu DG, Chung HT. (2004) 3Hydroxyanthranilic acid, one of metabolites of tryptophan via indoleamine 2,3dioxygenase pathway, suppresses inducible nitric oxide synthase expression by enhancing heme oxygenase-1 expression. Biochem Biophys Res Commun 320(4), 1156–62.

25. Tan PH, Chan C, Xue SA, Dong R, Ananthesayanan B, Manunta M, Kerouedan C, Cheshire NJW, Wolfe JH, Haskard DO, Taylor KM, George AJT. (2004) Phenotypic and functional differences between human saphenous vein (HSVEC) and umbilical vein (HUVEC) endothelial cells. Atherosclerosis 173, 171–83. 26. Skehan P, Storeng R, Scudiero D, Monks A, McMahon J, Vistica D, Warren JT, Bokesch H, Kenney S, Boyd MR. (1990) New colorimetric cytotoxicity assay for anticancerdrug screening. J Natl Cancer Inst 82(13), 1107–12.

Section III Migration of T Cells In Vivo

Chapter 11 Visualisation of Lymphoid Organ Development Henrique Veiga-Fernandes, Katie Foster, Amisha Patel, Mark Coles, and Dimitris Kioussis Abstract This chapter provides information on imaging tools that can be employed to visualise and study lymphoid organ development. We focus on the use of genetically modified mouse models that take advantage of fluorescent protein expression in discrete cell populations, thus allowing live cell imaging during lymphoid organogenesis. We discuss approaches that allow characterisation of the cell types involved in the formation of lymphoid organs, including (i) functional assays in explant organ cultures and (ii) highresolution whole-mount immunostaining methods, which are useful for the characterisation of specific cell populations in the context of the whole developing organ. Key words: Lymphoid organogenesis, genetically modified animals, fluorescent proteins, live cell imaging, time-lapse microscopy, stereo fluorescent microscopy, confocal laser scanning microscopy, imaging.

1. Introduction The lymphoid system is organised into discrete capsulated organs or accumulations of diffuse lymphoid tissue. Lymphoid structures are crucial for proper immune cell development and function and are classified as either primary or secondary lymphoid organs. Primary lymphoid organs are sites where lymphopoiesis occurs. In mammals, T cells develop and mature in the thymus, while B cells develop in the foetal liver and in the bone marrow. During T and B cell differentiation, lymphocytes acquire their repertoire of specific antigen receptors, which in the periphery form the basis of adaptive immune responses to pathogens. The F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_11, © Springer Science+Business Media, LLC 2010

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thymus rudiment is first colonised by lymphocyte progenitor cells between embryonic day 10.5 (E10.5) and E12.5 (1). In addition to the haematopoietic cells, the thymic parenchyma is composed of blood vessels, connective tissue and epithelial cells, which form a highly specialised microenvironment (2). Neural crest cells (NCCs) have also been shown to play a major role in the formation of the thymus rudiment contributing to the connective tissues in the thymic capsule and surrounding blood vessels in the thymus (3–5). Secondary lymphoid organs, such as lymph nodes (LN) and Peyer’s Patches (PP), are crucial players in the initiation of immune responses, providing optimal conditions for cellular and molecular interactions that lead to lymphocyte activation, differentiation and homeostasis. The development of these structures occurs during embryonic life, but in some pathological situations lymphoid structures can also be formed in adulthood (tertiary lymphoid organs). During foetal lymphoid organogenesis haematopoietic “inducer” cells interact with stromal “organiser” cells giving rise to lymphoid organ primordia (6–10). Haematopoietic inducer cells are the progeny of a foetal liver progenitor CD3– CD4– cKit+ Il7Rα+ α4β7+ cell (11) that is thought to differentiate into CD3– CD4+ cKit+ Il7Rα+ α4β7+ lymphoid tissue “inducer” cells (LTis) (6, 8, 10). At the sites where secondary lymphoid structures develop, “inducer” cells interact with a cell type from mesenchymal origin, the “organiser” cell (LTo), inducing its full maturation (CD45– Vcam+ Icam+ ). It is believed that upon this initial productive interaction a positive feedback loop involving IL7/IL7R and LT/LTR signalling promotes the formation of lymphoid organ primordia (6, 8, 10). Interestingly, haematopoietic cells that colonise lymphoid organ primordia are heterogeneous (9). Haematopoietic CD45+ CD3– CD19– cells can be subdivided in two main categories: CD4+ CD11c– (LTi) and CD4– CD11c+ (9, 12). The latter cell subtype plays an important initiator role in PP development, hence their name Lymphoid Tissue Initiator cells (LTin), and the tyrosine kinase receptor RET, which is crucial for the enteric nervous system formation, is involved in this process. In embryonic gut explant cultures, the RET ligand ARTN is a strong inducer for CD4– CD11c+ cells to aggregate and probably causes the recruitment of LTi cells and the maturation of stromal organiser cells leading to ectopic lymphoid tissue formation (9, 12). Herein, we address some of the methodological approaches that allow the visualisation of lymphoid organogenesis and the characterisation of the cell types involved in these processes. In particular, we discuss the use of reporter mice expressing genetically encoded fluorescent proteins in a specific cell type, allowing live cell/organ imaging during the development of the immune

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system. We further describe a methodology that is useful for the characterisation of specific cell populations, both at a single cell level and in the context of the whole developing lymphoid organ.

2. Materials 2.1. Cell/Organ Culture

1. Dissection preparation components: dissection scissors, fine dissecting forceps, 50 ml Falcon tubes, 35 mm sterile NUNC Petri dishes, ice bucket, ice, pipette (Integra Biosciences), pipettes ranging from 5 to 50 ml and 100 mm sterile Petri dishes (NunclonTM Surface). 2. Air-buffered Iscove’s Modified Dulbecco’s Medium (AirBuffered IMDM, Invitrogen, Paisley, UK) is supplemented with 2.1 mg/ml sodium chloride (Fisher Scientific, Leicestershire, UK), 0.06 mg/ml penicillin (Sigma-Aldrich, UK), 0.1 mg/ml streptomycin (Sigma-Aldrich) and filter sterilised. 3. RPMI (Sigma-Aldrich) supplemented with 10% foetal calf serum (BioSera, UK), 0.06 mg/ml penicillin, 0.2 mg/ml streptomycin, 2 mM glutamine (all Sigma-Aldrich) and filter sterilised. 4. Calcium- and magnesium-free phosphate-buffered saline (PBS) is prepared by dissolving 8 g/l sodium chloride, 0.2 g/l potassium chloride, 1.15 g/l disodium hydrogen orthophosphate and 0.2 g/l potassium dihydrogen (all Fisher Scientific) in double-distilled water and autoclaved to sterilise. 5. 0.5% SuperQ (Sychem, UK) and 70% Ethanol (Fisher Scientific) diluted in water is used to clean tissue culture equipment and surfaces before use. 6. Lyophilised Artemin (Peprotech, NJ, USA) is reconstituted at 200 ng/μl in water and stored in single use aliquots at –20◦ C. 7. Bovine serum albumin (Sigma-Aldrich) is dissolved at 100 ng/μl in PBS and stored in single use aliquots at –20◦ C. 8. Lyophilised GDNF family receptor alpha 3 (GFRα3, R&D Systems, USA) is reconstituted at 200 ng/μl in PBS containing 0.1% bovine serum albumin and stored in single use aliquots at –20◦ C. 9. Agarose beads are Affi-gel Blue Gel beads (BioRad, CA, USA) 150–300 μm.

2.2. Immunohistochemistry

1. 4 g paraformaldehyde (PFA, Fisher Scientific) is dissolved in 90 ml double distilled water at 68◦ C (see Note 1), adding

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one to two drops of 5 N sodium hydroxide to increase solubility. Once transparent and cooled to room temperature, 10 ml of 10× PBS are added, the pH adjusted to 7.3 with hydrochloric acid and the solution filtered to remove any un-dissolved particles, aliquot and store at –20◦ C. 2. Triton X-100 (Sigma-Aldrich) is dissolved in PBS at 0.1% (PBST, see Note 2). 3. NuSieve GTG agarose (low melting point, BMA, Rockland, ME, USA). 4. Agarose, type 1 (high melting point, Sigma-Aldrich). 5. Benzyl alcohol (Aldrich) and benzyl benzoate (SigmaAldrich) are mixed in a 1:2 dilution (BABB). Methanol (Romil, UK) and BABB are mixed in a 1:1 dilution (see Note 3). 6. Nail varnish (Boots, Nottingham, UK, see Note 4).

3. Methods 3.1. Reporter Mouse Strains Allowing Visualisation of Lymphoid Organogenesis

Current reporter mouse strains used to visualise lymphoid organogenesis are primarily based on expression of genetically encoded fluorescent proteins (FP) under the control of tissuespecific regulatory elements. Fluorescent proteins are invaluable tools to track the dynamics of cellular distribution and behaviour during organogenesis by allowing live imaging of specific cell types. Several strategies can be used to ensure FP expression in specific cell types and some of the more commonly used approaches are listed below. Transgenesis frequently results in multiple copy number mouse strains that may ensure high levels of FP expression. However, in the absence of a locus control region in the transgenic construct, the integration site may result in position effect variegation and unexpected expression patterns of the transgene (13, 14). A more recent transgenic approach takes advantage of the Bacterial Artificial Chromosome (BAC) technology, which allows the use of large genomic DNA sequences (usually around 100 Kb) flanking the transgene. BAC transgenes appear to be less susceptible to position effects, but are normally low copy number and the levels of expression may not be sufficient for live cell imaging. A more reliable system consists of knocking-in the coding sequence of a FP into a specific gene locus. In principle, under these circumstances, the reporter gene mimics the exact physiological expression pattern and levels of the endogenous gene

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locus where the FP was knocked in, but as in other single gene copy integrants levels of FP expression might be inadequate for live cell imaging. Finally, fate-reporter systems can also be used to visualise lymphoid structures. For this purpose, one mouse line expressing Cre recombinase under the control of a given gene promoter and regulatory elements is bred to another that has a FP gene knocked into an ubiquitously expressed gene locus (ROSA26 for example) but is preceded by a triple polyadenylation signal, which acts as a transcription stop, flanked by two loxP sites (15–17). In mice resulting from this breeding cells that express transgenic Cre delete the transcriptional stop in front of the FP and indelibly mark these cells and their progeny by FP expression (see also below).

3.1.1. Mouse Models Driving Direct Expression of FP During Lymphoid Organogenesis

Several mouse models have been used to shed light into lymphoid organogenesis. Rather than describing them in detail, here we discuss some examples that illustrate both the potential and the caveats that are intrinsic to these approaches. Human CD2-GFP transgenic mice (hCD2-GFP) express green fluorescent protein (GFP) under the control of the human CD2 promoter and locus control region (LCR) (17). These mice express GFP in a population of haematopoietic cells aggregating in embryonic sites where lymph nodes and PP will develop in the adult (7, 9) (Fig. 11.1a). CD45+ GFP+ haematopoietic cells in these mice are detected in the foetal liver, blood, intestines and LN anlagen in embryos from day E11.5 onwards. GFP expression in these structures allows micro-dissection of anlagen LN and PP from these embryos and flow cytometric analysis. In these mice phenotypic analysis of GFP-positive cells from days E15.5 and E16.5 shows that this population includes the CD4+ CD3– Il7Rα+ c-Kit+ CD11c– LTi cells and a phenotypically distinct population of CD4– CD3– Il7Rα– c-Kit+ CD11c+ cells, which follow a pattern of distribution and aggregation similar to LTi cells (Fig. 11.1b). Human CD2-DsRed transgenic mice (9) can also be used to follow early events of lymphoid organogenesis; however, DsRed has the advantage of being excited at a higher wave length, where tissue auto-fluorescence is low; thus, signal to noise ratio is improved (18). Murine CD4+ CD3– Il7Ra+ c-Kit+ CD11c– LTi cells are characterised by the expression of the retinoic acid-related orphan receptors (RORs) RORγ and RORγt (19, 20). Using homologous recombination in embryonic stem cells, Eberl and colleagues inserted the coding sequence of eGFP into exon 1 of RORγt (21). In this mouse strain LTi cells express eGFP, but live cell imaging is hindered by low expression levels.

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Fig. 11.1. (continued)

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3.1.2. Cell Fate Reporter Mouse Models for Visualisation of Lymphoid Organogenesis

Fate-reporter systems that express Cre recombinase under the control of hCD2 regulatory elements can be used to visualise events in lymphoid organogenesis. Following breeding to ROSA26 lox-tpa-lox eYfp (15, 16) mice, hCD2 drives the expression of Cre deleting the transcriptional stop in front of the eYfp, thus marking these cells and their progeny by eYFP expression. Analysis of eYFP expression can then be determined by flow cytometry or fluorescence microscopy. Similarly, other Cre expressing lines can be used to study the contribution of a particular cell type during lymphoid organogenesis. As an example, Wnt1-Cre (22) transgenic mice can be bred to ROSA26 lox-tpalox eYfp (15, 16) mice in order to visualise neural crest-derived cells during thymic development (4, 5) (Fig. 11.1c). In a similar breeding, Foxn1-Cre (23) can be used to visualise epithelial cells in the thymus and Tie2-Cre (24) to highlight endothelial cells.

3.2. Live Cell Imaging During Lymphoid Organogenesis

When considering live cell imaging the expression intensity and sensitivity of detection of the FP is a critical factor. Low levels of expression might be sufficient for flow cytometry studies or immunohistochemistry, but will yield poor or undetectable signal for live cell tracking. The protocols addressed in this and the following sections are based on wide-field fluorescence stereo microscopy, since this set-up offers an excellent compromise between sensitivity and versatility.

3.2.1. Microscope Set-Ups for Live Cell Imaging in Intact Lymphoid Organs

Live cell imaging of lymphoid organogenesis requires technology providing maximum sensitivity while preserving tissue integrity and anatomical information. Conventional wide-field microscopy using deconvolution, single-photon confocal microscopy and spinning disc confocal microscopy offer insufficient depth penetration and engender high levels of phototoxicity to be useful for live cell imaging of lymphoid organogenesis. Conversely, wide-field fluorescent stereo microscopy and multiphoton confocal microscopy can provide useful and complementary levels of analysis during lymphoid organogenesis. It is not our aim to

 Fig. 11.1. (continued) Reporter mouse strains for visualisation of lymphoid organogenesis. (a) hCD2-GFP+ embryos were dissected and anlagen lymph node and intestines analysed by stereo fluorescence microscopy. Results show the distribution of GFP+ cells in a lymph node at E15.5 (left) and in a Peyer’s patch primordium at E16.5 (right). (b) E15.5 GFP+ lymph nodes were micro-dissected and digested with collagenase. Cell suspensions were stained with the relevant monoclonal antibodies and analysed by flow cytometry. (c) Wnt1-Cre/ROSA26 lox-tpa-lox eYfp newborn mice were dissected and the thymus analysed by confocal microscopy for the expression of eYFP. Results show a peri-vascular eYFP-positive cell network of neural crest origin. Left: magnification ×10; right: magnification ×40.

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extensively discuss these different microscope set-ups, but rather to balance advantages and inconveniences of these two systems for the study of organogenesis: i. Stereo microscopy has several key advantages when being used in live cell imaging of developing lymphoid organs. It has a wide field of view, low phototoxicity (due to lower magnifications) and very good depth resolution. The main disadvantage of this methodology is the loss of information concerning movement in the z-plane. However, in our opinion this is offset by its simplicity of use and its enormous versatility. ii. Multiphoton microscopes have been heavily used to image lymphocyte and dendritic cell behaviour in adult tissues. This set-up has the advantage of having a very good x, y, z resolution without causing phototoxicity. Furthermore, in addition to imaging to depths up to 250 μm, multiphoton microscopy can generate second harmonic images to directly visualise collagen-containing structures, without the need for additional fluorescent markers. Due to its very good depth penetration and z resolution, this approach is best suited when studying cell interactions during organogenesis. Irrespective of the used microscope set-up, environmental conditions of the sample are critical to preserve the normal behaviour of haematopoietic and stromal cells in tissue explants. Small changes in temperature, oxygen levels or other conditions lead to rapid changes in cellular motility and behaviour. The use of temperature-controlled sample blocks and high oxygen pressure is therefore necessary. 3.2.2. Embryo Dissections and Visualisation of Developing Lymphoid Structures 3.2.2.1. Embryo Dissection and Sample Preparation

The protocols described in this section relate to embryos expressing GFP under the control of the human CD2 regulatory elements. The same approach can however be used for any other reporter system. 1. Mouse embryos are obtained from time matings using super ovulated F1 B6/CBA adult females to hCD2-GFP males. Pregnant mice are killed at selected dates post coitus in a CO2 chamber followed by cervical dislocation. 2. Dissection medium: air-buffered IMDM (Iscove’s Modified Dulbecco’s Medium), to be kept on ice for the duration of the experiment. 3. Dissection preparation components: dissection scissors, fine dissecting forceps, 50 ml Falcon tubes, 35 mm sterile NUNC Petri dishes, ice bucket, ice, pipette (Integra Biosciences), pipettes ranging from 5 to 50 ml, 100 mm sterile TM Petri dishes (Nunclon Surface) and small scissors.

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4. Intact uterus is removed and placed in a 50 ml falcon tube containing air-buffered IMDM. All procedures henceforth are performed on ice. 5. Embryos are separated from the placenta and yolk sac and placed in a 100 mm Petri dish containing air-buffered IMDM. 6. A wide-field stereo fluorescent microscope set-up is used for image capturing with software package for image acquisition and analysis such as Openlab (Improvision). Image editing can be done using Volocity (Improvision) and Adobe Photoshop (Adobe). Filters on the stereo microscope are set to 519 nm for GFP visualisation. 7. Confirmation of GFP expression in the embryos is determined by the presence of fluorescent inguinal and brachial lymph nodes. 8. GFP-positive embryos are micro-dissected and intestines are removed including stomach and caecum. Care has to be taken so that the gut is free of any extra tissue other than the mesenterium (see Note 5). 9. The intestines are then placed in cold air-buffered IMDM and left on ice until microscopic analysis. 10. Images are taken using a stereo fluorescent microscope setup and appropriate image acquisition and analysis software (see Step 6). 3.2.2.2. Time-Lapse Microscopy of Developing Lymphoid Structures

In time-lapse microscopy the same cells or the same structures are photographed at regular time intervals over several minutes or hours. This approach allows insight on the cellular dynamics during organogenesis, mainly focusing on motility and cellular interactions. Here we describe the general principles for embryonic explant intestine culture and the basic procedures for stereo microscope time-lapse imaging. 1. Follow the points 1–9 from Section 3.2.2.1. 2. Culture medium: RPMI-1640 (Sigma) with 10% foetal calf serum, 0.06 mg/ml penicillin, 0.2 mg/ml streptomycin and 2 mM glutamine. Culture preparation components: Nalgene filter units (0.45 μm) and 500 ml sterile flasks; 35 and 100 mm sterile Petri dishes (NunclonTM Surface); 70 μm nylon cell strainers (BD FalconTM ); laminar flow cabinet in tissue culture class 1 facilities; humidified incubator at 37◦ C, 5% CO2 and 95% air, 5% SuperQ, 70% ethanol and racks for holding falcon tubes. 3. Prepare samples in a hood that should be thoroughly disinfected with 0.5% SuperQ followed by 70% ethanol.

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4. The wall of the 70 μm strainer is removed by cutting along the bottom edge such that the base mesh remains as an intact surface (one mesh per plate). 5. RPMI medium is warmed to room temperature. 6. Using a 20 μl tip, a single intestine is gently lifted and placed centrally on the mesh in a 35 mm plate (Fig. 11.2a). 7. Fine dissecting forceps are used to spread out the intestine such that all the intestinal folds are flat on the mesh and the mesenterium lays exposed. 8. In the hood, 2 ml of RPMI medium is added to each place/sample. 9. Samples are taken to the stereo microscope where they are kept at 37◦ C using a temperature control unit and temperature-controlled block (Scientific Systems Design) (see Note 6). Images are taken every minute for a predefined length of time. Image acquisition and analysis can be performed using Openlab (Improvision). Volocity software from Improvision can also be helpful to create movie sequences from acquired data. 3.2.2.3. Migration Assays in Explant Organ Cultures

Insights on the role of cellular and molecular players during organogenesis can be obtained by expression analysis studies and imaging methods. Mounting evidence has emerged suggesting that cell migration within intact organs is a key mechanism, by which immune cells interact with each other or in response to particular molecules (25, 26). Interfering with a molecule’s expression pattern may elucidate its possible role, but while some of these experiments can be easily performed in vitro using cell suspensions, in vivo or ex vivo explant organ cultures are technically more difficult to perform during embryogenesis. Here we describe a functional assay for the formation of ectopic lymphoid structures in the intestine. Although the example we describe takes advantage of RET-positive haematopoietic cells that aggregate near a glial-derived neurotrophic factor ligand (GFL) source, these assays can potentially be used with any other type of

 Fig. 11.2. (continued) Lymphoid organogenesis in explant organ cultures. (a) E15.5 hCD2-GFP+ embryos were dissected and a single gut was placed centrally on a mesh in a 35 mm plate. (b) E15.5 GFP+ intestine spread out on a mesh showing the mesenterium lays exposed. Four agarose beads were placed on the mesenterium using a 20 μl tip. (c) E15.5 GFP+ intestines were cultured in vitro with agarose beads impregnated with BSA (left) or ARTN+GFRα3 recombinant proteins (right). Analysis was performed over 72 h by stereo microscopy. (d) E15.5 GFP+ intestines were dissected and analysed by time lapse. Results show detail of time-lapse analysis. Cell tracks are shown for different cells analysed over 90 min.

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molecules and/or cell types. This method is based on the use of agarose beads impregnated with soluble factors and incubated with explant embryonic intestine cultures. 1. Bead impregnation should be performed 24 h prior to the start of experiment. For the experiments shown here, artemin 200 ng/μl (Peprotech), bovine serum albumin 100 ng/μl (Sigma) and GFRα3 200 ng/μl (R&D Systems) recombinant proteins were used. Beads are Affi-gel Blue Gel (Bio-Rad) 150–300 μm beads. 2. Eppendorf tubes are labelled appropriately and placed on ice. 20 μl of beads solution is placed in each tube. The tubes are then centrifuged at 1,300 rpm for 10 s and any liquid in the tube is removed using a P20 pipette. 3. Beads are washed by putting 20 μl of water in each tube followed by centrifugation at 1,300 rpm for 10 s and removal of any excess liquid. At this phase, beads should remain at the bottom of the tubes and tubes are returned to ice. 10 μl of BSA or 5 μl Artemin and 5 μl GFRα3 is added to its respective tube. Tubes are then flicked gently and incubated at 4◦ C for 24 h before the start of the experiment. 4. Embryo dissection and preparation follows the same rules as in Section 3.2.2.1. 5. Culture medium: RPMI-1640 (Sigma) with 10% foetal calf serum, 0.06 mg/ml penicillin, 0.2 mg/ml streptomycin and 2 mM glutamine. Culture preparation components: Nalgene filter units (0.45 μm) and 500 ml sterile flasks; 35 and 100 mm sterile Petri dishes (NunclonTM Surface); 70 μm nylon cell strainers (BD FalconTM ); laminar flow cabinet in tissue culture class 1 facilities; humidified incubator at 37◦ C, 5% CO2 , 5%; SuperQ, 70% ethanol and racks for holding falcon tubes. 6. Prepare samples in a hood that should be thoroughly disinfected with 0.5% SuperQ followed by 70% ethanol. 7. The wall of the 70 μm strainer is removed by cutting along the bottom edge such that the base mesh remains an intact surface (one mesh per plate). 8. RPMI medium is warmed to room temperature. 9. Using a 20 μl tip, a single intestine is gently lifted and placed centrally on the mesh in a 35 mm plate (Fig. 11.2a). Fine dissecting forceps are used to spread out the intestine such that all the intestinal folds are flat on the mesh and the mesenterium lays exposed. Using a fine tip on a micropipettor, a minute amount of beads (4–5) is placed on the mesenterium at distance from the mesenteric lymph node (Fig. 11.2b).

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10. In the hood, 2 ml of RPMI medium are added to each plate/sample. 11. Samples are photographed using a stereo fluorescent microscope and Openlab software (Fig. 11.2b). 12. Samples are then incubated in a humidified incubator at 37◦ C, 5% CO2 for 24 h, after which images are taken for the first time point. 13. Medium is replaced every 48 h and time-point images are taken up to 96–120 h (Fig. 11.2c). 14. If time-lapse imaging is required for this assay then samples are taken to the stereo microscope where they are kept at 37◦ C using a temperature-controlled block (Scientific Systems Design). Images are taken every minute for a predefined length of time. Image acquisition and analysis can be performed using Openlab (Improvision). Volocity software from Improvision can also be helpful to create movie sequences from acquired data (Fig. 11.2d, Supplementary video 1). 3.3. Whole-Mount Staining During Lymphoid Organogenesis

Whole-mount staining offers particular advantages when studying lymphoid organogenesis. First, the embryonic organs are in general small. Second, many of the cellular components of the developing lymphoid organs are individually identifiable by their morphology and/or localisation. Finally, whole-mount immunostainings can be rendered in three dimensions, allowing precise identification of cellular networks within the studied organ (Fig. 11.3a, b, Supplementary video 2, Supplementary video 3). Several factors may, however, limit this methodology: (i) some staining reagents penetrate the tissue very slowly and thus

Fig. 11.3. Whole-mount staining of lymphoid organs. (a) E13.5 hCD2-GFP+ embryos were fixed, immunostained for GFP, optically cleared and analysed by confocal microscopy. Results show the thymic lobes (T) and the developing brachial lymph nodes (LN). Magnification ×10. (b) The adult thymus was fixed, immunostained for endomucin, optically cleared and analysed by confocal microscopy. Results show the vessel and capillary endomucin-positive network within the thymus. Magnification ×40.

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tissues should not exceed 1,000 μm thick, (ii) clearing wholemount organs/tissues for microscopic visualisation is a compromise between tissue transparency and preservation of staining; indeed, fluorochrome selection is a crucial factor on this method and (iii) antibodies may not recognise cognate epitopes after fixation. In this section we report a whole-mount staining method that produces very good anatomic resolution in developing intact lymphoid organs. 3.3.1. Tissue Preparation and Sectioning

1. Tissues and organs are dissected into Petri dishes containing air-buffered IMDM and put onto ice. 2. Specimens are placed into bijous containing 4 ml 4% paraformaldehyde PFA (see Note 7). For 4% PFA preparation: i. 90 ml dH2 0 is boiled in the microwave, removed and left on the bench to cool until 68◦ C (see Note 1). ii. 4 g paraformaldehyde powder are added while continually stirring. iii. 20 μl 1 M NaOH are added to increase PFA solubility. Once transparent, the PFA is transferred to ice and allowed to cool down to room temperature. iv. When PFA has reached room temperature 10 ml 10× PBS are added. v. The pH is adjusted with HCL to 7.3. vi. The PFA is aliquoted to small volumes and stored at –20◦ C (see Note 1). 3. Tissues are washed three times with 4 ml PBS–Triton X-100 0.1% (PBST, see Note 2) at room temperature for 10 min each. Samples can be stained whole mount, or cut to make sections. For the latter proceed to point 4. 4. 8% agarose gel is prepared by dissolving 4 g low melting point and 4 g high melting point agarose powder in 100 ml dH2 0. 5. Agarose is melted in the microwave on low power until all the powder has dissolved and only a few bubbles remain. 6. Agarose is cooled down to 45◦ C and poured into small Petri dishes. 7. Using forceps, the sample is embedded into the dish and left to cool at 4◦ C for 1 h. 8. Sections are cut to 100 μm thickness using a Vibratome and put into 12-well plates, placing 1 ml PBST into each well and storing at 4◦ C.

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All staining steps in this section are carried out in 12-well plates under mild agitation, each well containing a volume of 1 ml. For whole-mount staining, the time required for antibody incubation, washing and post-fixation should be adjusted according to the thickness of each specimen and to the quality of antibodies. 1. 10% serum is prepared by diluting 1 ml of serum from the same species as the secondary antibody to be used, in 9 ml PBST. 2. Specimens are incubated in 10% normal serum PBST for 30 min. 3. Primary antibody is diluted to a pre-determined concentration in 10% normal serum, PBST. Specimens are incubated in primary antibody for 2 h at room temperature, or overnight at 4◦ C. 4. Specimens are washed three times with PBST for 20 min each. 5. Fluorescently conjugated secondary antibodies are diluted to pre-determined concentration in 10% normal serum PBST. Specimens are incubated for 2 h at room temperature, or overnight at 4◦ C in the dark. 6. Specimens are washed three times with PBST for 20 min each. 7. If staining with a second primary antibody is required, steps 3–6 are repeated. PFA fixation reduces fluorescence from FP; therefore, when staining sections from fluorescent reporter mice an antibody detecting the fluorescent protein is required. 8. Once staining is complete, specimens are fixed in 4% paraformaldehyde for 10 min at room temperature.

3.3.3. Optical Clearing of the Sample

Acquiring clear optical sections through a thick specimen as light passes through it is limited by reflection, refraction, scattering and absorption, which determine its refractive index. It is possible to improve image quality and penetration of light in a biological specimen by removing water, which has a low refractive index compared to the tissue and replacing it with an aqueous solution which has a higher refractive index. The use of BABB matches the refractive index of the tissue with the suspending medium allowing for the visualisation of morphology in tissue samples of up to 1,000 μm thick. 1. Specimens are dehydrated in successive concentrations of methanol in PBST, starting with 25% methanol, then, 50 and 75% for 10 min each and 100% for 1 h. 2. In a glass container, BABB (Benzyl Alcohol and Benzyl Benzoate in a 1:2 dilution) (see Note 3) and 100% methanol

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are mixed to a dilution of 50% BABB and 50% methanol (MeOH). 3. Specimens are incubated in BABB:MeOH for 10–15 min at room temperature. 4. BABB:MeOH is replaced with 100% BABB and incubated for 10–15 min at room temperature. 5. Specimens are mounted in a depression slide with 0.7 mm depth or similar, and 100 μl BABB are added into the depression to prevent air bubbles forming. A coverslip is placed on top, gently pressed on the edges to remove excess liquid and blotted onto a tissue. The edges of the coverslip are sealed with nail varnish and left in the dark at room temperature until the nail varnish has solidified. Slides can be stored at 4◦ C in the dark (see Note 4). 3.3.4. Image Acquisition

Different microscope set-ups can be used to acquire images of whole-mount immunostained samples. For a quick screening or simple identification of specific cell structures, stereo fluorescent microscopy may be sufficient, but confocal laser scanning microscopy is imperative for high-resolution rendering. In this section we focus on a few points that may improve acquisition when using a confocal microscope platform. Samples are scanned at wavelengths appropriate to the fluorochromes used. To reduce the likelihood of bleed-through signal, it is important not to excite more than one fluorescent protein at any one time; a sequential scan function in which each laser is activated in turn should therefore be used. Using confocal microscopy it is possible to image the structure of the organ or tissues by collecting background fluorescence at 520 nm. In order to achieve this, the sample should be excited with a 488 nm laser and emission collected at 490–520 nm. It is important to note that if this step is required, fluorescent antibodies which excite and emit between 470 and 530 nm cannot be used. A stack of serial optical images can be acquired at the optimised z distance for each objective to allow for three-dimensional reconstitution. It is therefore important to check the specifications of each objective lens for z resolution. Images should be saved into 8-bit TIFF format, which can be read by most imaging software packages.

4. Notes 1. Paraformaldehyde solubility in water can be improved by increased temperature and pH. Paraformaldehyde can

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however decompose at temperatures above 68◦ C. Single use aliquots of 4% paraformaldehyde at –20◦ C can be stored for periods up to 4 weeks. 2. PBST must be used fresh and therefore prepared before each set of experiments. 3. All steps involving BABB must be carried out in glass containers and pipettes, since BABB dissolves plastic. BABB is an irritant; thus, it should always be handled with gloves. 4. Any nail varnish can be used to seal the edges of coverslip to the slide to prevent tissue sections drying out. Slides must be fully sealed and dried before taking to the microscope as BABB is potentially damaging to various parts of the microscope. 5. Embryo dissection is a critical step for the maintenance of organ integrity. Care must be taken when the intestine or any other organ is manipulated. Indeed we found that excessive manipulation leads to erosion and excessive background fluorescence of the specimens, hindering further imaging analysis. 6. Time-lapse imaging requires controlled environment conditions. When using temperature-controlled units and temperature-controlled blocks, either commercial or bespoke, please ensure that temperature is maintained constant throughout the duration of the time lapse. Small temperature oscillations can cause drift of the focal plan and result in out of focus frames. As a rule we set the focus plan only when the block temperature is constant. 7. Paraformaldehyde fixation times vary according to size of tissues or organs. As guidance, E15.5 intestine will require 15 min fixation at room temperature. These conditions may however vary according to the tissue and antibodies being used.

Acknowledgments Work in this chapter was funded by the Medical Research Council (MRC), UK. We wish to thank Vassilis Pachnis for helpful discussion; T. Norton and K. Williams for technical assistance. We also wish to thank Prof. D. Vestweber for kindly providing endomucin antibody. H.V.-F. and K.F. were supported by a grant from the European Union: Molecular Imaging LSHG-CT-2003-503259 Supplementary Video 1. Time-lapse video showing mobility of GFP cells in the wall of the gut (low magnification). This

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video shows a time-lapse sequence of E15.5 intestines. Time-lapse images were taken for 90 min. Supplementary Video 2. Three-dimensional reconstitution of an embryo section of E13.5. This sample was immunostained with anti-GFP (green) and the embryo structure is depicted in grey color. This video shows the developing lymph nodes and thymus. Magnification ×10. Supplementary Video 3. Three-dimensional reconstitution of an adult thymus section. This sample was immunostained with anti-endomucin (red). This video shows the vessel and capillary endomucin-positive network within the thymus. Magnification ×40. References 1. Owen JJ, Ritter MA. (1969) Tissue interaction in the development of thymus lymphocytes. J Exp Med 129, 431–42. 2. Petrie HT. (2002) Role of thymic organ structure and stromal composition in steadystate postnatal T-cell production. Immunol Rev 189, 8–19. 3. Le Douarin NM, Jotereau FV. (1975) Tracing of cells of the avian thymus through embryonic life in interspecific chimeras. J Exp Med 142, 17–40. 4. Foster K, Sheridan J, Veiga-Fernandes H, Roderick K, Pachnis V, Adams R, Blackburn C, Kioussis D, Coles M. (2008) Contribution of neural crest-derived cells in the embryonic and adult thymus. J Immunol 180, 3183–9. 5. Muller SM, Stolt CC, Terszowski G, Blum C, Amagai T, Kessaris N, Iannarelli P, Richardson WD, Wegner M, Rodewald HR. (2008) Neural crest origin of perivascular mesenchyme in the adult thymus. J Immunol 180, 5344–51. 6. Mebius RE. (2003) Organogenesis of lymphoid tissues. Nat Rev Immunol 3, 292–303. 7. Coles MC, Veiga-Fernandes H, Foster KE, Norton T, Pagakis SN, Seddon B, Kioussis D. (2006) Role of T and NK cells and IL7/IL7r interactions during neonatal maturation of lymph nodes. Proc Natl Acad Sci USA 103, 13457–62. 8. Mebius RE. (2007) Lymphoid organogenesis: educating stroma. Immunol Cell Biol 85, 79–80. 9. Veiga-Fernandes H, Coles MC, Foster KE, Patel A, Williams A, Natarajan D, Barlow A, Pachnis V, Kioussis D. (2007) Tyrosine kinase receptor RET is a key regulator of

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Peyer’s Patch organogenesis. Nature 446, 547–51. Vondenhoff MF, Kraal G, Mebius RE. (2007) Lymphoid organogenesis in brief. Eur J Immunol 37(Suppl 1), S46–52. Yoshida H, Kawamoto H, Santee SM, Hashi H, Honda K, Nishikawa S, Ware CF, Katsura Y, Nishikawa SI. (2001) Expression of alpha(4)beta(7) integrin defines a distinct pathway of lymphoid progenitors committed to T cells, fetal intestinal lymphotoxin producer, NK, and dendritic cells. J Immunol 167, 2511–21. Fukuyama S, Kiyono H. (2007) Neuroregulator RET initiates Peyer’s-patch tissue genesis. Immunity 26, 393–5. Festenstein R, Tolaini M, Corbella P, Mamalaki C, Parrington J, Fox M, Miliou A, Jones M, Kioussis D. (1996) Locus control region function and heterochromatininduced position effect variegation. Science 271, 1123–5. Kioussis D, Festenstein R. (1997) Locus control regions: overcoming heterochromatininduced gene inactivation in mammals. Curr Opin Genet Dev 7, 614–9. Zambrowicz BP, Imamoto A, Fiering S, Herzenberg LA, Kerr WG, Soriano P. (1997) Disruption of overlapping transcripts in the ROSA beta geo 26 gene trap strain leads to widespread expression of beta-galactosidase in mouse embryos and hematopoietic cells. Proc Natl Acad Sci USA 94, 3789–94. Srinivas S, Watanabe T, Lin CS, William CM, Tanabe Y, Jessell TM, Costantini F. (2001) Cre reporter strains produced by targeted insertion of EYFP and ECFP into the ROSA26 locus. BMC Dev Biol 1, 4.

Visualisation of Lymphoid Organ Development 17. de Boer J, Williams A, Skavdis G, Harker N, Coles M, Tolaini M, Norton T, Williams K, Roderick K, Potocnik AJ, Kioussis D. (2003) Transgenic mice with hematopoietic and lymphoid specific expression of Cre. Eur J Immunol 33, 314–25. 18. Hoffman RM. (2005) The multiple uses of fluorescent proteins to visualize cancer in vivo. Nat Rev Cancer 5, 796–806. 19. Kurebayashi S, Ueda E, Sakaue M, Patel DD, Medvedev A, Zhang F, Jetten AM. (2000) Retinoid-related orphan receptor gamma (RORgamma) is essential for lymphoid organogenesis and controls apoptosis during thymopoiesis. Proc Natl Acad Sci USA 97, 10132–7. 20. Sun Z, Unutmaz D, Zou YR, Sunshine MJ, Pierani A, Brenner-Morton S, Mebius RE, Littman DR. (2000) Requirement for RORgamma in thymocyte survival and lymphoid organ development. Science 288, 2369–73. 21. Eberl G, Marmon S, Sunshine MJ, Rennert PD, Choi Y, Littman DR. (2004) An essential function for the nuclear receptor RORgamma(t) in the generation of fetal lymphoid tissue inducer cells. Nat Immunol 5, 64–73.

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22. Danielian PS, Muccino D, Rowitch DH, Michael SK, McMahon AP. (1998) Modification of gene activity in mouse embryos in utero by a tamoxifen-inducible form of Cre recombinase. Curr Biol 8, 1323–6. 23. Gordon J, Xiao S, Hughes B 3rd, Su DM, Navarre SP, Condie BG, Manley NR. (2007) Specific expression of lacZ and Cre recombinase in fetal thymic epithelial cells by multiplex gene targeting at the Foxn1 locus. BMC Dev Biol 7, 69. 24. Kisanuki YY, Hammer RE, Miyazaki J, Williams SC, Richardson JA, Yanagisawa M. (2001) Tie2-Cre transgenic mice: a new model for endothelial cell-lineage analysis in vivo. Dev Biol 230, 230–42. 25. Bajenoff M, Egen JG, Qi H, Huang AY, Castellino F, Germain RN. (2007) Highways, byways and breadcrumbs: directing lymphocyte traffic in the lymph node. Trends Immunol 28, 346–52. 26. Bajenoff M, Germain RN. (2007) Seeing is believing: a focus on the contribution of microscopic imaging to our understanding of immune system function. Eur J Immunol 37(Suppl 1), S18–33.

Chapter 12 Single-Cell Analysis of Cytotoxic T Cell Function by Intravital Multiphoton Microscopy Thorsten R. Mempel Abstract Novel nonlinear optical imaging modalities, most prominently multiphoton microscopy, allow for the direct in vivo visualization of dynamic biological processes in experimental animals at high spatial and temporal resolution. This has led to important new insights into the orchestration of adaptive immune responses. Here we describe in detail a technique that allows for the monitoring of the cytotoxic function of CD8+ effector T cells in tumor-draining lymph nodes of mice at the single-cell level in vivo using multiphoton intravital microscopy. Key words: Cytotoxic T lymphocytes, T cell receptor, cytotoxicity, multiphoton intravital microscopy, in vivo imaging, popliteal lymph node, tumor, surrogate target cells.

1. Introduction Cellular behavior in vivo is determined by a multitude of extrinsic factors that are impossible to replicate in their full complexity outside of the living organism. In order to audit and refine hypothesis based on findings from in vitro studies, in vivo experiments are required. Multiphoton intravital microscopy (MP-IVM) is a powerful imaging modality that allows for the direct dynamic observation of biological processes in their physiological tissue environment at cellular and subcellular resolution. The physical process of multiphoton excitation hereby allows for deeper optical penetration into tissues, compared to conventional fluorescence techniques, as well as for prolonged observation while limiting phototoxicity and photobleaching (1, 2). The F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_12, © Springer Science+Business Media, LLC 2010

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study of the migratory behavior and function of immune cells is particularly amenable to this technology (3, 4). Here we describe in detail a technique that allows investigation of the efficiency and dynamics of contact-mediated killing by CD8+ effector T cells in the setting of an anti-tumor immune response (5). The technique relies on the footpad implantation of tumors expressing a model antigen and adoptive transfer of T cell receptor (TCR)-transgenic T cells into mice (6). In the tumor-draining popliteal lymph node (popLN) the transferred T cells are activated and differentiate into cytotoxic T lymphocytes (CTL). When B cells pulsed with the peptide-antigen recognized by the TCR transgenic T cells are injected into the same mouse as surrogate CTL target cells, they will be lysed by CTL in the tumor-draining lymph node (LN). By fluorescently tagging CTL and B cells, their interactions can be visualized by MP-IVM. Manipulations that are hypothesized to affect cytotoxic function can be tested for their impact on CTL behavior in vivo using this method.

2. Materials 2.1. Induction of an Antigen-Specific Anti-Tumor CD8+ T Cell Response

1. A tumor cell line that expresses an antigen recognized by an MHC class I-restricted TCR expressed in an available transgenic mouse strain [e.g., CT44, which expresses influenza hemagglutinin (HA) as a model antigen]. 2. A complementary TCR transgenic mouse strain (e.g., CL4) that also expresses enhanced green fluorescent protein (EGFP) in CD8+ T cells (e.g., by expressing the DPE–GFP transgene (5)). 3. Reagents for immunomagnetic cell purification of CD8+ T cells (e.g., mouse CD8 T cell-negative selection kit from Miltenyi Biotec, Bergisch Gladbach, Germany). 4. Recipient mice of the same genetic background as the TCR transgenic strain and the tumor cell line (e.g., Balb/C).

2.2. Preparation and Injection of Surrogate CTL Target Cells

1. Spleen and lymph node cells from a mouse of the same genetic background as the recipient animal (e.g., Balb/C). 2. Reagents for immunomagnetic cell purification of follicular B cells (e.g., mouse anti-CD43 beads from Miltenyi Biotec, Bergisch Gladbach, Germany). 3. Hoechst 33342 and CellTracker Orange, both from Molecular Probes/Invitrogen, Carlsbad, CA.

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1. Ketamine HCl solution, 100 mg/mL (e.g., Ketaset) 2. Xylazine HCl solution, 20 mg/mL (e.g., Rompun) 3. NaCl 0.9% for injection.

2.4. Microsurgical Preparation of the Mouse Popliteal Lymph Node for Intravital Microscopy

1. Custom-built microscope stage (see Fig. 12.1a, j, Note 1) 2. Small animal clipper (e.g., Pocket Pro, Wahl, Sterling, IL) 3. Depilation crème (e.g., Nair, Church & Dwight Co., Princeton, NJ) 4. Cotton tip applicators 5. 2 × 2 in. gauze sponges 6. Surgical instruments: Straight “tough cut” iris scissors (1), Vannas spring scissors with 3–5 mm blades (1), Dumont #5 Forceps standard tip, straight, 0.1 × 0.6 mm, Inox, 11 cm total length (2), Dumont #5 Forceps, Biologie tip, 0.05 × 0.02 mm, Inox, 11 cm total length (2) (e.g., from Fine Science Tools, Foster City, CA) 7. 18 mm #1 round cover glasses 8. Ethyl-2-cyanoacrylate glue (e.g., Crazy Glue) 9. Adhesive tape 10. Suture material (e.g., 5-0 braided silk, Roboz Surgical, Gaithersburg, MD) 11. High-viscosity vacuum grease (e.g., Dow Corning, Midland, MI) 12. Plasticine modeling clay 13. Miniature K type thermocouple (e.g., 5SC-TT-K-40-72 from Omega, Stamford, CT)

2.5. Multiphoton Intravital Microscopy of the Mouse Popliteal Lymph Node

1. Upright multiphoton microscope equipped with (a) at least three non-descanned PMT detectors (e.g., Ultima IV, Prairie technologies, Middleton, WI; TriMScope, LaVision Biotec, Bielefeld, Germany; TCS SP5, Leica, Wetzlar, Germany, LSM 710 NLO Zeiss, Jena, Germany), (b) a high numerical aperture objective lens (e.g., Olympus XLUMPLFL20XW 20X, 0.95 NA, water immersion, 2 mm working distance), and (c) a femtosecond-pulsed infrared laser (e.g., DeepSee, Spectra-Physics/Newport, Mountain View, CA; or Chameleon Ultra II, Coherent, Santa Clara, CA). 2. Small water bath (e.g., Precision 280 series, Thermo Scientific, Waltham, MA) 3. Roller pump (e.g., Masterflex L/S with Easyload II, Cole Parmer, Vernon Hills, IL)

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Fig. 12.1. Equipment and procedure of the popLN preparation for MP-IVM. (a) 1: Clipper. 2: Depilation crème. 3: Gauze sponges. 4: Custom-built microscope stage. 5: Pedestal for positioning of right hindleg. 6: Adhesive tape. 7: Suture. 8: Surgical instruments. 9: Dissembled holding clamp for immobilization of trochanter major. 10: Dissembled holding clamp for immobilization of spine. 11: Vacuum grease loaded in syringe. 12: Plasticine. 13: Cover glasses. 14: Mounting device for cover glass and micromanipulator. 15: Crazy Glue. 16: Circular metal tube for temperature control. 17: Thermocouple. (b) Shaved areas of the experimental animal. (c) Depilated area. (d) Positioning of experimental animal prior to the surgical preparation. Upper insert: Site for spine clamp application. Lower insert: Site for hip fixation. (e–i) Steps of the popLN exposure and creation of a closed chamber. Dashed line: position of skin incision. Small arrowheads: outline of lateral marginal vein. Large arrowhead: position of a characteristic branch of the marginal vein that assists in locating the popLN. Arrow: position of popLN. (g) Ring of vacuum grease (h) Cover glass glued to mounting device (highlighted by a circular dashed line). (i) Circular metal tube sealed to cover glass. (j) Fully assembled preparation. (k) Schematic of the temperature control system.

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4. Digital thermometer (e.g., model 710, BK Precision Corporation, Yorba Linda, CA) 5. Polyethylene tubing (e.g., Intramedic PE160, Becton Dickinson, Franklin Lakes, NJ). 2.6. Image Processing and Quantitative Analysis of Surrogate Target Cell Death

1. Image analysis computer workstation with high graphics processing capability 2. Image analysis software that allows processing and analysis of 4-D (3-D and time) data sets (e.g., Volocity, Improvision, Coventry, England; or Imaris, Bitplane, Zurich, Switzerland). As a more cost-effective alternative to these commercial products, the freeware program ImageJ with its wide array of freely available plugins (http://rsb.info.nih.gov/ij/) can substitute for some of the most relevant functions of image processing and analysis.

3. Methods 3.1. Induction of an Antigen-Specific Anti-Tumor CD8+ T Cell Response

1. The tumor cell line is harvested from culture when subconfluent and injected at an appropriate number (e.g., 106 ) suspended in serum-free buffer subcutaneously into the dorsum of the right foot. 2. The same (day 0) or the following day (day 1) EGFP+ CD8+ T cells that express a transgenic TCR specific for the relevant tumor-expressed antigen are harvested and purified using standard immunomagnetic cell selection techniques and 105 cells injected intravenously in serum-free medium.

3.2. Preparation and Injection of Surrogate CTL Target Cells

1. On the day of the imaging experiment (between days 5 and 7 after the T cell transfer) follicular B cells are harvested from pooled spleen and LN cells of a donor animal using negative immunomagnetic selection against the CD43 surface protein. 2. Purified B cells are fluorescently stained (CellTracker Orange: 10 μM, 15 min; Hoechst 33342: 1.25 μM, 30 min, both at 37◦ C in PBS with 1% FBS), incubated with the appropriate recombinant antigenic peptide at a saturating concentration (typically 1 μM or less) for 60 min at 37◦ C in T cell medium, washed, and kept on ice until injection.

3.3. Surgical Anesthesia

Mice are anesthetized initially by intraperitoneal injection of a mixture of ketamine, xylazine, and NaCl 0.9% (to achieve a dose of 150 mg/kg body weight for ketamine and 10 mg/kg for xylazine). A surgical plane of anesthesia is achieved if the mouse

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does not react to firm pinching of the footpad. Repeat injections during the surgical procedure and later on during the imaging session can be carried out intramuscularly to achieve slower release and more even plasma levels. 3.4. Microsurgical Preparation of the Mouse Popliteal Lymph Node for Intravital Microscopy

1. The right mouse leg, flank, and back are shaved using the clippers (Fig. 12.1b) and depletion crème briefly (<30 s) apply to the dorsal aspect of the right leg, avoiding maceration of the skin (see Note 2). Thoroughly wipe off the crème using moist gauze and repeat if required (Fig. 12.1c). The goal is to remove all hair from this area since fragments of hair remaining in the imaging field produce undesirable autofluorescence. Shaving the left upper leg facilitates intramuscular injections when the mouse is under the microscope. 2. The mouse is positioned on the microscope stage using adhesive tape attached to the left hindleg, the left foreleg, and the tip of the tail and by applying tension to the right hindleg using a sling of suture attached to one of the toes (Fig. 12.1d). 3. The tip of the right trochanter major (a prominent bony pivot of the upper leg bone) is identified through palpation and exposed though a 1 cm skin incision in the flank. The attached connective tissue is teased away and the first holding clamp applied (Fig. 12.1d). During this, the right leg should be in a rotational position so that the popliteal fat pad, which is visible though the skin, is facing straight upward. 4. The dorsal skin is incised over a distance of about 1 cm to expose the upper lumbar spine and a small fraction of its circumference freed of the attached musculature. The second holding clamp is attached to its dorsal aspect (Fig. 12.1d). 5. Using the iris scissors, the skin overlying the popLN is longitudinally incised over a length of about 1 cm, using the lateral marginal vein as a landmark structure (Fig. 12.1e). Performing blunt dissection, using the fine forceps, some the connective and fat tissue to the left of the lateral marginal vein is gently teased away and the popLN is readily exposed (Fig. 12.1f). During all times, the LN should be kept moist by applying 0.9% NaCl (see Note 3). 6. Plasticine blocks are molded onto the leg from both sides at the level of the popLN. Applying some pressure from both sides pushes the popLN into a slightly elevated position, making it more accessible without the need for extensive tissue dissection (Fig. 12.1 g). Then a ring of

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vacuum grease is applied to the dry skin around the incision (Fig. 12.1 h). The resulting basin is filled with 0.9% NaCl. 7. A clean cover glass glued to a micropositioning device mounted on the stage is lowered onto the popliteal fossa to seal off the pool of NaCl solution and thus create a closed chamber, taking care not to inclose air bubbles (Fig. 12.1 h). The cover glass can gently touch the surface of the popLN, but excessive pressure will perturb its blood perfusion and should be avoided. 8. A second ring of vacuum grease is applied to the rim of the cover glass and used to seal a piece of metallic tubing (e.g., made from 16G needle tips) that is shaped to an incomplete circle to align with the cover glass (Fig. 12.1i). The metal tubing rests on another block of plasticine and is attached to flexible plastic tubing that will later be connected to the circular water heating system (Fig. 12.1j, k) (see Note 4). 10. The thermocouple is positioned between the mouse leg and the cover glass in close proximity to the popLN (Fig. 12.1i, j). 11. The stage is transferred to the intravital microscope. The heating element is connected to the roller pump and the waterbath and the pump turned on. The thermocouple is plugged into the digital thermometer. 3.5. Multiphoton Intravital Microscopy of the Mouse Popliteal Lymph Node

1. Through the eyepieces of the microscope the popLN can be distinguished from the surrounding tissue structures by its oval shape and its green autofluorescence under mercury arc lamp illumination (see Note 5). Negatively contrasted blood vessels can be assessed for robustness of perfusion. Any residual movement of the specimen that is noticeable at this point will preclude acquisition of high-resolution 3-D image stacks and should be corrected by readjustment of the preparation under the dissecting microscope. 2. Lymphocyte migration is highly temperature dependent (7). Since the water immersion objective lens functions as a heat sink, the ambient popLN temperature must be actively maintained close to 37◦ C. Perfusing the circular metal tube overlying the cover glass is a cost-effective and convenient approach, since the temperature can be quickly adjusted by changing the flow rate (Fig. 12.1 k). 3. The laser source is tuned to 920 nm and the microscope equipped with appropriate filter combinations to detect blue, green, and orange-red fluorescence emission (e.g., using 460/50, 525/50, and 620/100 nm band-pass filters). After switching to viewing by multiphoton fluorescence excitation, backscattered second harmonic signals from the

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collagen-rich organ capsule and the reticulum are visible in the blue channel and serve as convenient landmark structures to assess the current location within the popLN. The green channel will show fluorescence of CTL-associated EGFP as well as low levels of autofluorescence. The latter will also be detectable in the blue and red channel. 4. To test the viability of the preparation, the baseline migratory behavior of EGFP+ CTL is recorded. To this end, a field of view containing the T cell area close to the T/B border is chosen (typically 150–200 μm away from the LN capsule) and a stack of optical sections (e.g., 11 sections spaced 4 μm apart in the z-axis, corresponding to 40 μm of depth) repetitively recorded (e.g., at 15 s intervals) for 15 min without optical zoom (when using a ×20 lens this typically results in a field size of 500–700 μm). Most commercial image acquisition software allows subsequent browsing through the acquired image sequence along the time axis to visually assess cell motility. At this point attention should also be paid to the stability of the preparation. Either a high frequency jitter caused by the respiratory movement of the mouse or the pulsation of a nearby arterial vessel, but also a slow, gradual drift of the specimen need to be corrected by readjustment of the preparation under the dissection microscope. 5. To obtain images that resolve subcellular detail of the target cells, an optical zoom of ×3 is used (field size of 208 × 208 μm on our instrument). At this point it is important to identify a location where injected B cells will emerge from high endothelial venules (HEV) after intravenous injection (see Note 6). The optimal microscope settings (laser power, PMT gain, offset etc.) to obtain balanced signals of EGFP, CellTracker Orange, and Hoechst 33342 have been determined in pilot experiments and are now loaded. 6. A sufficient number (e.g., 2 × 107 ) of surrogate target B cells are injected either retro-orbitally or though an intravenous catheter (e.g., in the tail vein or the jugular vein) and a time-lapse recording started immediately afterward. Ideally, a continuous recording of several hours should be generated, but because of practical limitations, typically multiple sequential recordings have to be obtained (see Note 7). 3.6. Image Processing and Quantitative Analysis of Target Cell Death

Since naïve follicular B cells are highly motile in LN, their irreversible loss of motility can be used as a read-out for loss of cellular function as a result of CTL-induced apoptosis. The labeling of nuclear DNA by Hoechst 33342 and of the cytoplasm by CellTracker Orange furthermore permits the detection of loss of structural integrity of the target cells on the basis of changes in fluorescent properties (see Note 8).

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In our studies we found that encounter of a peptide-pulsed B cell by a CTL leads to the formation of a stable conjugate in about 70% of cases; the conjugate maintains motility for an average of 10 min at the migratory speed of unconjugated B cells with the B cell leading the way; cessation of motility is followed, after an average of another 10 min, by loss of structural integrity of the B cell, as detected by a decrease in red fluorescence and an concomitant increase in blue fluorescence. The CTL then dissociates from the B cell. Alterations in CTL cytotoxic function can be measured through changes in the duration of the various steps of this process (see Fig. 12.2) (5).

Fig. 12.2. Monitoring target cell apoptosis by MP-IVM. (a) Selected images from a recording of a peptide-loaded B cell interacting with a CTL. Top panels show the B cell track. The bottom panels show the individual red, green, and blue channels from the recordings of the B cell-CTL conjugate. Note the decreasing red cytoplasmic signal and the prominent membrane bleb (large arrowhead) as well as the increasing blue nuclear signal at the end of the sequence. The small arrowhead indicates a green signal resulting from spectral spillover from the Hoechst fluorescence due to increased intensity. (b) Analysis of the data shown in (a). Instantaneous velocity and the ratio of red to blue fluorescence permit the definition of motility arrest and structural lysis as sequential steps during B cell apoptosis. Gray-shaded area indicates duration of the CTL-B cell contact.

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Algorithms for automated object identification for cell tracking typically rely on the fluorescence intensity of the object. Since target B cells undergo changes in fluorescent properties during apoptosis in the described experimental system, manual tracking is required. To track an individual cell, a region of interest (ROI) defining the cell’s dimension is drawn for all frames in which it appears in the imaged tissue volume. This ROI is used to determine the cells position (as center of gravity or geometric center) and its integrated red and blue fluorescence. Information on the calibrated spatial position in the x, y, and z dimension and on the red- and blue-integrated fluorescence for all individual time-points of measured B cell tracks are exported as tab-delimited.txt-files. These can then be computed to obtain parameters describing cell motility (e.g., instantaneous migratory velocity) and structural integrity (e.g., the ratio of red to blue fluorescence) in spreadsheet calculation programs such as Excel (Microsoft corporation, Redmond, WA) or matrix-based computing environments such as Matlab (Mathworks, Natick, MA). Since instantaneous migratory velocities fluctuate over a wide range, we subject them to a time-averaging algorithm to smooth the data, using the mean of the instantaneous value and the five preceding and five following values. Since red and blue fluorescence emission is scattered to different degrees in tissue and the relative detected intensities therefore vary with tissue depth, we normalize, for each recording, the calculated ratios of fluorescence to the average values obtained from all viable B cells in the same field of view.

4. Notes 1. Our microscope stage is manufactured using a Plexiglass plate as basis, onto which various elements for the fixation of the mouse are mounted. These elements for the most part consist of materials available in any hardware store that are shaped to suit their particular function. We found small steel corner brackets, angle style steel shelf support pegs, screws and nuts of various calibers, silicone-based windshield sealer, and casting resin very useful items. The only expensive element is a 3-D micromanipulator, which allows for the controlled positioning of the cover glass on the popLN. 2. Our choice of the popLN for MP-IVM studies over other skin-draining LNs was based on two observations: (i) the quantity of footpad-injected reagents that drains to the LN can be controlled more accurately with the popLN than

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with other skin-draining LNs and (ii) due to the popLN’s remote location to the animal’s trunk, the respiratory movements, which would inevitably cause motion artifacts during the slow image acquisition achieved by laser-scanning microscopy, could be shielded off effectively without applying pressure to the surrounding tissues. Likely as a consequence of this, we more reproducibly detected robust lymph flow in the popLN model. 3. During initial attempts with this procedure, sparing the afferent lymph vessels during the surgical dissection may be a technical challenge. This can be helped by footpad injection of a dye (e.g., Evans blue) in order to outline the draining lymphatic vasculature. Typically three afferent lymph vessels can be identified during the procedure. Blunt dissection (insertion of closed forceps into the tissue followed by their release) minimizes bleeding. If a persistent bleeding still occurs, it can usually be stopped by grabbing and applying pressure on the vessel with forceps for ∼30 s. Perfusion of the popLN occurs through perforating deeper arteries and veins and is therefore not affected by damage to superficial blood vessels. 4. At this step it is important that the metal tube rests strain free on the cover glass. Otherwise slow release of tension (e.g., during warming and softening of the plasticine block) can lead to a gradual drift of the specimen, precluding acquisition from the same depth of the LN. 5. If a conventional epifluorescence unit is unavailable, locating the popLN can be helped by drawing a circle around it on the dry cover glass with a fine marker pen. Oblique bright field epi-illumination with a handheld flashlight then allows viewing of the specimen though the eyepieces of the microscope. Despite poor image contrast the drawn circle helps to identify the popLN. 6. Although the delineation of LN microenvironments is less defined in tumor-draining compared to resting LNs, identification is helped by the collagen fiber reticulum, which is sparser in B follicles and dense at the cortical ridge, the part of the T cell area that borders the B follicle. Here the greatest numbers of HEV are also located. 7. The duration of the recording is limited by (i) the requirement to obtain access to the animal in order to maintain surgical anesthesia, (ii) evaporation of the immersion water, and (iii) restrictions of the either the acquisition software or the computer hardware on the maximum size of the recorded data files. Besides the total length of the recording, the file size can typically be adjusted through the pixel resolution,

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the number of z planes, and the cycle time. We typically use a pixel resolution of 256 × 256, 11 optical sections, and a cycle time of 15 s, since this provides us with a good compromise between file size, image detail, and the ability to follow individual motile cells for sufficient time to obtain meaningful data. 8. The observed decrease in red fluorescence may be explained by loss of labeled cytoplasmic proteins during cell death. The concomitant increase in blue fluorescence may be caused by unquenching of Hoechst 33342 that is densely packed on the DNA of healthy cells, but may be partially released during chromatin condensation and nuclear disintegration. 9. In a recently published study, a fluorescent protein-based, genetically encoded FRET-based apoptosis probe was used to monitor cell death by MP-IVM (8). This probe provides the opportunity to tag target cells that are not amenable to ex vivo labeling and adoptive transfer. References 1. Denk W, Strickler JH, Webb WW. (1990) Two-photon laser scanning fluorescence microscopy. Science 248, 73–6. 2. Helmchen F, Denk W. (2005) Deep tissue two-photon microscopy. Nat Methods 2, 932–40. 3. Cahalan MD, Parker I, Wei SH, Miller MJ. (2002) Two-photon tissue imaging: seeing the immune system in a fresh light. Nat Rev Immunol 2, 872–80. 4. Pittet MJ, Mempel TR. (2008) Regulation of T-cell migration and effector functions: insights from in vivo imaging studies. Immunol Rev 221, 107–29. 5. Mempel TR, Pittet MJ, Khazaie K, Weninger W, Weissleder R, von Boehmer H, von Andrian UH. (2006) Regulatory T cells reversibly suppress cytotoxic T cell func-

tion independent of effector differentiation. Immunity 25, 129–41. 6. Klein L, Trautman L, Psarras S, Schnell S, Siermann A, Liblau R, von Boehmer H, Khazaie K. (2003) Visualizing the course of antigen-specific CD8 and CD4 T cell responses to a growing tumor. Eur J Immunol 33, 806–14. 7. Miller MJ, Wei SH, Parker I, Cahalan MD. (2002) Two-photon imaging of lymphocyte motility and antigen response in intact lymph node. Science 296, 1869–73. 8. Breart B, Lemaitre F, Celli S, Bousso P. (2008) Two-photon imaging of intratumoral CD8 T cell cytotoxic activity during adoptive T cell therapy in mice. J Clin Invest 118, 1390–7.

Chapter 13 Imaging Interactions Between the Immune and Cardiovascular Systems In Vivo by Multiphoton Microscopy Owain R. Millington, James M. Brewer, Paul Garside, and Pasquale Maffia Abstract Several recent studies in immunology have used multiphoton laser-scanning microscopy to visualise the induction of an immune response in real time in vivo. These experiments are illuminating the cellular and molecular interactions involved in the induction, maintenance and regulation of immune responses. Similar approaches are being applied in cardiovascular research where there is an increasing body of evidence to support a significant role for the adaptive immune system in vascular disease. As such, we have begun to dissect the role of T lymphocytes in atherosclerosis in real time in vivo. Here, we provide step-by-step guides to the various stages involved in visualising the migration of T cells within a lymph node and their infiltration into inflamed tissues such as atherosclerotic arteries. These methods provide an insight into the mechanisms involved in the activation and function of immune cells in vivo. Key words: Brachiocephalic artery, imaging, lymph node, multiphoton microscopy, T cells.

1. Introduction Atherosclerosis has been recognised as an inflammatory disease and much of the current research is focused on investigating how this inflammation is regulated. Adaptive immunity is highly involved in atherogenesis (1, 2), but we still have much to learn about its effective role in pathology. The adaptive immune system is a complex collection of cells, which interact in distinct locations in a tightly regulated manner in order to allow the rapid induction of immunity and prevent the induction of F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_13, © Springer Science+Business Media, LLC 2010

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inappropriate responses. Failure of this regulation is associated with development of many inflammatory and autoimmune diseases such as atherosclerosis and rheumatoid arthritis. Importantly, the stochastic generation of antigen receptor diversity produces a low frequency of antigen-specific cells. In order to provide effective immune surveillance, these cells must therefore migrate throughout the body in search of cognate antigen presented upon dendritic cells (DCs). The contact between an antigenpresenting cell and a naïve T cell can have important consequences for the induction of immunity since failure to establish stable cell–cell interactions prevents the optimal activation of T cell responses (3–6), which may be of benefit in therapeutic applications. Analysing the cellular interactions during the induction, maintenance and effector stages of an immune response therefore provides insight into how antigen-specific immunity may be manipulated for therapeutic benefit. Indeed, molecules involved in these interactions (such as CTLA-4, CD28 and CD3) have recently been targeted for therapeutic use. While a range of imaging techniques have been applied to immunology over recent years, few have provided such an insight into the cellular interactions involved in the induction of immune responses as multiphoton laser-scanning microscopy (MPLSM) (3–16). Studies using MPLSM have demonstrated that naïve T cells show rapid motility within the lymph node (LN), contacting multiple DCs per hour, allowing them to effectively scan for antigen (7–9). Following antigen recognition, T cells reduce their velocity and remain associated with the antigen-presenting DC (8–10). The dynamics and kinetics of this interaction have important implications in the induction of immunity, with a stable interaction being required for effective T cell priming, whereas shorter/less contact results in the failure of immunity and/or tolerance (3–6). More recently, the role of specific molecules involved in the activation and regulation of these DC:T cell interactions have been investigated, providing a mechanistic insight into immune activation (11, 17). However, in the majority of inflammatory-based pathogeneses, it is the somatic tissues which are the target of disease. While many important studies have investigated the role of cells within the LNs or in the tissues of these diseases, few have examined the interaction between these sites. Thus, it is often assumed that observations made in the LN apply to cells in the tissue and vice versa (e.g. in vivo observations of CTL activity, which may not carry out killing in the LN under normal conditions). Clearly, this may not actually be the case, and therefore it is important to examine the cellular behaviour of cells in the tissue and in the LN and eventually to image the migration between them. As such, we have begun to investigate the relationships between the immune and the cardiovascular systems by imaging the LN and artery (16) in real time

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in vivo. Lymphocytes infiltrate the arterial wall before and during development of atherosclerosis (16, 18), but we still need to investigate their potential antigens, their contact-dependent activities and cross-talk with other cells within or outside the atherosclerotic vessels. A major hurdle is the limited knowledge regarding secondary lymphoid tissue where priming of atherogenic T cells should take place, such as the draining LNs. Thus, imaging immune cell behaviour directly in intact atherosclerotic vessels as well as the associated lymphoid tissues (if any) could provide important information for the development and use of immune-specific therapies in vascular pathology, as well as an improved understanding of the migration and interactions of cells involved in immune pathology. Here we provide a description of how samples are prepared for imaging in order to analyse the behaviour of T cells in vivo within not only the lymph node but also a potential target tissue, the atherosclerotic blood vessel. While most studies to date have used adoptively transferred populations of antigen-specific T cells, there is an increasing use of mice in which fluorescent reporters are expressed by specific subsets of cells or induced under certain conditions and these could be used in the following protocols. Unfortunately, there are currently few MPLSM systems that provide the tuneable excitation laser, accurate scan head and sensitive detection system essential for in vivo imaging. Hence, most groups use customised microscope systems along with environmental control systems to maintain tissue viability. The methods and notes described here should help in establishing imaging systems for the visualisation of T cell migration within the lymph node and the target tissue it drains.

2. Materials 2.1. Mice

A selection of mice has been used in MPLSM imaging experiments, depending on the exact experimental question. Commonly, T cell receptor (TCR) transgenic animals are used as donors for adoptive transfer, allowing the visualisation of a population of antigen-specific T cells in the recipient animal. Typically, we have used the ovalbumin (OVA)-specific DO11.10 mice on a scid background. Since these mice are unable to generate T or B cells of any other specificity, we can be sure that all the cells analysed are the antigen-specific cells of interest. Alternatively, CD4/CD8 T cell isolation must be performed (19). For vascular imaging, C57BL6- or OVA-specific OT-II mice (on a C57BL6 background) are used as lymphocyte donors for adoptive transfer and apolipoprotein E-deficient mice (ApoE–/– ,

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B6.129P2-ApoEtm1Unc ), showing a marked increase in total plasma cholesterol levels, as recipient animals. In these animals, monocyte attachment to endothelial cells is observed at 6 weeks, foam cell lesions develop as early as 8 weeks and fatty streaks in the proximal aorta and carotid are found at 3 months of age. The lesions increase with age and progress to lesions with less lipid but more elongated cells, typical of a more advanced stage of preatherosclerotic lesion. 2.2. CFSE and CMTPX Stock

1. 5- (and 6-) carboxyfluorescein diacetate, succinimidyl ester (5 (6)-CFDA, SE; Invitrogen, Paisley, UK) is prepared as a 10 mM stock solution in DMSO and kept frozen at –20◦ C in aliquots. 2. Cell TrackerTM Red CMTPX (Invitrogen) is prepared as a 20 mM stock solution in DMSO and kept frozen at –20◦ C in aliquots.

2.3. Media

1. RPMI 1640 (Sigma-Aldrich, Poole, UK). 2. HBSS (Sigma-Aldrich). 3. Complete medium [RPMI 1640, 10% foetal calf serum (FCS), 2 mM L-glutamine, 100 U/mL penicillin, 100 mg/mL streptomycin, 1.25 mg/mL fungizone; all obtained from Invitrogen]. 4. CO2 -independent medium (Invitrogen).

2.4. Other Items

1. Nitex membranes (Cadisch & Sons, London, UK) or 40 μm cell strainers (BD Bioscience, Oxford, UK). 2. Plastic coverslips. 3. 3MTM VetbondTM Tissue Adhesive 1469SB.

3. Methods 3.1. Removal of Lymph Nodes

1. Cull the mouse with an increasing concentration of CO2 . 2. Spray the mouse with 70% ethanol to prevent fur from contaminating the region to be dissected. 3. Lie the cadaver ventral side up, pin the mouse’s paws to the dissection board and place under a light source. 4. Make an incision in the skin of the mouse from the abdomen to mandible. 5. Make further incisions from the central one towards each paw. 6. Separate skin from the abdominal muscles and pin out.

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7. Dissect lymph nodes (e.g. inguinal, brachial, axillary, cervical and mesenteric) and spleen if required (see Note 1). 3.2. Preparation for Adoptive Transfer

1. Harvest peripheral and mesenteric LNs from donor mice into RPMI 1640. 2. Mash the LNs through Nitex mesh or BD cell strainers into 50 mL tube, using a syringe plunger. 3. Wash fresh media through the Nitex to fill the tube. 4. Spin the cells at 400×g for 5 min and resuspend the pellet in an appropriate volume of HBSS for counting cells (see Note 2). 5. Count the cells using trypan blue and a haemocytometer. 6. Spin the cells again and resuspend at a concentration of 107 lymphocytes per mL in appropriate pre-warmed buffer containing dyes. a. If labelling cells with CFSE, resuspend in HBSS. b. If labelling with CMTPX, independent media.

resuspend

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c. Add 0.5 μL of stock CFSE/CMTPX per millilitre of buffer (see Note 3). 7. Immediately transfer cells to 37◦ C water bath and incubate with occasional agitation to prevent cell aggregation. a. For CFSE (0.5 μL stock solution/1 × 107 cells/mL) labelling, incubate for 10–15 min at 37◦ C (loose lid). b. For CMTPX (0.5 μL stock solution/1 × 107 cells/mL) labelling, incubate for 45 min at 37◦ C (loose lid). 8. Wash cells by topping up tube with FCS-containing buffer and centrifuging at 400×g for 5 min (see Note 4). 9. Resuspend pellet in RPMI 1640 and count cells using trypan blue. 10. Spin cells and resuspend at appropriate cell concentration for adoptive transfer (see Note 5). 11. Transfer into congenic-recipient mice by intravenous injection. 3.3. Preparation of LN for Imaging

A step-by-step visual guide to LN imaging is shown in Fig. 13.1 (15). 1. Collect the appropriate LN for imaging and place in CO2 independent medium warmed to room temperature (see Notes 6–8). 2. When ready for imaging prepare the LN by rolling on paper towel (Fig. 13.1c). This dries the LN a little and also allows removal of any fat still attached to the LN.

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Fig. 13.1. Procedure for ex vivo imaging of LN. Isolated LNs are adhered to unbreakable plastic coverslips (a) using veterinary-grade superglue (b). The LN is rolled on blue roll to remove fat (c). A thin layer of Vetbond is spread over slide using tissue paper (d and e) and the LN secured on this (f). The glue is cured by immersing in media (g) before trimming the slide to size (h). Finally, a small amount of grease is spread on the underside of the coverslip to secure in the imaging chamber (i).

3. Place a small drop of Vetbond glue onto the unbreakable plastic coverslip (Fig. 13.1d) and spread using the corner of tissue paper to give a very thin layer over the coverslip (Fig. 13.1e). 4. Add the LN onto the coverslip directly without rolling over the glue-covered surface. Press down gently on the LN with forceps to secure it onto the coverslip but avoid putting too much force on it and damaging the capsule (Fig. 13.1f) (see Note 9). 5. Cure the Vetbond by immersing in the CO2 -independent medium (Fig. 13.1 g). The plastic coverslip can now be cut down to size in order to fit into the imaging chamber (Fig. 13.1 h).

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6. Apply a small amount of grease to the underside of the coverslip (Fig. 13.1i) to adhere it into the imaging chamber. 3.4. Preparation of Brachiocephalic Artery for Imaging

1. Cull and prepare the mouse as described above (Section 3.1, points 1–3). 2. Make an incision in the skin of the mouse from the abdomen to the top of the thorax. 3. Open the abdominal wall below the ribcage. 4. Lift the sternum with tweezers and cut the diaphragm. Then cut away the lower part of the ribcage to partially expose the heart. 5. Draw blood from the heart by sticking the needle of a 1 mL syringe into the apex of the left ventricle (see Note 10). 6. Make a small incision in the right atrium for drainage. 7. Stick a 10 mL syringe into the apex of the left ventricle and slowly flush the blood from the mouse with 10 mL of PBS. 8. Dry the mouse by turning it over and pressing it against paper towels; return to the ventral position and re-pin the appendages. 9. Place the mouse under a dissecting microscope. 10. Remove the remaining ventral part of the ribcage, carefully cut the right clavicle leaving the brachiocephalic artery and its branches intact (Fig. 13.2a). Using microdissection scissors, remove all of the fat from around the ascending aorta and brachiocephalic artery. 11. Remove the fat and tissue surrounding the heart. 12. Cut the brachiocephalic artery distal to where it branches into the right subclavian and right carotid and then cut it at its branching site from the aorta to disclose a Y-shaped specimen (Fig. 13.2b); place in PBS to clean (see Note 11). 13. Take the cleaned piece of the brachiocephalic artery and place in CO2 -independent media warmed to room temperature. 14. When ready for imaging place a small drop of Vetbond glue onto the unbreakable plastic coverslip and spread using the corner of tissue paper to give a very thin layer over the coverslip (Fig. 13.1d, e). 15. Glue down the tail of the Y-shaped specimen of brachiocephalic artery onto the coverslip first and gently lower the artery down from the opposite limb to secure it to the slide. Proceed as described above (Section 3.3, points 5 and 6).

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Fig. 13.2. Dissection of the brachiocephalic artery (a) and vessel isolation for imaging (b). Several red-labelled lymphocytes are detectable mainly in adventitia (c–e, ∼10– 30 μm below the outer surface) and within the autofluorescent elastic fibres (green) in the tunica media (f, ∼30–50 μm below the outer surface) as shown by the set of longitudinal sections of ApoE–/– mouse brachiocephalic artery.

3.5. Perfusion System

The tissue ready for imaging is placed in a custom-built perfusion system to bathe tissue in warmed, oxygenated medium. Unfortunately, no off-the-shelf product is currently suitable for this. However, individual components are relatively easy to combine into a system that allows control over the flow rate, temperature and oxygenation of medium throughout the imaging period (20). 1. RPMI 1640 is pre-warmed in a microwave to around 60◦ C and added to the reservoir, where it is kept warm (see Note 12). 2. A mixture of 95% O2 and 5% CO2 is bubbled through the warmed media at a slow, steady rate. 3. Medium is pumped into the imaging chamber (1.5 mL volume) at a flow rate of 1–2 mL per minute, ensuring adequate perfusion of medium/oxygen and removal of waste. 4. Set the inline heater to warm the media above 37◦ C and monitor the temperature in the imaging chamber. Adjust until the temperature is between 35 and 36◦ C.

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Again, few complete MPLSM systems are commercially available ready for use and many groups have customised existing systems for their own use. Of the reported setups used, several groups (3, 4, 7, 9, 10, 20) describe systems similar to ours, consisting of the following: 1. Solid-state, tunable titanium:sapphire laser system (Chameleon, Coherent Inc., Glasgow, UK). 2. The laser beam is routed into a multiphoton excitation laserscanning system (Radiance 2000MP, Bio-Rad Laboratories) through an upright microscope (E600-FN, Nikon). 3. The objective lens used for all imaging investigations is a CFi-60 Nikon Fluor 40×/0.8 NA water-dipping objective lens. 4. The sample is illuminated with 770–830 nm light, with ∼210 fs pulse duration and 76 MHz repetition frequency (see Note 13). 5. Excited fluorescence passes through a laser blocking filter (E625SP, Chroma Technologies, Rockingham, VT) and detected with a multi-alkali cathode photomultiplier tube (S20 PMT) in an external or non-descanned configuration as part of the Radiance 2000MP system. 6. A filter block can be used to separate fluorescence at 500–550 nm (green dyes or autofluorescence) from higher emitted wavelengths (red-labelled cells). 7. The scans are acquired with 500 lines per second and between 256 × 256 and 1024 × 1024 pixel boxes, for a frame rate of 1.95 fps (see Notes 14–16).

3.7. Analysis

Several image analysis software packages have been used in the analysis of MPLSM data including Imaris (Bitplane, Zurich, Switzerland), MetaMorph (Molecular Devices, Downingtown, PA), Video Savant (IO Industries, Ontario, Canada), Picviewer (John Dempster, University of Strathclyde, Glasgow, UK), Adobe AfterEffects (San Jose, CA), ImageJ (NIH, Bethesda, MD) and Volocity (Improvision, Coventry, UK). Although many of these tools perform several similar analyses, Volocity software allows the visualisation and rendering of images as well as the semiautomated quantitation of cell movement and behaviour within the 3D images (Fig. 13.3). While the exact measurements required depend on the data being analysed, there are certain steps that are commonly used: 1. Objects (cells) are identified based on their fluorescence, identifying the centroid location of all the individual cells in the sample. 2. Objects are linked over time by tracking an object over at least eight time points.

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A.. A

B. B.

D.. D

C. C.

Fig. 13.3. Analysis of T cell migration and interactions in the LN. Using software, individual cells are identified within the 3D image and then tracked over time (a). Multiple cell tracks can be plotted relative to each other to effectively show migration distances or polarised movement (b). The velocity of each track can then be graphed to compare the movement of different samples [e.g. to show the difference between naïve T cells and those recognizing antigen (c)]. Alternatively, the amount of interaction between red T cells and green DCs can be calculated as a colocalisation coefficient and compared between two different samples (d).

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3. Many data sets can be generated simply by tracking – e.g. track velocity, displacement, meandering index, motility coefficient. 4. It is also possible to measure the interaction between DC and T cells using the colocalisation tool to identify green and red voxels. This represents the amount of interaction between cells in sample and varies in different conditions. 5. Results can be displayed in a variety of formats, including showing all tracks in an x, y, z plot, plotting the speed of individual cells or calculating the amount of interaction across the sample (Fig. 13.3).

4. Notes 1. The inguinal lymph node is typically found in the skin flap just above the leg and you can find it by looking at junction of blood vessels that run above it. Axillary lymph nodes are located right along a major blood vessel and sometimes dissection can be tricky because of bleeding. Superficial cervical LN along the side of the jaw muscles of the mouse should not be confused with salivary glands. Take care when isolating the mesenteric LN, since this is often associated with a lot of fatty tissue, which is not required. 2. If using large numbers of donors, it is often worth centrifuging the supernatant from the first wash as this can increase cell yield. 3. It is best to add the fluorescent dye to the appropriate buffer and vortex prior to using to resuspend the cells. There is occasionally precipitation of CMTPX in CO2 independent medium and can require thorough vortexing to prevent clumping when added to cells. 4. The inclusion of FCS in washing buffer is important to remove any remaining free fluorescent dye. 5. Typically transfer 2–5 × 106 antigen-specific cells in a volume of 0.2 mL per recipient. 6. Care is required to avoid damaging the capsule of the LN to preserve structure. However, it is important to avoid collecting associated fat which can decrease the quality of images obtained. A good way to tell if fat is removed is to place the LN in medium; if there is any fat it will float. 7. Generally use the LN draining the site of antigen challenge (e.g. when immunizing subcutaneously in the footpad, image the popliteal LN).

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8. Medium should be warmed to room temperature in advance. Lymph nodes can be harvested from several mice and maintained in CO2 -independent medium for several hours. Keep tissues at room temperature and avoid placing on ice. 9. Normally place the LN onto the slide in a transverse orientation (i.e. on its side, with an apparent “kidney” shape). 10. Draw blood by slowly pulling the plunger, twisting the needle. Do not draw for longer than a minute as blood will start to clot. 11. When handling the brachiocephalic artery, it is important to touch only at its ends and not in the middle to avoid damaging the adventitial layer. 12. Heating the medium as gas is bubbled through it helps prevent the build-up of bubbles in the imaging chamber after the inline heater. These bubbles can often form on the tissue or objective lens, obscuring the images. 13. Excitation wavelengths from the Ti:sapphire laser are in the range 770–830 nm, depending on fluorochrome. For single-colour (CFSE) and/or autofluorescence imaging, excitation around 770–780 nm is used. For imaging both CFSE- and CMTPX-labelled cells, excitation is at 830 nm. Green autofluorescence of adventitial connective tissue and elastin fibres in the arterial media is always visible in the range 770–830 nm. Depending on laser power, green autofluorescence of superficial tissue nuclei can be detected at an excitation of 770–780 nm. 14. When setting up LN for imaging, focus to the top of the LN and then z-step down through the tissue to image at a known depth. Typically this is around 200–500 μm from the top of the LN. When setting up an artery for imaging, focus to the top of the adventitia (autofluorescent connective tissue) and then z-step down through the tissue to image at a known depth (Fig. 13.2c–f). Lamina adventitia is a major site of arterial wall lymphocyte infiltration into atherosclerotic arteries; typically this is around 0–30 μm from the top of the vessel. Lymphocyte infiltration is randomly distributed throughout the arteries. 15. Typically scan an area of 256 × 256 pixels (284 μm × 284 μm) in x and y, with z-depth of 35–50 μm (in 2.55 μm steps). 16. The depth of the stack imaged is determined by how fast cells are moving in the LN – faster cells require fast imaging in order to track in software. Generally, imaging a cell once every ∼30 s is sufficient for accurate tracking.

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Acknowledgments The authors would like to thank Dr. Gary Dever (Centre for Biophotonics, Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde) for the experimental support. This work was supported by a BHF project grant (PG/06/083/21198 awarded to PG, JMB, PM and Prof Iain McInnes) and from Wellcome Trust project grants (GR066890MA awarded to JMB, PG and Prof R.S. Phillips; GR068895 awarded to PG, JB, Prof. A. Gurney, Mr. D. Wokosin). PM is supported by a Capacity Building Award in Integrative Mammalian Biology funded by the BBSRC, BPS, KTN, MRC and SFC. OM is funded through an RCUK fellowship. References 1. Robertson AK, Hansson GK. (2006) T cells in atherogenesis: for better or for worse? Arterioscler Thromb Vasc Biol 26, 2421–32. 2. Hansson GK, Libby P. (2006) The immune response in atherosclerosis: a double-edged sword. Nat Rev Immunol 6, 508–19. 3. Hugues S, Fetler L, Bonifaz L, Helft J, Amblard F, Amigorena S. (2004) Distinct T cell dynamics in lymph nodes during the induction of tolerance and immunity. Nat Immunol 5, 1235–42. 4. Shakhar G, Lindquist RL, Skokos D, Dudziak D, Huang JH, Nussenzweig MC, Dustin ML. (2005) Stable T cell-dendritic cell interactions precede the development of both tolerance and immunity in vivo. Nat Immunol 6, 707–14. 5. Zinselmeyer BH, Dempster J, Gurney AM, Wokosin D, Miller M, Ho H, Millington OR, Smith KM, Rush CM, Parker I, Cahalan M, Brewer JM, Garside P. (2005) In situ characterization of CD4+ T cell behaviour in mucosal and systemic lymphoid tissues during the induction of oral priming and tolerance. J Exp Med 201, 1815–23. 6. Millington OR, Gibson VB, Rush CM, Zinselmeyer BH, Phillips RS, Garside P, Brewer JM. (2007) Malaria impairs T cell clustering and immune priming despite normal signal 1 from dendritic cells. PLoS Pathog 3, 1380–87. 7. Miller MJ, Wei SH, Parker I, Cahalan MD. (2002) Two-photon imaging of lymphocyte motility and antigen response in intact lymph node. Science 296, 1869–73. 8. Mempel TR, Henrickson SE, Von Andrian UH. (2004) T-cell priming by dendritic

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cells in lymph nodes occurs in three distinct phases. Nature 427, 154–59. Miller MJ, Hejazi AS, Wei SH, Cahalan MD, Parker I. (2004) T cell repertoire scanning is promoted by dynamic dendritic cell behavior and random T cell motility in the lymph node. Proc Natl Acad Sci USA 101, 998–1003. Celli S, Garcia Z, Bousso P. (2005) CD4 T cells integrate signals delivered during successive DC encounters in vivo. J Exp Med 202, 1271–78. Schneider H, Downey J, Smith A, Zinselmeyer BH, Rush C, Brewer JM, Wei B, Hogg N, Garside P, Rudd CE. (2006) Reversal of the TCR stop signal by CTLA-4. Science 313, 1972–75. Millington OR, Zinselmeyer BH, Brewer JM, Garside P, Rush CM. (2007) Lymphocyte tracking and interactions in secondary lymphoid organs. Inflamm Res 56, 391–401. Okada T, Miller MJ, Parker I, Krummel MF, Neighbors M, Hartley SB, O’Garra A, Cahalan MD, Cyster JG. (2005) Antigenengaged B cells undergo chemotaxis toward the T zone and form motile conjugates with helper T cells. PLoS Biol 3, e150. Garside P, Brewer JM. (2008) Real-time imaging of the cellular interactions underlying tolerance, priming, and responses to infection. Immunol Rev 221, 130–46. Matheu MP, Parker I, Cahalan MD (2007) Dissection and 2-Photon Imaging of Peripheral Lymph Nodes in Mice. J Vis Exp http://www.jove.com/index/Details. stp?ID=265

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16. Maffia P, Zinselmeyer BH, Ialenti A, Kennedy S, Baker AH, McInnes IB, Brewer JM, Garside P. (2007) Images in cardiovascular medicine. Multiphoton microscopy for 3dimensional imaging of lymphocyte recruitment into apolipoprotein-E-deficient mouse carotid artery. Circulation 115, e326–8. 17. Celli S, Lemaitre F, Bousso P. (2007) Realtime manipulation of T cell-dendritic cell interactions in vivo reveals the importance of prolonged contacts for CD4+ T cell activations. Immunity 27, 625–34. 18. Galkina E, Kadl A, Sanders J, Varughese D, Sarembock IJ, Ley K. (2006) Lymphocyte recruitment into the aortic wall before and

during development of atherosclerosis is partially L-selectin dependent. J Exp Med 203, 1273–82. 19. Matheu MP, Cahalan MD (2007) Isolation of CD4+ T cells from Mouse Lymph Nodes using Miltenyi MACS Purification. J Vis Exp http://www.jove.com/index/ Details.stp?ID=409 20. Huang JH, Cárdenas-Navia LI, Caldwell CC, Plumb TJ, Radu CG, Rocha PN, Wilder T, Bromberg JS, Cronstein BN, Sitkovsky M, Dewhirst MW, Dustin ML. (2007) Requirements for T lymphocyte migration in explanted lymph nodes. J Immunol 178, 7747–55.

Chapter 14 Applying an Adaptive Watershed to the Tissue Cell Quantification During T-Cell Migration and Embryonic Development D. Zhu, S. Jarmin, A. Ribeiro, F. Prin, S.Q. Xie, K. Sullivan, J. Briscoe, A.P. Gould, Federica M. Marelli-Berg, and Y. Gu Abstract Cell and particle quantification is one of the frequently used techniques in biology and clinical study. Variations of cell/particle population and/or protein expression level can provide information on many biological processes. In this chapter, we propose an image-based automatic quantification approach that can be applied to images from both fluorescence and electron microscopy. The algorithm uses local maxima to identify labelling targets and uses watershed segmentation to define their boundaries. The method is able to provide information on size, intensity centroids and average intensity within the labelling partitions. Further developed from this method, we demonstrated its applications in four different research projects, including recruitment enumeration of circulating T cell in non-lymphoid tissues, cell clustering in the early development of the chick embryo, gold particle localization and clustering in electron microscopy, and registration/co-localization of transcription factors in neural tube development of early chick embryo. The advantages and limitations of the method are also discussed. Key words: Automatic cell quantification, image segmentation, fluorescence microscopy, electron microscopy, watershed segmentation.

1. Introduction Cell quantification is one of the techniques frequently used in biology and clinical haematology. Biomedical researchers rely on cell counts and fluorescence intensity to identify the changes in cell populations, protein expression, and level of aggregations and clustering. Traditionally, cell quantification relies on manual enumeration. Cells are incubated in special counting chambers F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_14, © Springer Science+Business Media, LLC 2010

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with grids or haemacytometers and then visualized and counted by technicians under bright-field microscopes. An experienced operator can provide further quantitative information about clusters and morphological variations of cell population. Manual counting is very flexible, inexpensive and less technically demanding and therefore is still the most widely used method in clinical pathology and haematology (1). However, manual counting is quite labour intense and tedious as a routine clinical practice, especially when hundreds of images need to be quantified for statistical significance. It also suffers from inconsistency, lack of reproducibility and inter-observer variation. Those difficulties have prompted the development of automatic cell quantification. Compared with manual counting, automatic quantification has advantages of fast, light work load and good repeatability (2). It also provides morphological information (size, brightness and cell/particle shape) that would otherwise be difficult to evaluate by manual enumeration. Recently, cost-effective alternatives have emerged (1, 3–5). Among them, the most widely used are imaging-based microscopic approaches. A typical set-up is mainly composed of a bright-field or fluorescence microscope, an image acquisition system (a digital camera, a computer and acquisition software) and corresponding cell quantification software. Such an approach inherits the advantages of manual counting and overcomes most of the shortcomings mentioned above. With only a moderate investment to upgrade a traditional cell inspection system, it has a vast potential in both biological research and clinical practice. The performance and the application field of imaging-based systems vary with their quantification approaches. For instance, cell segmentation is mainly achieved from 2D images, based on the information on intensity distribution, cell geometrical profile, cell size and sub-cellular structures. A typical intensity-based cell segmentation system relies on a global threshold to separate background and counting targets into binary pixel clusters, where those “white pixels” represent cell candidates (6, 7). The algorithm works well with a picture having uniform background, clear cell separation and high signal-to-noise ratio. Unfortunately, images seldom satisfy these conditions. A tissue image usually suffers from variations in cell expression level, uneven background, auto-fluorescence, cell overlapping and clustering (8). To deal with such “real” situations, more sophisticated approaches based on image local environment, gradients of intensity profile and intensity similarity between neighbouring pixels have been developed. Corresponding techniques are often referred to as adaptive thresholding, edge detection and region growing, respectively. An adaptive thresholding varies the threshold locally to overcome uneven background (9). Edge detection picks up the cell border from intensity gradients, which circumvents the

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problem of uneven brightness caused by different cell expression level (10). Region growing searches the neighbourhood pixels for similar grey levels and progressively follows the intensity profile for the cell region (11, 12). So far, many successful applications have been reported (13–16). However, it is accepted that the above techniques have limitations. Adaptive thresholding requires reasonable signal level. It may also have difficulty in finding a reasonable threshold when cells overlap to form a complex profile. Edge detection is sensitive to image details and is prone to noise level and background variation. Region growing requires seeds to start the process. A correct selection of enough seeds can be difficult in an automatic system. It is of great interest to develop automatic algorithms that are more robust to variations of cell type, cell density, microscope set-ups and image quality. Such an approach may require a combination of a number of techniques and employ comprehensive information including multiple fluorescence labelling, temporal expressional variation and the preknowledge of possible cell shapes (17–19) (see Note 1). Watershed segmentation is one of the region-growing approaches. It treats an image as a topographic surface where the grey levels of the image represent altitudes. This topographic analogy considers bright and dark image areas as hills and valleys of a landscape. During segmentation, the landscape is flooded with water flowing from high-altitude area into domains of low altitude where accumulating water forms lakes. With the water flowing in, those lakes expand and are eventually merged with adjacent lakes. The dividing lines at the merging moment are termed as watershed, while those lakes are termed as catchment basins. The merging process carries on until the water level reaches the highest peak of the landscape (brightest pixel of an image). In a cellular fluorescence image, the watersheds segment the topographic surface into catchment basins that correspond to individual cells. It is generally accepted that applying automatic watershed transformation to a biological image is limited by over-segmentation and irregular boundaries. Over-segmentation is usually caused by texture of sub-cellular details, while irregular boundaries are the results of image noise. Since watershed was implemented in 1991 (20), many efforts have been made to combat these problems. Algorithms that combine the transformation with pre-knowledge such as cell shape, texture and grey-level gradients have been developed to meet different applications (21). In this chapter, we proposed an adaptive watershed approach, which combines a local maximum detection algorithm with the grey-level-based watershed (referred to as local maximum seeded watershed or LMSW). Local maximum detection was used to identify labellings from the maxima of their intensity profile. Then these peak locations are seeded into the watershed to segment cell area from the background. Using the segmented image,

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information about intensity centroid, cell partition area and average fluorescence intensity are quantified. Detailed descriptions of the algorithm and four application examples are outlined thoroughly in Section 3. The advantages and shortcomings of LMSW are addressed in Section 4. As the following descriptions are applicable to both fluorescence-labelled cell and EM particle, we use “labelling target” to represent all counting subjects hereafter unless specified.

2. Materials 2.1. Tissue Sample Processing and Visualization in Memory T-Cell Migration

1. Generation and culture of HY (male antigen)-specific memory T cells has been described in detail elsewhere (22). The number of T cells required per animal is 3 × 106 –107. 2. IFN-γ (Peprotech EC, London, UK; 600 U/mouse): To be injected i.p. to induce local upregulation of MHC molecules and local antigen presentation 48 h prior to the adoptive transfer of T cells. 3. Labelling: 5 μM cell-linker PKH26 (Sigma, St Louis, MO, USA). 4. Sampling: Tissue samples were embedded in optimal cutting temperature (OCT) compound (Raymond A Lamb, Eastbourne, UK), snap-frozen and stored until analysis (23). 5. Peritoneal membranes or frozen tissue sections were laid onto Polysine-coated microscope slides (VWR International, Lutterworth, UK), left to dry and mounted in Vectorshield mounting medium with DAPI (Vector Laboratories, Peterborough, UK). 6. Imaging: Fluorescence images were taken by Zeiss Axiovert S100TV fluorescence microscope with MicroMax CCD camera (Princeton Instruments, USA). The objective used was Plan-Neofluar 10×/0.30 Ph1.

2.2. Preparation and Imaging of Embryonic Sections in Gene Expression Studies

1. In ovo electroporation has been described in detail elsewhere (24). 2. Neural tube samples are fixed in 4% paraformaldehyde. 3. DAPI staining: Used to highlight the nuclei of targeted cells were detected by indirect immunofluorescence. 4. Secondary antibodies for immunofluorescence: Hox and Alexa 594-conjugated secondary antibody (Molecular Probes). 5. Optical sections: Acquired from flat-mounted sample using a Leica TCS-SP1 confocal microscope with a PL Fluotar 40×/1.0 oil objective.

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6. LCS software (LCS Lite© , Leica Microsystems): For tissue analysis. 2.3. Sample Processing and Imaging of RNA Polymerase II

1. Cells: HeLa cells, including Cajal and promyelocytic leukaemia (PML) nuclear bodies (25). 2. Sample processing: Cells were fixed with 4% paraformaldehyde, embedded in sucrose, frozen, cryosectioned (120 nm thick), captured on drops of sucrose and transferred to formvar-coated nickel grids for immuno-gold labelling. 3. Antibodies: Specific mouse anti-Pol II antibodies and conjugated with 5-nm gold particles, while Cajal or PML nuclear bodies were labelled with rabbit anti-coilin or antiPML antibodies and conjugated with 10-nm gold particles (Molecular Probes), counterstained with uranyl acetate before EM imaging. 4. Imaging: The images were acquired on a JEOL-1011 TEM (JEOL UK Welwyn Garden City, Herts, UK) with a cooled slow-scan KeenView CCD (1392 × 1024 pixels; Soft Imaging System, Munster, Germany).

2.4. Preparation and Imaging of Embryonic Tissue for the Expression Analysis of Transcription Factors

1. Staging of chick embryos has been described elsewhere (26). 2. Preparation of tissue samples: Embryos were fixed for 1 h with 4% paraformaldehyde in 0.12 M phosphate buffer (PB) at pH 7.4. The embryos were cryoprotected in 30% sucrose in 0.12 M PB overnight at 4◦ C, embedded in 7.5% gelatin/15% sucrose in 0.12 M PB and stored at –80◦ C. 3. Sections: Obtain frozen 14 μm transverse sections of the embryos. 4. Antibodies: Rabbit α-Nkx2.2 1:4000 (27); guinea pig α-Olig2 1:8000 (28); and mouse α-Pax7 1:20 (DSHB) and visualized with secondary antibodies conjugated to FITC, Cy3 and Cy5 (Jackson Labs). Cell nuclei were visualized with DAPI. Imaging: Images were taken from a Leica TCS-SP2 AOBS confocal microscope using PL APO 40×/1.25 oil Ph3 objective.

3. Methods 3.1. Sample Preparation and Imaging 3.1.1. Imaging of Tissue-Infiltrating Memory T Cells

Based on a well-established model of lymphocyte trafficking (22), T cells (1 × 107 ) were injected intravenously in syngeneic female recipient mice to assess their constitutive recirculation in various non-lymphoid tissues. HY-specific T cells (3 × 106 ) were also employed to assess the infiltration of antigenic inflamed tissue (peritoneal membrane and cavity) following intraperitoneal

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injection in female or male recipients. As infiltration of the peritoneal membrane by specific T cells is dependent on major histocompatibility complex (MHC), molecule upregulation and antigen presentation, in these experiments, mice were treated with an i.p. injection of IFN-γ (Peprotech EC, London, UK; 600 U/ml) 48 h prior to the administration of HY-specific T cells. Injected cells were labelled with 5 μM cell-linker PKH26 (Sigma, St Louis, MO, USA). Tissue samples were harvested after 24 h and embedded in optimal cutting temperature (OCT) compound (Raymond A Lamb, Eastbourne, UK), snap-frozen and stored until analysis (23). Peritoneal membranes or frozen tissue sections were laid onto Polysine-coated microscope slides (VWR International, Lutterworth, UK), left to dry and mounted in Vectorshield mounting medium with DAPI (Vector Laboratories, Peterborough, UK). The fluorescence images were taken by Zeiss Axiovert S100TV fluorescence microscope with MicroMax CCD camera (Princeton Instruments, USA). 3.1.2. Analysis of Transcription Factors in Embryonic Tissue

Mouse Hox4 gene was misexpressed within the neural tube of a 2-day-old chick embryo using in ovo electroporation (24). After electroporation, the embryos were incubated for 48 h. Then, the neural tube was dissected and fixed in 4% paraformaldehyde. Nuclei of the neuroepithelial cells were visualized using DAPI staining and the nuclei of targeted cells were detected by indirect immunofluorescence using a primary antibody specified to Hox and Alexa 594 conjugated secondary antibody (Molecular Probes). Optical sections were acquired from flat-mounted sample using a Leica TCS-SP1 confocal microscope with a PL Fluotar 40×/1.0 oil objective. A depth average projection of eight sections equivalent to 7 μm in depth beneath the ventricular surface of the neuroepithelium was obtained using LCS software (LCS Lite© , Leica Microsystems) to enhance the signal-to-noise ratio.

3.1.3. TEM Imaging of RNA Polymerase II

The high-magnification EM images were taken from nuclei of HeLa cells, including Cajal and promyelocytic leukaemia (PML) nuclear bodies (25). Briefly, cultured cells were fixed with paraformaldehyde, embedded in sucrose, frozen, cryosectioned (120 nm thick), captured on drops of sucrose and transferred to formvar-coated nickel grids for immuno-gold labelling. The different forms of largest subunit RNA polymerase II were labelled with specific mouse anti-Pol II antibodies and conjugated with 5-nm gold particles, while Cajal or PML nuclear bodies were labelled with rabbit anti-coilin or anti-PML antibodies and conjugated with 10-nm gold particles. The labelled sections were counterstained with uranyl acetate before EM imaging. The images were acquired on a JEOL-1011 TEM (JEOL UK Welwyn Garden City, Herts, UK) with a cooled slow-scan KeenView CCD (1392 × 1024 pixels; Soft Imaging System, Munster, Germany).

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3.1.4. Preparation and Imaging of Embryonic Tissue for the Expression Analysis of Transcription Factors

Chick embryos were staged (26) and fixed for 1 h with 4% paraformaldehyde in 0.12 M phosphate buffer (PB) at pH 7.4. The embryos were cryoprotected in 30% sucrose in 0.12 M PB overnight at 4◦ C, embedded in 7.5% gelatin/15% sucrose in 0.12 M PB and stored at –80◦ C. Immunofluorescent staining was performed on frozen 14 μm transverse sections of the embryos with the following primary antibodies: rabbit α-Nkx2.2 1:4000 (27); guinea pig α-Olig2 1:8000 (28); and mouse α-Pax7 1:20 (DSHB) and visualized with secondary antibodies conjugated to FITC, Cy3 and Cy5 (Jackson Labs). Cell nuclei were labelled with DAPI. Images were taken from a Leica TCS-SP2 AOBS confocal microscope using PL APO 40×/1.25 oil Ph3 objective. For each image, three optical sections from 1 μm depth were taken and used to generate an averaged projection that produced the best signal-to-noise ratio.

3.2. Designing an Adaptive Watershed-Based Algorithm for Tissue Cell Quantification

The LMSW approach presented here is an intensity-based automatic quantification system. It relies on local maximum intensity to identify targets and use watershed to segment labelling area. The overall measurement consists of four steps, including preprocessing (optional), target identification, watershed segmentation and labelling quantification (Fig. 14.1). Unlike most watershed-based methods that rely on the segmentation result to identify targets, the approach developed in our lab separates identification and segmentation into two steps. Labelling targets are first identified by using local maximum intensity detection. Then they are employed as seeds for watershed segmentation. Since targets are identified first with great accuracy, the over-segmentation problem in classic watershed approaches has been greatly improved. This technique was originally developed to quantify single labelling systems, such as positive population, intensity centroids and expression level-related average fluorescence intensity. It has been further developed into the quantifications of multiple labelling, clustering, TEM particle enumeration and target registration. Experiments described in Section 2 are four examples of projects in which LMSW has been involved. These examples demonstrate its potentiality and flexibility as in every case the technique has been re-developed to meet application requirements (see Note 2 & Note 3).

3.2.1. General Description

3.2.2. Local Maximum Detection in Target Identification

On a digital image taken from a fluorescence sample, labelling targets resemble groups of glowing pixels of different size, shape, brightness, density and intensity profiles (Fig. 14.1). In each group, there is at least one pixel brighter than all others. This pixel is defined as the peak of its intensity profile, which usually has three distinctive features: (1) the pixel value is the maximum

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Fig. 14.1. LMSW procedures illustrated with an example: (a) LMSW processing flowchart; (b) fluorescence image candidate; (c) pre-processing with image smooth and thresholding; (d) target identification result (black marks correspond to the identified cells); (e) target segmentation result. Image size is 278 × 127 pixels. Scale bar in (b) is 5 μm. Microscopic images shown hereafter were not enhanced in any way apart from LMSW process. They were cropped from original image to maintain reasonable visibility.

among its neighbours or within local area, (2) spatially growing from this point to include more pixels, the area increases with the decrease in grey levels and (3) peaks from neighbouring groups are rarely connected with each other, although two groups would sometimes have certain overlap. These features enable us to use the peak as a countable feature. If each target has a single-peak intensity profile, the number of peaks represents the number of labelling targets. Detection of peaks is a grey-level searching approach involving the full dynamic range of an image (0–255 for 8-bit image).

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Fig. 14.2. Illustration of local maximum detection where x and y correspond to two spatial dimensions of an image and z corresponds to the fluorescence intensity level.

The overall dynamic range can be divided into a number of searching levels (16 levels in Fig. 14.2, shown in different grey levels for discrimination). At each level, a global threshold is set to discriminate all pixels. Pixels brighter than the threshold remain unchanged, while those dimmer are set to zero. Searching through the remaining pixels in each group, a local maximum can be located as the peak of the group. This process resembles water fluctuation in a reservoir with the thresholds corresponding to water levels and labelling targets being the hills submerged in the water. When the water recedes from the maximum level, the top parts of high hills start to emerge as islands. Low hills remain invisible until the water level decreases further. The overall landscape can be revealed only when the water is drained. With the searching level progressing from high to low (or water level receding), peaks with different maxima will emerge and be detected. While new peaks are being discovered, peaks detected from previous rounds will also be detected repetitively. As the locations and pixel values from the same peaks remain invariant, only those newly detected will be countable. By searching through the whole intensity range of an image, all targets with distinctive profiles (stiff or smooth, and bright or dim) may be identified. It is worth mentioning that using local maximum as labelling identity can potentially generate over-identification, especially when the image is noisy. A local maximum can be the peak of a true target, a speckle of image noise or texture of labelling details. The detection has to be combined with other

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information such as size and distance between peaks to retrieve the correct identification. In practice, a two-step selection approach was used. First, a size criterion (one-fifth of minimum labelling size) was used to examine the size of groups where new local maxima were detected. Only maxima from pixel groups with higher than criterion size are included. The examination effectively excludes noise-related speckles whose profiles are usually sharp and short with a small, slow-growing section area. Second, maxima are re-examined after the local maximum detection. Profiles of all peak candidates were sectioned at the full-width halfmaximum (FWHM) and projected onto a 2D plane. Their section sizes and intensity centroids were then calculated. Comprehensive information on section overlaps, distance between intensity centroids and correlation between locations were used to examine the neighbourhood and to exclude false identities.

3.2.3. Implementation of Watershed Segmentation

We implemented the original grey-level-based approach proposed by Vincent and Soille (20) because the identification result can be naturally used as seeds of segmentation. To start the process, images need to be grey level inverted before the segmentation so that peaks of intensity profiles become local minima or bottom of catchment basins, which are in turn seeded into the watershed process. Water starts to rise from those seeds and expands with the increase in water level. A watershed is built when neighbouring catchment basins merge with each other. Catchment basins that do not have seed points would not have watersheds built when they merged with others. In this way, the oversegmentations caused by texture of labelling details are greatly reduced.

3.2.4. Image Pre-processing

Unlike commercial cell-counting software, pre-processing is not always needed for LMSW quantification. However, if an image is “noisy”, an image smooth process (low-pass filtering) may help to reduce the quantification error. The smooth process is also needed to overcome the influence of texture of labelling details. In practice, smooth options (a Gaussian filter or a median filter with different kernel size) and a global thresholding are incorporated into the software. A special background subtraction is also available for TEM images with strong background from heavy metal counterstaining or from sub-cellular structures.

3.2.5. Algorithm Implementation

The algorithm was implemented in a Java environment (V1.6© , Sun Microsystems Inc.). While the identification, the segmentation and the quantification of labelling targets are done automatically, the pre-processing options and parameters can be adjusted interactively. Depending on image type and quality, an operator can decide the best approach with selections of image smooth,

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background subtraction and global thresholding (Fig. 14.1). Little prior knowledge of experiments and imaging set-up are required for running the program. Four parameters are usually required for the process, including maximum and minimum target widths in the image, background threshold and step width of local maximum search. A noisy image or an image showing rich texture of labelling details needs to be smoothed in advance. In this case, decisions on filter type (Gaussian and median) and kernel size are also required. We found that most of the parameters remain the same for images taken from the same experiment or experiments with the same set-up. Therefore, setting up parameters is required only for the first image. Results are presented as images with marked target locations of positive counts (Fig. 14.1d) and as a data spreadsheet, including intensity centroid (x, y), target partition size and average brightness (Fig. 14.1f) (see Note 4). In the near future, we will incorporate LMSW into Image J as a plug-in for free download. 3.2.6. Applications 3.2.6.1. Recruitment Enumeration of Circulating T Cell in Non-lymphoid Tissues

Productive T-cell immunity requires both the activation and the migration of specific T cell to the antigenic tissues. In recent studies, we have quantified the number of adoptively transferred T cells in a variety of tissues. In order to gain statistical significance, LMSW was employed to quantify the migration of fluorescently labelled T cells. During the experiment, we found that, with PKH26 labelling (PE) located mainly in the cytoplasm, the cell individuals were optically difficult to resolve. This is caused by the dense cell population which leads to plasma membrane overlap between cells (Fig. 14.3). In order to circumvent the problem, we employed DAPI-stained DNA as the counting subject, while using PKH26 labelling (PE) as a mask to include solely positive cells. During the process, the PE channel was globally thresholded to eliminate background. Then it was binarized with pixels brighter than the threshold set as “white” showing labelled cells as a positive population. This mask is applied to the DAPI-labelled counting channel to exclude nuclei of non-positive T cells from which the remaining T-cell population was enumerated. To estimate the enumeration accuracy, we use manual counting as a reference. Two experienced biologists were invited to count 83 images independently, with no pre-selection and preprocessing. The results from two operators were averaged to generate the “true count” for each image. The averaged absolute differences between two operators were used as errors of manual counting as they reflected inter-observer variation. For the total of 83 images, the average inter-observer variation was 1.81%. After the automatic identification, results were checked by the same team. All miscounts and overcounts were added in absolute values as the quantification error. Errors from two independent

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Fig. 14.3. Quantification of T-cell migrations in non-lymphoid tissues: (a) original PKH26 labelling channel; (b) original DNA-stained DAPI channel; (c) quantification results with white marks representing positive identification; and (d) data spreadsheet of positive locations. Original image size: 329 × 284 pixels. Scale bar in (a) is 5 μm.

Fig. 14.4. Comparison of cell identification between manual and LMSW approaches: (a) counting correlation; (b) relative counting errors of LMSW. The averaged relative error is 6.97% ± 2.66% for total of 83 images (88% of images achieved less than 10% error). The linear correlation between manual counting (X) and LMSW counting (Y) is Y =0.9608 × X + 3.8498.

operators were averaged and ratioed with the average manual counts to generate the relative error of each image (Fig. 14.4). The average difference between manual counting and LMSW counting is around 7% (see Note 5 & Note 6).

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During embryonic development, groups of cells become specialized to form individual organs. Cells within an organ tend to adhere to one another more strongly than they do to unrelated neighbouring cells from a different organ. Within the developing hindbrain, another type of differential adhesion leads to neural cells being segregated into segmental groups called rhombomeres. A family of homeodomain-containing transcription factors called Hox genes are known to be responsible for endowing neurons within each rhombomere with a distinct cell identity. As a part of a large project on Hox genes investigation (29), we were interested in testing whether experimentally induced expression of a Hox protein would be sufficient to make embryonic chick cells adhere to one another, thus forming large multi-cellular clusters. In order to quantify the cell aggregation, we developed a clustering module to measure the number and size of clusters based on the number of cells and their locations quantified from LMSW. When forming clusters, the cell density becomes spatially uneven. Inside each cluster, cells are close to each other, which is in contrast to the distinctive margin between clusters. We used closest neighbour criteria to sort cell populations so that cells with gaps smaller than a predefined distance are classified into the same group. The procedures are detailed as follows: 1. Mixtures of Hox-expressing and non-expressing cells were prepared and fixed according to Section 2.2. 2. Samples were imaged in depth-resolved sections on TCS SP1 confocal microscope. 3. Average projection was applied to bring eight sections into the same 2D image (Section 3.1.2). 4. Images were segmented and quantified using the LMSW approach (Fig. 14.5).

Fig. 14.5. Hox-induced cell clustering: (a) original image showing Hox-expressing cells mixed with non-expressing cells (unlabelled) and (b) clustering result. Original image size: 512 × 512 pixels. Scale bar in (a) is 20 μm.

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5. The distances between intensity centroids of all cells were calculated. 6. Cells were grouped together if the gaps between their intensity centroids were smaller than the predefined criteria. 7. The number of cells in each cluster was counted. 8. A cluster histogram vs. size of cluster was generated for each image. Combining the clustering histogram with other experimental results (unpublished data), we concluded that Hox-expressing cells form statistically larger clusters than do non-expressing cells. 3.2.6.3. Application to TEM Images

Similar to immunofluorescence labelling in optical microscopy, immuno-gold labelling is widely used in electron microscopy (EM) for ultra-structural studies of cellular antigens (25). Electron microscopists use different sizes of gold particles (such as 5, 10 and 20 nm) to distinguish different cellular components in sub-cellular compartments. The counts, aggregation and locations of certain size particles can reveal the presence or the colocalization of certain sub-cellular components. Quantification of gold particles has slightly different technical focuses. On the one hand, it does not have the implications caused by the texture of sub-cellular details or the difficulties from cell overlapping. On the other hand, since most EM cryosections are thin and nearly transparent to electron beams, heavy metal counterstaining (such as lead citrate and uranyl acetate) is used to scatter electrons and enhance contrast of structure differences. The enhancement complicates the counting as the background pattern is so strong that sometimes it is difficult to distinguish gold particles from the particle-like background (Fig. 14.6a). An effective background subtraction is required before particle identification to remove the background and enhance the particle signal. Inspired by Sanchez-Marin (30), we developed an approach which (1) applied a low-pass filter on the original image to generate its background image, (2) subtracted the background image from the original, (3) convolved the result with a mean filter kernel to smooth the very high-frequency background and speckles generated during the process, and (4) re-projected the output image to the full intensity scale for the best signal contrast. This approach worked effectively with particles because their size and shape similarities were reflected in the Fourier domain as relatively concentrated frequency band. In contrast, background of a TEM image is mainly in the lower frequency domain. If the filters are designed properly, the background subtraction can remove most of the background while maintaining the major frequency of particles (Fig. 14.6b). We used a mean filter with the kernel size equivalent to two times of the maximum particle size

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Fig. 14.6. Illustrations of particle quantification and clustering in a TEM image: (a) TEM image candidate; (b) result of background subtraction; (c) segmentation result; and (d) particle clustering result with dark polygons corresponding to clusters of 10-nm gold particles and white polygons corresponding to 5-nm gold particles. Original image size is 941 × 740 pixels. Scale bar in (a) is 0.1 μm.

(empirical) measured from the image. The kernel size for highfrequency smooth matches full-width half-maximal radius of the microscope’s theoretical point spread function that is defined as σ = 0.21λ/NA (31). With standard imaging set-ups recommended by microscope manufacturers, σ should be around 1 pixel. In practise, 3 × 3 pixels is the closest kernel size. We used this procedure to explore the presence of different phosphorylated forms of RNA polymerase II (Pol II) and nascent RNA in Cajal and promyelocytic leukaemia (PML) bodies. In a fluorescence image, Pol II appeared to be perfectly co-localized with Cajal and PML bodies, suggesting that these nuclei compartments may play a role in Pol II assembly and transcription activity (25). However, these nuclear compartments are too small to be resolved by an optical microscope, and the co-localization assumption needs to be further explored using higher resolution tools. Image shown in Fig. 14.6a was taken on a JEOL-1011 TEM. The sample was made into ultrathin cryosections (about 120 nm) to enhance the depth resolution. Gold particles (5 and 10 nm) were used to label Pol II and Cajal or PML bodies (Section 3.1.3). After the acquisition, the particles in the image were first background subtracted using a 25 × 25 kernel. Then LMSW was used to identify and segment particles. The size information from the segmentation was drawn into a his-

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togram where they were classified into two groups corresponding to 5 and 10 nm. They are clustered separately according to the near-neighbouring criteria described in Section 3.2.6.2 (Fig. 14.6d). The quantification provided high-resolution (25 nm) details about the number of Pol II at each site, the size of Cajal or PML compartments, the distance between Pol II clusters, and the extent of overlap between Pol II and Cajal or PML compartments. Quantifications over 91 images indicate that Pol II clusters did not significantly overlap with Cajal or PML compartments (Fig. 14.6d). The perfect co-localization observed on the fluorescence microscope was due to insufficient optical resolution (about 250 nm) and therefore could not provide appropriate evidence of Cajal and PML involvement in Pol II assembly and transcription activity. 3.2.6.4. Registration and Co-localization of Transcription Factors in Embryonic Development

The specification of neurons in the central nervous system depends on the correct patterning of progenitor cells. In ventral regions of the neural tube, the spatial organization of progenitor cells is controlled by the morphogen Sonic hedgehog (Shh). The long-range activity of Shh drives the differential expression of a set of transcription factors including Olig2, Nkx2.2 and Pax7 along the dorsoventral axis, thus delineating the progenitor domains that will generate distinct neuronal subtypes. This is a dynamic process, depending on both the concentration and the duration of Shh signalling (32), leading to the progressive generation of domains with more ventral identities and increasingly precise boundaries. Systematic characterization of the spatial and temporal changes in transcription factor expression, therefore, provides insight into the dynamics of neural tube development. LMSW was introduced to automatically quantify the levels of expression and spatial distribution of transcription factors with single-cell resolution in the chick neural tube. Each sample was stained with antibodies for three transcription factors and corresponding secondary antibodies labelled with different fluorophores to generate multi-channel images that had to be combined in a single plane. This could be achieved by directly overlapping the three channels if the channels were perfectly registered. However, centroids calculated from the labelling of transcription factors may not accurately describe the dynamics of cellbased patterning. The variation of centroids is caused not only by cell movement but also by the expression changes of transcription factors in each nucleus. Specifically, the differences in the expression levels and the distribution of transcription factors inside the nuclei may influence the location of the intensity centroid. The transcription factors that are co-expressed in the same nucleus also showed different patterns. When projected into the same plane, the expression of these transcription factors did not fully overlap,

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generating inconsistency in the choice of cell centroids. In order to correctly assign the nuclei position across channels, we need a reference channel, independent of the transcription factors and relatively stable during development. DNA labelling satisfies these conditions and effectively marks the entire nuclear region. We set up a four-channel system, with DAPI-stained DNA and three fluorophore-visualized transcription factors. The intensity centroids in the DAPI channel describe cell locations, while average intensities from transcription factor channels represent their relative expression levels. Specifically, after acquisition of multi-channel fluorescence images (Section 3.1.4), the positive nuclei in each channel were quantified using LMSW. Each transcription factor channel was then registered to the DAPI channel.

Fig. 14.7. Illustrations of multi-channel registration and co-localization: (a) original DNA channel; (b) original Nkx2.2 channel; (c) original Olig2 channel; (d) original Pax7 channel; (e) registration of positive expressed cells localized by black marks; and (f) projected segmentation result with highlights (in saturated white colour) corresponding to co-expressed cells. Original image size is 639 × 842 pixels. Scale bar in (d) is 20 μm.

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During the registration, intensity centroids from the two channels were compared and the pair with closest proximity was chosen. If the distance was smaller than the corresponding cell width, the pair was registered as from the same nucleus. Finally, nuclei that had non-zero average intensities in more than one transcription factor channel, but with the same intensity centroids, were identified as co-expressed cells. Figure 14.7a–d shows the four-channel images taken at HH stage 20 embryos stained with DNA, Nkx2.2, Olig2 and Pax7. Figure 14.7e illustrates the spatially registered positive nuclei (marked with black dots), with Pax7+ nuclei located dorsally, Olig2+ nuclei located in the middle and Nkx2.2+ nuclei located ventrally. Nuclei co-expressed Olig2 and Nkx2.2 were highlighted in Fig. 14.7f, marking the boundary between two domains of expression (see Note 7). As the project progresses, we hope to generate a spatiotemporal atlas of progenitor markers in the neural tube, using the quantified expression. This will allow us to document and analyse the dynamics and reliability of neural patterning in embryos.

4. Notes 1. Image-based automatic quantification of tissue or embryonic cells usually suffers from uneven expression level, unpredictable cell concentration, cell overlap and low image contrast. Automatic particle quantification from EM images requires tackling problems of strong/uneven background, significant signal variation and sometimes poor signal-tonoise ratio. We have developed an intensity-based approach to address the above difficulties. 2. The approach can be used for both fluorescence and EM images. It is designed to quantify the labelling targets with information of positive counts, target size, intensity centroid and average intensity. However, it is not able to measure the sub-cellular structure or its shape patterns. The examples presented in this chapter used DNA staining or gold particle labelling as the counting subject because of their relatively smooth intensity profile and less overlap with each other. The approach is not however limited to nuclear staining. We have used PKH26 labelling in the cytoplasm to directly analyse T-cell infiltrates (33). LMSW relies mainly on the intensity profile so that the analytical error is influenced by the signal pattern on an image rather than by the specific sub-cellular staining. Although the signal pattern is ultimately determined by the structure of a labelling target, its profile on an image is also related to the experimental

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design. For example, the texture can be minimized by optimizing microscopic settings such as objective and pixel binning. It is a common practice in clinical haematology to use low-magnification objectives in cell counting to maximize the field of view. Depending on the sub-cellular labelling, LMSW can achieve a good estimation when the size of a counting target is less than 300 square pixels. The LMSW approach is relatively robust with non-perfect images such as those with strong background or high level of noise. As with most available algorithms, quantification accuracy of LMSW is mainly influenced by images, although its tolerance to image quality is much wider. It is especially capable of handling images from thick embryo/tissue samples, showing out-of-focus, uneven background and weak signal. With the local maximum detection, it can deal with images where both bright and dim targets are present with variable size and profile, therefore covering the full dynamic range of intensity detection. 3. The algorithm is less capable of dealing with images with rich cellular details or multi-peak profiles. The error is associated with overcount during target identification, as each peak may be treated as a candidate. The issue is further complicated by the fact that a multi-peak profile has little difference to the profile caused by target overlap. A successful application needs to keep a balance between overcounting from labelling details and miscounting from overlapped individuals. 4. The quantification results provided by LMSW include number of labelling targets, intensity centroids, partition size and average brightness. Among them, measurements of number of labelling targets (target identification) and partition size (segmentation) determine the quantification accuracy. 5. The error of the target identification is mainly the contribution of overcounts and miscounts. Miscounts were caused by faint targets in a noisy image or under uneven background. Any attempt to remove the background will inevitably delete some background-like weak targets. Miscounts can also be caused by low-contrast borders of overlapping individuals. The complication of shape, expression level, extent of overlapping and the number of targets overlapped together can result in an unpredicted shape with a variety of combined intensity profiles. For instance, a dim target overlapped with a bright one may be overseen if its intensity profile is so flat that its peak has little difference with that of the overlapping boundary of its neighbour. 6. Overcounts happened mainly when a target shows multiple intensity peaks. It may also be caused by speckles or debris, whose intensity profiles are target-like. At the

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current stage, we introduced three major steps to tackle the problem. (1) During the pre-processing, labelling structures can be smoothed. (2) In each cycle of local maximum detection, the size of inspected pixel cluster is examined (Figs. 14.1 and 14.2). (3) A comprehensive examination is applied at the end of the target identification. With the above efforts, we have managed some “difficult” images and have achieved averagely better than 90% accuracy in T-cell migration project. However, as shown in Fig. 14.4b, the identification rate from difficult images is generally worse than those with smooth intensity profiles. 7. The segmentation error is mainly related to boundary identification. Under the influence of background noise, the boundary area becomes indistinguishable when the signal level is comparable with noise. Wrong segmentation also happens in overlapped cells when the sub-cellular texture is more distinctive than the true partition. As we are still tackling these problems, size measurement presented in LMSW has only statistical and relative meaning. Averaged labelling size in an image can be used to compare between samples from the same experiment or experiments with the same settings. Relative size can also be used to classify labelling groups if the number of labelling types is known in advance (Section 3.2.6.3). From our experiences, the average brightness derived from size measurement seems to have more biological significance, as it reflects the expression level of labelling targets.

5. Conclusions We have presented an image-based automatic cell/particle quantification approach, which can be used for the analysis of fluorescence and EM images. The algorithm uses local maximum for labelling identification and grey-level-based watershed for the target segmentation. The four examples demonstrated that LMSW can not only be used for cell/particle enumeration but also be adopted to deal with a wide range of comprehensive applications. References 1. Aune MW, Sandberg S. (2000) Automated counting of white and red blood cells in the cerebrospinal fluid. Clin Lab Hematol 22, 203–10. 2. Sims AJ, Bennett MK, Murray A. (2002) Comparison of semi-automated image

analysis and manual methods for tissue quantification in pancreatic carcinoma. Phys Med Biol 47, 1255–66. 3. Agarwal A, Sharma RK. (2007) Automation is the key to standardized semen analysis using the automated SQA-V

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Direct crossregulation between retinoic acid receptor band Hox genes during hindbrain segmentation. Development 132, 503–13. 30. Sanchez-Marin FJ. (1999) Automatic segmentation of contours of corneal cells. Comput Biol Med 29, 243–58. 31. Thomann D, Rines DR, Sorger PK, Danuser G. (2002) Automatic fluorescent tag detection in 3D with super-resolution: application to the analysis of chromosome movement. J Microsc 208, 49–64.

32. Dessaud E, Yang LL, Hill K, Cox B, Ulloa F, Ribeiro A, Mynett A, Novitch BG, Briscoe J. (2007) Interpretation of the sonic hedgehog morphogen gradient by a temporal adaptation mechanism. Nature 450, 717–21. 33. Jarmin SJ, David R, Ma L, Chai J, Dewchand H, Takesono A, Ridley AJ, Okkenhaug K, Marelli-Berg FM. (2008) T cell receptor-induced phosphoinositide-3-kinase p110δ activity is required for T cell localization to antigenic tissue in mice. J Clin Invest 118, 1154–64.

Section IV Monitoring T-Cell Migration in Human Diseases

Chapter 15 Identifying Homing Interactions in T-Cell Traffic in Human Disease Patricia F. Lalor, Stuart M. Curbishley, and David H. Adams Abstract Description of the molecular mechanisms which regulate the traffic of lymphocyte populations over recent years [for useful reviews see (1, 2)] has significantly enhanced our understanding of the processes underlying acquired immunity and also permitted the development of therapies targeted at specific leukocyte subpopulations. Such therapies are dependent upon a detailed knowledge of the molecular regulation of lymphocyte adhesion to and migration through endothelium in specific tissues. Whereas animal models have been central to understanding the underlying mechanisms, it is crucial to confirm and extend observations in man by using analysis of tissues and in vitro cell-based models. In this chapter, we discuss expertise developed in our laboratory for the isolation of specific lymphocyte and endothelial populations from explanted human liver tissue specimens. We then move on to provide specific examples of assays such as the Stamper–Woodruff assay, the transmigration assay and the tissue-specific endothelial static and flow-based adhesion assays, which can be used to interrogate the tissue-specific adhesion and migration of lymphocyte subsets. Although our own experience is with human liver tissue, the general principles apply to analysing any organ of interest. Key words: Lymphocyte, adhesion, migration, endothelium, liver, chemokine, inflammation.

1. Introduction The inappropriate accumulation of lymphocytes in inflammatory or autoimmune diseases leads to tissue damage as a consequence of three separate but related events: (1) increased leukocyte recruitment from the circulation; (2) the survival of recruited cells in tissue; and (3) the activation of damaging effector mechanisms by tissue-infiltrating cells. For many years, research was focused on what activates the lymphocyte and how such cells F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_15, © Springer Science+Business Media, LLC 2010

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cause tissue damage and relatively little was understood about how these cells were recruited into tissue but studies over the last 15 years have elucidated the molecular basis of lymphocyte homing and demonstrated how and why lymphocytes are recruited to particular tissues. These studies explain why T cells and B cells are distributed in particular compartments in lymph nodes and how they are brought together with antigen-presenting cells during an immune response. They have also shown that different signals recruit specific subsets of lymphocytes to particular tissues such as the mucosal system in the gut and the skin. This unravelling of the molecular basis of lymphocyte homing has revolutionised the pathogenesis of immune-mediated disorders and provided novel therapeutic targets, some of which are in clinical trials. The ability to manipulate cell populations at sites of inflammation and modify disease processes underpins experimental cell therapy. For instance, in vitro expanded antigen-specific T cells can be infused into immunodeficient patients to reconstitute beneficial anti-viral immune responses or into patients with cancer to boost antitumour immunity and anti-inflammatory regulatory T cells have potential to suppress inflammatory diseases (3, 4). The identification of the homing receptors that guide cells to target tissues allows their therapeutic targeting. Hence, monoclonal antibody therapies directed against the α4-integrin (5) and α4β7 integrin (6) are currently under investigation for treatment of inflammatory bowel disease. Recent evidence that the nature of the antigen-presenting cell and microenvironmental signals determine which homing receptors are expressed by B and T cells during antigen-driven activation shows how such homing receptors can be imprinted on specific lymphocyte subsets. Most studies have been done in mice but despite the difficulty in isolating dendritic cells from human tissues, we and others have succeeded in demonstrating that these principles also apply to humans using primary human tissue-derived cells (7, 8). Identification of specific immunopathogenic or immunoprotective populations of cells has increased the possibility of modulating the traffic of such cells as a therapeutic strategy (9, 4). However, in order for such therapies to be tested in vitro and in vivo, it is necessary to have robust mechanisms for isolating and phenotyping cells and for measuring and modulating their adhesion to organ-specific substrates. We will focus on our expertise modelling lymphocyte recruitment to the human liver to illustrate methods for measuring T-cell interactions in human disease. We begin by describing processes for isolation of T-cell and dendritic cell populations from human tissue and then describe adhesion assays based upon the use of tissue sections or hepatic endothelial populations which can be used to dissect adhesive and transmigratory mechanisms. Though these examples focus on one specific

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organ (i.e. the liver), all the techniques have the potential to be translated into other organ systems with minor modifications.

2. Materials 2.1. Isolation of Lymphocytes and Dendritic Cells from Human Liver Tissue

1. Tissue storage medium: Either unmodified DMEM or RPMI-1640 (both from Invitrogen, UK) depending on whether non-parenchymal cell populations (endothelium, biliary epithelium) or leukocytes, respectively, will be isolated from the tissue. 2. Culture medium: RPMI 1640 media (Invitrogen, UK) supplemented with 10% (v/v) heat-inactivated foetal calf serum (FCS, Invitrogen, UK) and 60 μg/ml benzylpenicillin, 100 μg/ml streptomycin and 2 mM L-glutamine (all Sigma, UK). 3. Wash buffer: RPMI containing 0.1% bovine serum albumin solution. 4. “Stomacher”, Seward 400C Laboratory Blender and blender bags (both from Seward Co, UK). 5. OptiprepTM (Axis-Shield): Sold as 60% iodixinal (w/v) solution. Diluted to 13.5% in RPMI-1640 for DC enrichment. 6. DC purification by lymphocyte depletion with anti-CD3 R Dynabeads and magnetic positive selection on CD11c R (Stemcell Technologies) accordexpression using EasySep ing to manufacturer’s instructions.

2.2. Stamper–Woodruff Adhesion Assay

1. Poly-L-lysine solution, 10% (v/v, in dH2 O). This solution can be stored at +4◦ C until required and re-used to coat microscope slides as necessary for up to a month. 2. Acetone. 3. Adhesion assay medium: RPMI or PBS containing physiological levels of Ca and Mg and 0.1% BSA. 4. Detection antibody: Anti-CD3 mAb (e.g. M7254 for lymphocytes; Dako, UK) or anti-CD31 (e.g. JC70A for EC; Dako, UK). Both are used at pre-determined optimal concentrations ranging from 1 to 10 μg/ml and should be raised in the same species and of the same isotype to permit detection of both with the same secondary reagents. 5. Tris buffer: 0.05 M Tris-buffered saline, pH of 7.5 or 8.2 depending upon the stage of staining procedure. 6. Secondary and tertiary antibodies for APAAP staining: Rabbit anti-mouse antibody (Z0259: 1/25 dilution of manufacturer’s stock; Dako, UK), mouse APAAP (D0651: 1/50 dilution of manufacturer’s stock; Dako, UK).

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7. Chromogenic visualisation system, Fast Red Substrate (KO699; Dako, UK). 8. Water-soluble mountant: Immunomount (Shandon, UK). 2.3. Endothelial Isolation from Human Liver Tissue

1. Collagenase solution: 2 mg/ml type 1A collagenase (Sigma, UK) aliquotted into volumes sufficient for 1 cell prep and stored at –20◦ C until required. 2. Nycodenz solution: 25% (w/v) Nycodenz (Axis-Shield, UK) can be stored at +4◦ C ready for use. 3. Mouse anti-human HEA125 monoclonal antibody (PROGEN Biotechnik, Heidelberg, Germany) used at 10 μg/ml. R 4. Sheep anti-mouse Dynabeads (Invitrogen, UK) – beads are added at an optimal bead-to-target cell ratio according to manufacturer’s instructions.

5. Mouse anti-human CD31 monoclonal antibody (BBA7, R+D systems, UK) used at 10 μg/ml. 6. Complete endothelial culture medium: Human endothelialSFM (Invitrogen, UK) supplemented with 10% (v/v) heatinactivated human serum (HIHS, HD Supplies, UK), 60 μg/ml benzylpenicillin, 100 μg/ml streptomycin and 2 mM L-glutamine (all from Sigma, UK) and 10 ng/ml of vascular endothelial growth factor (VEGF) and hepatocyte growth factor (HGF, both from Peprotech, UK). 7. Trypsin solution: Tryp-LE (Invitrogen, UK) reagent is supplied at working concentration and can be stored at +4◦ C ready for use. 8. Collagen solution: Ready-coated tissue culture plasticware (e.g. Biocoat collagen I-coated flasks, BD-Biosciences, UK). 2.4. Adhesion and Transmigration Assay Systems

1. Microslide capillary tubes (VD/3530-050: Fisher Scientific, UK) have excellent optical qualities and defined internal dimensions (300 μM × 4 mm × 5 cm, see Fig. 15.1). The slides are prewashed with nitric acid and coated with APES (Sigma, UK) according to standard protocols prior to use to maximise attachment of cells and proteins to the glass. We coat the slides with collagen solution (see above) prior to loading with cells. 2. VCAM-1 solution: A stock solution of purified recombinant human VCAM-1 (R+D Systems, UK) is prepared according to manufacturer’s instructions (500 μg/ml–1 mg/ml) and aliquotted for storage at –20◦ C. Note that recombinant proteins have a limited shelf life and should not be subjected to repeated freeze–thaw cycles. We typically use final working concentrations of 1–50 μg/ml in PBS supplemented with 0.1% BSA.

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Fig. 15.1. Flow-based adhesion assay system used for measurement of lymphocyte adhesion and transmigration across endothelial monolayers. (a and b) Commercially available microcapillary tubes with excellent optical qualities are seeded with a confluent monolayer of endothelium. Cells are usually pre-stimulated with proinflammatory cytokines prior to assay. Capillary tubes are then connected into a flow-assay system (c) comprising a phase contrast, an inverted microscope, a syringe pump, a digital recording system and an electronic valve (to permit transition between perfusion of wash buffer and cells). The system is maintained at 37◦ C throughout the experiment. Adhesive events are recorded in real-time for offline analysis.

3. Cytokine solutions: We typically use human TNF-α or IFN-γ (10–100 ng/ml) in complete endothelial media (Peprotech, UK). 4. CellTracker Green solution (C2925, Molecular Probes, UK): A stock solution is prepared (1 mg/ml), aliquotted and stored at –20◦ C for up to 6 months. 5. Cell culture inserts: We have used inserts from the following manufacturers: BD Biosciences, Greiner, Corning.

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6. Collagen solution: We use an in-house preparation of collagen but it is possible to buy cell culture inserts treated with matrix components (e.g. Biocoat collagen I-coated inserts, BD Biosciences, UK) or to preincubate inserts overnight with recombinant matrix components (e.g. Fibronectin F2006, Sigma, UK, 10 ng/ml–10 μg/ml assay-dependent concentration). 7. Chemoattractant solution: We typically use purified recombinant human chemokines (e.g. CXCL12, 10–500 ng/ml; Peprotech, UK). A stock solution is prepared (100 μg/ml), aliquotted and stored at –20◦ C for up to 6 months. Freezethawing should be avoided.

3. Methods 3.1. Lymphocyte and Dendritic Cell Isolation from Human Tissue

3.1.1. Sample Collection and Preparation

In order to dissect the mechanisms by which populations of lymphocytes are recruited to and retained within tissue microenvironments, it is necessary to isolate pure populations of lymphocytes to study. The study of populations of cells purified from blood can be informative but analysis of tissue-resident populations yields more specific information about the signals required for recruitment to sites of inflammation (10, 11). We have developed methods to isolate lymphocytes from human liver tissue, which avoid the use of proteolytic enzymes (12) that alter cell surface receptors (13). We have used a similar approach to isolate cells from the human spleen. 1. Slices of human liver (150–200 g) are removed under sterile conditions as soon as possible after explantation and stored in tissue storage medium in a sterile beaker at 4◦ C until required. 2. To aid mechanical digestion, tissue slices are placed on a sterile Petri dish and finely chopped using scalpels (∼5 mm3 , Fig. 15.2a), with repeated washes in an excess of PBS used to remove contaminating blood. Once free from blood, the sample is resuspended in a volume of 200 ml in RPMI. 3. The tissue suspension is transferred into a “Stomacher bag” and sealed using a heated bag sealer. For extra security, this bag can then be sealed into an additional bag to prevent leakage. 4. The tissue sample is placed into the Stomacher compartment (sealed edge uppermost) and the front panel closed (Fig. 15.2c). The Stomacher is run at 260 rpm for approximately 6 min to homogenise the tissue (see Note 1).

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Fig. 15.2. Isolation of lymphocytes from human liver specimens. Liver tissue is finely diced under sterile conditions (a) and mechanically digested in a “Stomacher” laboratory blender (c). The resultant cell homogenate is passed through a sterile sieve (b) to remove undigested tissue prior to cell purification.

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3.1.2. Cell Isolation from Liver Homogenate

1. The cell homogenate is passed through a fine nylon mesh (50 μm) to remove undigested clumps and fibrous material (Fig. 15.2b). The cells are successively washed in buffer and centrifuged (650×g; 5 min) until the supernatant is clear and the total volume reduced to 50 ml. 2. The cell suspension is then layered over a Lympholyte-H gradient (Cedarlane Laboratories, Canada) and centrifuged at 600×g for 30 min without brake. The lymphocyte layer that forms at the interface between the gradients is removed and washed twice in wash buffer (650×g; 5 min). The purified lymphocytes may then be cryopreserved immediately or used in functional assays. Typically, we obtain yields of 20– 50 × 106 cells but this varies depending on the nature of the starting tissue (see Note 1). 3. Viability is checked using trypan blue exclusion before use.

3.1.3. Isolation of Dendritic Cells

1. From liver homogenate: Following lymphocyte isolation as described above, dendritic cells can be enriched by a further density-gradient step (see Note 2). The cells are resuspended to 25 ml, and 5 ml is layered onto 10 ml of Optiprep solution (13.5%, v/v) before centrifuging at 650×g for 30 min. The cells at the interface contain the dendritic cell fraction, with lymphocytes forming the cell pellet. Both cell types are washed twice in buffer (650×g; 5 min) before further use. Typically, we obtain 2–8 × 106 cells. 2. From whole liver tissue: Thin slices (1 mm) are taken from explanted liver under sterile conditions and transferred to 75-cm2 tissue culture flasks in 20 ml culture media (approximately 15–20 slices 2 cm × 2 cm × 0.1 cm). Sufficient liver is prepared to add to 6–10 tissue culture flasks. The tissue slices are incubated at 37◦ C for 24 h to allow the dendritic cells to “walk out”. All non-adherent cells are then retrieved by passing the supernatant through a fine nylon mesh (50 μm) to remove tissue pieces before centrifuging at 650×g for 5 min to pellet cells. Dendritic cells are then enriched over an Optiprep gradient as described above. 3. From skin: Full-thickness skin pieces are floated on culture media and cultured overnight in the same way as the liver DCs. Non-adherent cells that migrate out of the tissue into the media are collected and enriched as described for liver tissue. 4. From lymph node: Lymph node is first transferred to a sterile Petri dish and finely chopped using a sterile scalpel before being passed through a 50-μm mesh and the supernatant collected. Any tissue remaining in the nylon mesh can be “forced” through using the plunger of a sterile syringe. The

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resulting supernatant is then collected and washed in buffer (650×g; 5 min) and enrichment of dendritic cells achieved using a 13.5% Optiprep gradient as detailed above. 3.1.4. Purification of Dendritic Cells

Should dendritic cell purity be critical, further enrichment of any of the samples described above can be achieved by positive or negative selection: 1. Positive selection of CD11c+ dendritic cells. Dendritic cells should be resuspended to not more than 10 × 106 per millilitre in buffer (at least 100 μl), labelled with anti-human CD11c-PE mAb (clone B-ly6; BD Biosciences, UK) and incubated for 15 min at room temperature with regular agitation. Cells are then washed once in buffer (650×g; 5 min) before being resuspended to not more than 10 × 106 per R millilitre in buffer (at least 100 μl). Anti-PE EasySep beads are then added according to manufacturer’s instructions and incubated for 15 min at room temperature with regular agitation. Positive magnetic selection is then achieved according to manufacturer’s instructions. 2. Negative selection by depletion of CD3 lymphocytes. Contaminating lymphocytes can be removed by resuspending enriched DCs to not more than 10 × 106 per millilitre in R buffer (at least 1 ml) before adding anti-CD3 Dynabeads (Invitrogen, UK) and incubating for 15 min at room temperature with frequent agitation. Magnetic separation is then performed according to manufacturer’s instructions and the enriched supernatant collected.

3.1.5. Downstream Use of Dendritic Cells

1. Recent evidence shows that dendritic cells are responsible for imprinting tissue-specific homing receptors on responding lymphocytes during antigen presentation and activation (7, 14). In order to study tissue-specific lymphocyte imprinting by dendritic cells, purified DCs are first irradiated to prevent proliferation of any contaminating T cells. Allogeneic naive T cells are then mixed with prepared DC at 1:10–1:100 DC:T cell. The mixed leukocyte reaction is performed according to standard protocols for 5 days following which the cells can be removed for phenotyping or alternative functional assays.

3.2. Stamper–Woodruff Adhesion Assay for Investigating T-Cell Binding to Ex Vivo Tissue

The Stamper–Woodruff assay was designed to investigate lymphocyte adhesion to blood vessels in lymph nodes (15). We have developed it to allow us to study how leukocytes bind to endothelial cells in human tissue sections. Although the assay is simple and does not take into account the effects of physiological blood flow, it permits the user to quantify adhesion events on different

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vascular structures within tissues using small numbers of lymphocytes (16, 17). It has the advantage of using vessels in their native state in tissue as an adhesive substrate, thereby avoiding problems associated with isolation and culture of endothelial cells, and it permits comparisons between “normal” and “injured” tissues. In can also be used to study binding to other structures as well as blood vessels. For example, we used the assay to investigate differences between lymphocyte binding to tumour stroma and peripheral, uninvolved liver in patients with liver tumours (18) (Fig. 15.3).

Fig. 15.3. Analysis of lymphocyte binding to tissue sections using the Stamper–Woodruff assay. (a) Adhesion of lymphocytes (some indicated with arrows on panel) to labelled vascular structures (pink staining) within a frozen section of liver tissue. (b) Adhesion to a sequential section from the same liver specimen is reduced in the presence of a functionblocking antibody (compare a with b). Adhesion of lymphocytes to hepatic tumour sections and surrounding uninvolved tissue (c and d) can be quantified using the same methodology.

3.2.1. Preparation of Tissue Specimens as Adhesive Substrates

1. Cubes of human liver (typically 1–2 cm3 ) are cut under sterile conditions as soon as possible after resection or explantation and snap-frozen by immersion in liquid nitrogen for storage at –80◦ C until required (see Note 3). 2. Samples for adhesion assay are prepared by cutting 10 μm sections with a cryostat (see Note 3) and mounting them on slides pre-coated with poly-L-lysine. Care should be taken to ensure that the tissue blocks remain adequately frozen throughout processing.

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3. Slides are left to air-dry for 1 h prior to fixing by immersion in acetone for 5 min. Once fixed, the slides should immediately be wrapped in aluminium foil (two slides, back to back per foil wrap) and stored at –20◦ C until required. 3.2.2. Preparation of Lymphocytes for Adhesion Assay

1. Tissue-derived or peripheral blood lymphocytes are resuspended to 1 × 106 cells/ml in adhesion assay media (see Note 4). 2. To inhibit specific adhesion receptors, cells are pretreated with optimal concentrations of function-blocking antibodies in wash buffer for 30 min prior to removal of unbound antibody by washing. 3. Visualisation of adherent cells is facilitated by pre-labelling with a detection antibody which can be localised using APAAP labelling at the end of the assay procedure. This step can also be used to study specific subpopulations of T cells without the need for additional purification steps. Labelling is carried out as in Step 2 above (see Note 5).

3.2.3. Adhesion Assay

1. Sections are warmed to room temperature whilst still wrapped. Once thawed, the section is outlined using a wax pen to retain liquid and pre-wetted with TBS. 2. If required, sections can be pretreated with pre-determined optimal concentrations of function-blocking antibodies in wash buffer for 30 min prior to removal of unbound antibody by washing. 3. Visualisation of target endothelial cells (or other cells of interest) is facilitated by pre-labelling with a cell-type-specific antibody detected using APAAP labelling at the end of the assay procedure. Labelling is carried out as in Step 2 above. 4. Excess wash buffer is blotted from the section and cell suspension added (typically 100 μl, see Note 4). The section is incubated at room temperature for 30 min to allow lymphocytes to adhere. 5. The sections are then carefully washed using wash buffer to remove unbound cells and fixed by immersion in acetone.

3.2.4. Visualisation of Adherent Cells and Endothelium and Quantification of Adhesion

1. Labelled lymphocytes and endothelium are detected using a standard APAAP protocol. Add 100 μl of rabbit anti-mouse antibody solution and incubate for 45 min at room temperature in a humidified chamber. 2. Wash the sections with an excess of TBS and add 100 μl of APAAP and incubate for 45 min at room temperature in a humidified chamber.

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3. Wash sections, blot excess liquid and add sufficient freshly made Fast Red Substrate to completely cover the section (typically 100–250 μl). Incubate until colour develops (typically 10 min) and wash sections with distilled water to cease reaction. 4. Sections are counterstained using haematoxylin and mounted in water-soluble mountant. 5. Adherent lymphocytes (stained pink) binding to structures of interest can be differentiated from tissue-resident cells within the section using light microscopy (Fig. 15.3). Binding to at least 10 similar structures or uniformly sized areas of interest is determined for each section and data expressed as absolute numbers of cells/unit area/structure or as a percentage of binding in the absence of blocking antibodies. 3.3. Tissue-Specific Endothelial Adhesion Assays: Endothelial Culture

Endothelial cells from different organs and even different branches of the vascular tree in the same organ (e.g. lymphatic vs. capillary) have distinct phenotypes (19, 20), which determine their ability to recruit leukocyte populations. Thus it is important to model the endothelial environment of the particular tissue you are most interested in rather than relying on “generic” endothelial cells such as human umbilical vein endothelial cells. We have developed a method to isolate a specific endothelial population, hepatic sinusoidal endothelial cells, from human liver tissue (21). These cells reside within the hepatic sinusoids which constitute the specialised “capillary” bed of the liver, in close association with underlying hepatocytes. Sinusoidal endothelial cells have a unique phenotype and diverse functions (22, 23) and are the main portal of entry for inflammatory cells entering the liver (24). Our protocol for isolating these cells is detailed below.

3.3.1. Sample Collection and Preparation

1. Slices of human liver (50–100 g) are collected into DMEM as described above for the lymphocyte isolation prep. Slices should be used within 24 h of tissue collection (see Note 6). The tissue is mechanically chopped (see Note 6) into small pieces and washed free of contaminating blood as described above. 2. The tissue chunks are digested in 25 ml of collagenase solution in a covered glass beaker at 37◦ C for up to 45 min (see Note 6). 3. After digestion, 5 ml of FCS is added to inactivate the collagenase, up to 50 ml of PBS is added to dilute the sample and the digest is passed through a sterile nylon mesh (Fig. 15.2b) to remove lumps and undigested tissue.

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1. Hepatocytes are removed by density-gradient centrifugation in which the cell mixture is gently overlaid onto Nycodenz and centrifuged at 450×g for 30 min. The non-parenchymal cell fraction which floats on the density material is collected and washed twice in PBS (see Note 7). 2. Endothelial cells are purified from this cell suspension using immunomagnetic selection. Contaminating biliary epithelial cells (BEC) are removed by incubation with HEA125 mAb for 45 min at 37◦ C, washed with PBS to remove unbound antibody and incubated with secondary antibody conjugated R to magnetic beads (sheep anti-mouse Dynabeads ) at 4◦ C for 30 min (see Note 8) to allow magnetic removal. 3. The remaining cells are washed with PBS and HSEC selected using an antibody against CD31 before resuspension in complete endothelial culture media in collagencoated 25-cm2 tissue culture flask in a humidified incubator. 4. Cell yields vary considerably depending upon the nature and size of the tissue sample. It may take up to 3 weeks for cells to become confluent after which they should be divided 1:3 using trypsin according to standard protocols. 5. If required, cells can be frozen in freezing media (1 × 75-cm2 flask of cells in 1.5 ml of media to make 3 × 500 μl aliquots) for long-term storage in liquid nitrogen.

3.4. Tissue-Specific Endothelial Adhesion Assays: Adhesion and Transmigration Assay Systems

3.4.1. Flow-Based Adhesion Assay

Once cultures of endothelial cells have been established, they can be used to measure adhesive interactions with T lymphocytes. Flow-based adhesion assay systems represent the most faithful model of adhesive interactions under conditions recapitulating the blood flow in a specific vascular bed (25, 26). These assays are complex to perform and require considerable investment in equipment and expertise, but are nevertheless briefly discussed below. However, a useful compromise is the use of “static” adhesion assays and transmigration assays through commercially available cell culture inserts. Here, users have the flexibility of choice of adhesive substrate (e.g. purified recombinant adhesion molecules or endothelial cells), proadhesive stimuli (e.g. cytokines such as TNF-α or IFN-γ) (10) and inhibitory reagents and some systems permit collection and phenotyping of transmigrated lymphocyte populations (27) (see below). Two of these types of assay are described below. 1. Lymphocytes are prepared as above and resuspended in adhesion assay media (0.5–1 × 106 cells/ml). 2. Monolayers of hepatic endothelium are cultured to confluence in commercially available, collagen-coated microcapillary tubes (Fisher Scientific, UK) (see Note 9 and

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Fig. 15.1a, b). Endothelial monolayers are pre-stimulated with proinflammatory cytokines such as TNF-α or IFN-γ for 4–24 h prior to performing the assay (see Note 10). 3. Microslides are connected to the flow system (Fig. 15.1c) and wash buffer perfused through the slide at physiological shear stress (0.05–0.2 Pa depending upon application) to ensure that debris is removed and the slide is perfusing properly. Known concentrations of lymphocytes (typically 1 × 106 per millilitre, see Note 11) are then perfused through the slide for a defined time and phase-contrast microscopic images of adhesive interactions recorded in real time for offline analysis. Both the leukocytes and the endothelium can be treated with function-blocking reagents prior to the assay. 4. Data are analysed by determining the number of adhesive events per unit area of endothelium and normalised per million cells perfused (Note 11). Use of phase-contrast microscopy permits classification of adhesion events as “rolling adhesion”, “static adhesion” and “transmigrated cells” (similar to morphology demonstrated in Fig. 15.4), which can be expressed as a proportion of total adhesion events. 3.4.2. Static Adhesion Assay

1. Lymphocytes should be prepared according to the protocol above and resuspended in adhesion assay media (0.5–1 × 106 cells/ml is ideal). 2. Adhesive substrates can be prepared according to specific user requirements. We typically coat the wells overnight with recombinant adhesion proteins such as VCAM-1 (Fig. 15.4a) or use confluent monolayers of endothelial cells grown in 24-well plates (Fig. 15.4b). If required, endothelial monolayers can be pre-stimulated with proinflammatory cytokines such as TNF-α or IFN-γ for 4–24 h, and cells may be fixed with methanol and stored at 4◦ C in PBS prior to the assay (see Note 12). 3. The actual adhesion assay procedure is similar to that described for Stamper–Woodruff assays above. Five hundred microlitres of lymphocyte suspension is applied to the well containing adhesion substrate (in the presence or absence of blocking reagents) and incubated for 30 min to 1 h. The wells are washed carefully to remove non-adherent cells and adherent cells fixed using methanol. Fixed wells can be stored in PBS at 4◦ C for up to 2 weeks prior to analysis. 4. Lymphocyte adhesion can be scored in several ways. The easiest method is to count adherent cells in uniform fields of view. This method also permits observation of cell

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Fig. 15.4. Analysis of lymphocyte binding using a “static adhesion assay”. (a) Adhesion of peripheral blood lymphocytes to recombinant VCAM-1 immobilised in a tissue culture plate. Adherent cells can be subclassified as adherent but unactivated (a), adherent and activated/shape changed (b) or adherent and migratory (c). Similarly, adhesion assays can be performed on endothelial monolayers (b) using the same methodology. These assays can be automated by using fluorescently labelled lymphocytes which can be quantified using a fluorescent plate reader. Adhesion of cells in test wells can be compared to numbers in a calibration curve. Panel (c) shows a typical calibration curve from such an experiment and demonstrates the correlation between cell number and fluorescence.

morphology (see Fig. 15.4) and thus quantification of activated or migratory cells. More rapid, automated cell counts can be determined using fluorescent plate readers (e.g. Synergy-HT, Wolf Labs, UK) in conjunction with fluorescently labelled leukocytes.

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5. For fluorescent labelling, we use CellTracker Green solution. Cells are labelled in RPMI/10% FCS at 2.5 μM for 40 min, washed twice in PBS, reconstituted to 10 ml in RPMI/10% FCS and rested for 30 min at 37◦ C to allow incorporation of the dye. Following a further wash in sterile PBS, lymphocytes are diluted in adhesion assay media (see Note 13) and used from Step 3 above. Final cell counts in each well are calculated by comparison with a calibration curve of known cell numbers set up in an additional 24-well plate (see Fig. 15.4c). 3.4.3. T-Cell Chemotaxis and Transmigration Assays Using Cell Culture Inserts

1. Lymphocytes are prepared as described above. 2. Adhesive substrates within the culture inserts (Fig. 15.5) can be prepared in a variety of ways. We typically use either confluent monolayers of endothelial cells (see Note 14) or coat the inserts with extracellular matrix components such as collagen or fibronectin to improve adhesion. If required, endothelial monolayers can be pre-stimulated with proinflammatory cytokines as described above. 3. Lymphocytes or endothelial cells can be pretreated with pre-determined optimal concentrations of function-blocking antibodies in wash buffer for 30 min prior to removal of unbound antibody by washing.

Fig. 15.5. Measurement of lymphocyte transmigration using commercially available cell culture inserts. (a) Typical inserts contain a porous membrane (1) contained within a plastic housing (2 ) which sits inside a well of a tissue culture plate. (b) A lymphocyte suspension is placed in the upper chamber (1) above the porous membrane (2 ) which separates the cells from a chemoattractant solution placed in the bottom well (3 ).

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4. If required a chemoattractant can be placed in the bottom chamber to promote migration. We typically use either recombinant chemokines (e.g. CXCL12) or a supernatant collected from stimulated cultured cells (see Note 15). 5. Migration is analysed by placing the cell culture insert (with or without endothelial monolayer or matrix coating) into a well in a tissue culture plate. The bottom well contains either media alone or chemoattractant solution. A known number of lymphocytes (typically 500 μl at 0.5–1 × 106 cells/ml) is placed into the upper chamber. The plate is incubated at 37◦ C for up to 4 h (see Note 16) to allow migration through the porous membrane into the bottom chamber. 6. Quantification of cell migration can be assessed in a variety of ways. The simplest method involves performing a cell count for cells remaining in the top chamber and present in the bottom chamber and comparing this to the starting number of cells applied. However, additional information can be obtained by using a flow cytometer to perform cell counts in combination with fluorescent labelling of key cell populations (for example, CD4 or CD8+, CD45RO or RA+) to yield information not only on total numbers of cells migrating but also on what populations of cells actually migrate (27).

4. Notes 1. Although this procedure is reasonably efficient, the yield of lymphocytes from each sample of liver varies considerably and is dependent upon the degree of inflammation present (i.e. the starting number of inflammatory cells present within the sample). Typically, we get yields of 20–50 × 106 cells. Recovery of lymphocytes from very fatty specimens is problematic and care should be taken to remove as much fat as possible by thorough washing before and after stomaching. It is possible to adjust the duration of homogenisation in the Stomacher to improve tissue dissociation but our experience suggests that altering the “speed” is not beneficial. If visual observation suggests the persistent presence of contaminating cells such as hepatocytes and red blood cells after gradient centrifugation, the lymphocyterich suspension can be further purified using an immunomagnetic selection technique. 2. When isolating dendritic cells, consideration should be given to the subgroup of DCs required. Isolation via

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mechanical dissociation (Stomacher method) will yield populations of both plasmacytoid and myeloid DCs. However, isolation via the “walk-out” method yields only myeloid DCs. Commercially available immunomagnetic isolation kit can be used to further subtype isolated cells should this be required. 3. Due to the repeated manipulations involved in performing an adhesion assay and staining of tissue sections, it is necessary to maintain tissue integrity throughout the preparation and assay. Rough handling of the sample can cause the tissue to detach from the microscope slide. We ensure that “chunks” of tissue are rapidly frozen after processing by immersion in liquid nitrogen and immediately transferred to –80◦ C storage. Liquid nitrogen is also used to transfer the tissue to the cryostat. Adequate air-drying of the cut section at room temperature ensures good adherence to the poly-L-lysine-coated slides and once wrapped and fixed, sections should be stored at –20◦ C. Upon thawing, once sections have been pre-wetted, they should not be allowed to dry out at any point. We have found that cutting sections of 10 μM thickness ensures that they withstand processing. 4. We find 1 × 106 cells/ml to be ideal to ensure satisfactory distribution of lymphocytes over the whole surface of the tissue during the assay and to minimise clumping of cells. This number may need to be modified for other cell populations according to user requirements. Users should ensure that physiological calcium and magnesium levels are maintained in their assay media to facilitate maximal adhesion. Presence of high concentrations of serum or albumin during the assay may promote cell clumping and cause formation of protein aggregates on the section which make subsequent visualisation difficult. We have also noted that freshly isolated cells or “activated” cell lines cultured in mitogens such as IL-2 may be very “sticky”, so it may be necessary to rest cells overnight to minimise non-specific adhesion and clumping. 5. If the antibodies used to detect endothelial cells and adherent lymphocyte subpopulations are raised in the same species, both can be detected in a single APAAP procedure. Optimal concentrations should be determined and it is important to make sure that the antibodies used to define cells/tissue structures are not function blocking. 6. Because we isolate cells from donors with a variety of different liver diseases, we find considerable variation in the yield of endothelial cells (22) and some preparations may fail to yield any usable cells. This may be because

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the tissue was damaged or necrotic or had not been maintained under stringent aseptic conditions or stored incorrectly prior to preparation. The likelihood of success is increased (i) by ensuring the shortest possible duration of tissue storage prior to cell isolation, (ii) by ensuring that tissue is stored under sterile conditions in the correct media and (iii) by using as large a piece of tissue as possible to work with. Cirrhotic/fibrotic tissues tend to be the most difficult to isolate cells from. This is because more digestion time is required to breakdown mature fibrous tissue and it is often best to compromise with a less complete digestion in order to obtain viable cells. Mechanical chopping of tissue is essential here rather than using the Stomacher; whereas cells loosely associated with tissue (i.e. resident leukocytes) are released undamaged by the Stomacher, architectural cells with tight junctions appear to be sheared by the actions of the machine. 7. We use a variety of density-gradient materials (including Nycodenz, percoll and metrizamide) from different manufacturers. The important parameter is to ensure that the correct density of solution is used. However, whilst more expensive, we currently favour Nycodenz for ease of use. 8. Maintenance of the tissue sample at 4◦ C whilst incubating with magnetic beads reduces phagocytosis of the beads by tissue macrophages and thus improves the purity of the final cell preparation. 9. It is essential to ensure that the endothelial monolayer is completely confluent prior to the flow-based assay as “patchy” cells have a tendency to detach from the substrate upon introduction of shear stress. Monolayer integrity is improved by collagen coating of the microslides, but we also use fibronectin for some cell types. At low shear stress (<0.05 Pa), leukocytes may adhere to exposed matrix if the endothelial monolayer is patchy, stressing the need for confluent monolayers. 10. Whilst leukocytes will bind well to unstimulated endothelial monolayers in “static” adhesion assays, the presence of shear stress confers the necessity for adequate expression of capture receptors in order to permit adhesion. Thus, we note little or no binding of most (but not all) leukocytes to unstimulated monolayers under flow. 11. Perfusion of a million lymphocytes per millilitre over cytokine-stimulated endothelium leads to reproducible numbers of adherent cells sufficiently well distributed over the monolayer to permit accurate analysis of adhesion events by eye or digitally. We find variation in the

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adhesion of different populations of cells (and interindividual variation) and thus optimal perfusion cell counts may need to be determined for each application. However, as long as the cell count, unit area, flow rate and perfusion period are recorded and standardised, adhesion can be normalised to adherent cells/mm2 /106 cells perfused for each experiment. 12. We often fix our cytokine-treated endothelial monolayers prior to performing adhesion assays as this permits flexibility in storage of samples and reduces problems associated with the loss of monolayers due to over enthusiastic washing. In our hands, we find no significant reduction in adhesion using this methodology. However, it is possible to perform the same assay using “live” monolayers which are then fixed after the adhesion stage of the experiment. 13. When using CellTracker Green-labelled lymphocytes in an “automated” adhesion assay, it is necessary to use phenolred-free media formulations to minimise problems associated with autofluorescence which can reduce the sensitivity of detection by the plate reader. 14. Our experience suggests that many problems associated with endothelial-based transmigration assays in culture inserts relate to confluency of the endothelial monolayer. Assays become less reproducible when either too many or too few cells are seeded onto the inserts. We recommend performing pilot experiments to determine the loading density required to give confluency and minimise “piling up” of endothelium at the centre of the wells. Endothelial attachment may be facilitated by precoating the inserts with matrix components such as gelatin or collagen. Such treatment may also help promote adhesion and transmigration of lymphocytes by improving their “traction” on the insert. However, most inserts are designed to be used in the absence of such treatment which may slow down transit of cells through the pores (compared to uncoated inserts). 15. Users will note variability in migration responses when lymphocytes from different patients are compared. This reflects inter-individual variation particularly of chemokine receptor expression. We have also noted some variability in the performance of different batches of chemokines and even individual culture inserts (particularly the lightimpermeable ones). Thus, we recommend pilot experiments to determine optimal doses of chemokines for each experiment and the use of at least three replicates of each condition in every experiment. Users should avoid using high concentrations of human serum in media for such

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assays as endogenous chemokines present may interfere with assay performance. 16. Data from kinetic transmigration assays suggest that most chemokine-dependent migration of human lymphocytes through culture inserts takes place within 3 h. Whilst more cells may go through given longer incubation periods, we have noted more non-specific migration when assays are left to run for too long. However, cells do take longer to penetrate membranes which are coated with endothelial cells. The number of recovered cells can be increased by gently scraping the bottom surface of the insert after incubation to recover cells which have transmigrated through the membrane but not fallen into the bottom reservoir. Also, users should pay attention to the pore size on their chosen insert. Non-specific movement of cells increases significantly if the holes are too large and we recommend using not larger than 5 μM for lymphocytes.

References 1. Ley K, Laudanna C, Cybulsky MI, Nourshargh S. (2007) Getting to the site of inflammation: the leukocyte adhesion cascade updated. Nat Rev Immunol 7, 678–89. 2. Marelli-Berg FM, Cannella L, Dazzi F, Mirenda V. (2008) The highway code of T cell trafficking. J Pathol 214, 179–89. 3. June CH, Blazar BR. (2006) Clinical application of expanded CD4+25+ cells. Semin Immunol 18, 78–88. 4. Cobbold M, Khan N, Pourgheysari B, et al. (2005) Adoptive transfer of cytomegalovirusspecific CTL to stem cell transplant patients after selection by HLA-peptide tetramers. J Exp Med 202, 379–86. 5. Ghosh S. (2003) Therapeutic value of alpha4 integrin blockade in inflammatory bowel disease: the role of natalizumab. Expert Opin Biol Ther 3, 995–1000. 6. Feagan BG, Greenberg GR, Wild G, et al. (2005) Treatment of ulcerative colitis with a humanized antibody to the alpha4beta7 integrin. N Engl J Med 352, 2499–507. 7. Mora JR, Iwata M, Eksteen B, et al. (2006) Generation of gut-homing IgA-secreting B cells by intestinal dendritic cells. Science 314, 1157–60. 8. Stagg AJ, Kamm MA, Knight SC. (2002) Intestinal dendritic cells increase T cell expression of alpha4beta7 integrin. Eur J Immunol 32, 1445–54. 9. Tacken PJ, de V I, Torensma R, Figdor CG. (2007) Dendritic-cell immunotherapy: from

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ex vivo loading to in vivo targeting. Nat Rev Immunol 7, 790–802. Curbishley SM, Eksteen B, Gladue RP, Lalor P, Adams DH. (2005) CXCR3 activation promotes lymphocyte transendothelial migration across human hepatic endothelium under fluid flow. Am J Pathol 167, 887–99. Eksteen B, Grant AJ, Miles A, et al. (2004) Hepatic endothelial CCL25 mediates the recruitment of CCR9+ gut-homing lymphocytes to the liver in primary sclerosing cholangitis. J Exp Med 200, 1511–7. Heydtmann M, Hardie D, Shields PL, et al. (2006) Detailed analysis of intrahepatic CD8 T cells in the normal and hepatitis C-infected liver reveals differences in specific populations of memory cells with distinct homing phenotypes. J Immunol 177, 729–38. Norris S, Collins C, Doherty DG, et al. (1998) Resident human hepatic lymphocytes are phenotypically different from circulating lymphocytes. J Hepatol 28, 84–90. Sigmundsdottir H, Pan J, Debes GF, et al. (2007) DCs metabolize sunlight-induced vitamin D3 to ‘program’ T cell attraction to the epidermal chemokine CCL27. Nat Immunol 8, 285–93. Stamper HB Jr., Woodruff JJ. (1976) Lymphocyte homing into lymph nodes: in vitro demonstration of the selective affinity of recirculating lymphocytes for highendothelial venules. J Exp Med 144, 828–33.

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16. McNab G, Reeves JL, Salmi M, Hubscher S, Jalkanen S, Adams DH. (1996) Vascular adhesion protein 1 mediates binding of T cells to human hepatic endothelium. Gastroenterology 11, 522–8. 17. Yoong KF, McNab G, Hubscher SG, Adams DH. (1998) Vascular adhesion protein-1 and ICAM-1 support the adhesion of tumorinfiltrating lymphocytes to tumor endothelium in human hepatocellular carcinoma. J Immunol 160, 3978–88. 18. Edwards S, Lalor PF, Tuncer C, Adams DH. (2006) Vitronectin in human hepatic tumours contributes to the recruitment of lymphocytes in an alpha v beta3-independent manner. Br J Cancer 95, 1545–54. 19. Aird WC. (2007) Phenotypic heterogeneity of the endothelium: II. Representative vascular beds. Circ Res 100(2), 174–90. 20. Yano K, Gale D, Massberg S, et al. (2007) Phenotypic heterogeneity is an evolutionarily conserved feature of the endothelium. Blood 109, 613–5. 21. Lalor PF, Edwards S, McNab G, Salmi M, Jalkanen S, Adams DH. (2002) Vascular adhesion protein-1 mediates adhesion and transmigration of lymphocytes on human hepatic endothelial cells. J Immunol 169, 983–92. 22. Lalor PF, Lai WK, Curbishley SM, Shetty S, Adams DH. (2006) Human hepatic

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sinusoidal endothelial cells can be distinguished by expression of phenotypic markers related to their specialised functions in vivo. World J Gastroenterol 12, 5429–39. Seternes T, Sorensen K, Smedsrod B. (2002) Scavenger endothelial cells of vertebrates: a nonperipheral leukocyte system for high-capacity elimination of waste macromolecules. Proc Natl Acad Sci USA 99, 7594–7. Xu XD, Ueta H, Zhou S, et al. (2008) Trafficking of recirculating lymphocytes in the rat liver: rapid transmigration into the portal area and then to the hepatic lymph. Liver Int 28, 319–30. Abbitt KB, Nash GB. (2001) Characteristics of leucocyte adhesion directly observed in flowing whole blood in vitro. Br J Haematol 112, 55–63. Cooke BM, Usami S, Perry I, Nash GB. (1993) A simplified method for culture of endothelial cells and analysis of adhesion of blood cells under conditions of flow. Microvasc Res 45, 33–45. Eksteen B, Grant AJ, Miles A, et al. (2004) Hepatic endothelial CCL25 mediates the recruitment of CCR9+ guthoming lymphocytes to the liver in primary sclerosing cholangitis. J Exp Med 200, 1511–7.

Chapter 16 Tracking Antigen-Experienced Effector T Cells In Vitro and In Vivo Claire L. Gorman, Claudia Monaco, Enrico Ammiratti, Anna-Chiara Vermi, Federica M. Marelli-Berg, and Andrew P. Cope

Abstract The TCR complex is a multisubunit complex, comprising at least eight transmembrane units. The clonotypic TCR α and β chains are responsible for antigen recognition, whilst the invariant chains of the CD3 complex (δ, ε and γ) and two zeta (ζ) polypeptides couple antigen recognition to downstream signal transduction pathways. TCRζ (CD247) functions as an amplification module in the TCR signalling cascade and is also essential for the assembly and surface expression of the TCR/CD3 complex. Loss of TCRζ expression is common in chronic infectious and inflammatory diseases, as well as in cancer. Previous work has indicated that TCRζlow -expressing cells phenotypically resemble antigen-experienced effector T cells. Here, we describe the derivation of a flow cytometry-based TCRζ expression index for the purpose of more precisely defining TCRζ expression, in addition to utilising a simple transmigration assay in the demonstration that TCRζdim T cells have intrinsic migratory properties that may explain their accumulation at sites of inflammation. Key words: T cells, T-cell receptor zeta, flow cytometry, cell migration.

1. Introduction The evaluation of mononuclear cell subsets by flow cytometry has revolutionised cellular immunology. Indeed, nowhere has this had a greater impact than in defining subsets of T and B lymphocytes. More recently, technologies and reagents have been developed that provide opportunities for exploring the expression of intracellular molecules that have an impact on cell function or responsiveness to extracellular cues. A particularly good example are those reagents that facilitate the analysis of TCR expression. F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_16, © Springer Science+Business Media, LLC 2010

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The TCR complex is a multisubunit complex, comprising at least eight transmembrane units. The clonotypic TCR α and β chains are responsible for antigen recognition (1), whilst the invariant chains of the CD3 complex (δ, ε and γ) and two zeta (ζ) polypeptides couple antigen recognition to downstream signal transduction pathways (2). The invariant chains possess immune-receptor tyrosine-based activation motifs (ITAMs); the CD3 γ, δ and ε subunits each have one ITAM, whilst each ζ chain possesses three ITAMs. Thus, TCRζ (CD247) functions as an amplification module in the TCR signalling cascade (3). The ζ chain is also essential for the assembly and surface expression of the TCR/CD3 complex as well as in positive and negative selection during thymic development of T cells, as demonstrated in studies of T cells from TCRζ-deficient mice (4, 5). A growing literature suggests that loss of TCRζ expression is a common finding in chronic infectious (6) and inflammatory diseases (7–9), as well as in cancer (10). This is particularly true in peripheral blood lymphocytes but has also been described at sites of inflammation, such as the rheumatoid synovial joint, but also in tumour infiltrating T cells. Functional correlates are less clear. While it may be supposed that loss of TCRζ expression in tumour infiltrates may have an impact on tumour immunity, the impact of attenuated TCR signals in chronic inflammation or infections is less clear. Previous studies from our laboratory have indicated that TCRζlow -expressing cells (or TCRζdim T cells) carry the phenotypic finger print of antigen-experienced effector T cells, retaining the capacity to express inflammatory cytokines while at the same time being relatively refractory to TCR stimulation (11). Here, we describe a method to define more precisely TCRζ expression through the derivation of a TCRζ expression index. This is important because careful scrutiny of TCRζ expression shows quite marked heterogeneity in T-cell and NK-cell subsets, even in healthy donors. We also exploit a simple transmigration assay to demonstrate that TCRζdim T cells have intrinsic migratory properties that are likely to explain their accumulation at sites of inflammation. We propose that this property may arise through failure of a default TCR-dependent stop signal that might otherwise preclude migration through activated endothelium.

2. Materials 2.1. Isolation of Peripheral Blood Mononuclear Cells (PBMCs) by Density-Gradient Centrifugation

1. Hanks buffered saline solution (HBSS; PAA Laboratories, Pasching, Germany). 2. Lymphoprep (for human PBMCs; Cedarlane Laboratories, Ontario, Canada). 3. Fresh whole venous blood.

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4. Serum-free RPMI 1640 containing L-glutamine (300 mg/ ml) and HEPES (5957.5 mg/ml, 25 mM). 5. Freezing mixture: 90% sterile filtered foetal calf serum (FCS) with 10% DMSO. 6. Trypan blue. 7. Haemocytometer. 8. Virkon (bleach). 2.2. Flow Cytometry 2.2.1. Cell Surface Staining of PBMCs

All antibodies were purchased from BD Biosciences Pharmingen (San Diego, CA, USA). 1. Mouse IgG1 -PerCP (clone X40). 2. Anti-CD3ε-PerCP (clone SK7). 3. Anti-CD4-PerCP (clone SK3). 4. Anti-CD8-PerCP (clone SK1). 5. Mouse IgG1 -APC (clone MOPC-21). 6. Anti-CD16-APC (clone 3G8). 7. Anti-CD56-APC (clone B159). 8. Anti-CD45RO-FITC (clone UCHL1). 9. FACs buffer: 2% BSA in 1× phosphate-buffered saline (PBS), plus 0.02% sodium azide. 10. 1× PBS. 11. FACS tubes (Falcon).

2.2.2. Intracellular Staining of PBMCs for TCR ζ Expression

1. Mouse IgG1 -PE (Immunotech, Beckman Coulter, Hialeah, FL). 2. Anti-TCRζ-PE (clone TIA-2; Immunotech, Beckman Coulter, Hialeah, FL). 3. Anti-TCRζ-PE antibody, clone 6B10.2 (Santa Cruz Biotechnology Inc., Santa Cruz, CA, USA).1 4. FACS buffer (as above). 5. PBS (1×). 6. Paraformaldehyde (2%). 7. Permeabilisation buffer (FACS buffer plus 0.5% saponin).

2.3. Preparation of Human Umbilical Vein Endothelial Cells (HUVECs)

1. Human umbilical vein samples. 2. Collagenase (0.25 mg/ml from Clostridium histolyticum; Boehringer Mannheim GmbH, Mannheim, Germany).

1 Clone 6B10.2 is an alternative anti-TCRζ-PE antibody, which recognises a different cytoplasmic domain epitope.

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3. RPMI 1640 containing 10% FCS, 10% newborn calf serum, heparin (90 μg/ml), and endothelial cell growth supplement (20 μg/ml). 4. TNF (10 ng/ml) (R&D systems GmbH, Wiesbaden, Germany). 5. Gelatine (1%). 6. Trypsin (0.05%)/EDTA (0.02%) in D-PBS. 7. Six-well (4.67 cm2 ) Transwell plates (Corning). 2.4. Transendothelial Migration Assay

1. HUVECs plated into six-well (4.67 cm2 ) Transwell plates. 2. Serum-free RPMI 1640 containing L-glutamine (300 mg/ml) and HEPES (5957.5 mg/ml, 25 mM). 3. Complete medium (RPMI 1640 plus L-glutamine and HEPES with 10% FCS, 1% penicillin/streptomycin and 1% sodium pyruvate added). 4. Chemokines: CXCL-10, 20 ng/ml; CCL5, 10 ng/ml or CXCL-12, 10 ng/ml (R&D systems GmbH, Wiesbaden, Germany). 5. Trypan blue. 6. Haemocytometer.

3. Methods 3.1. Isolation of PBMCs by Density-Gradient Centrifugation

1. Dilute whole blood 1:2 in HBSS in a 50-ml Falcon tube. 2. Put 20 ml Lympholyte in a fresh 50-ml Falcon tube. 3. Layer up to 30 ml of the diluted whole blood carefully onto Lympholyte. 4. Centrifuge at 750×g for 20 min at room temperature with the centrifuge brake off. 5. Carefully collect mononuclear cell layer at interface into a fresh 50-ml tube. Wash with an equal volume of HBSS. 6. Centrifuge at 750×g for 10 min, with the brake on. 7. Aspirate supernatant into container, ensuring thorough decontamination with Virkon. 8. Resuspend in 5–10 ml of serum-free RPMI and count live cells in a 10 μl sample, using Trypan blue. 9. Centrifuge cells at 450×g for 10 min at 4◦ C, with the brake on. 10. Aspirate and discard the supernatant. 11. Gently resuspend the pellet on ice in freezing mixture to a density of 5 × 106 –1 × 107 cells/ml in a labelled cryovial.

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12. Transfer cryovials on ice to –80◦ C freezer, storing them in a polystyrene box or wrapped in cotton wool. 13. The next day, transfer vials to liquid nitrogen stores. For freezing/thawing PBMCs, see Note 1. 3.2. Flow Cytometry 3.2.1. Cell Surface Staining

NB: All washes are performed at 450×g for 5 min at 4◦ C. 1. After isolation, resuspend the PBMCs/T cells in FACS buffer and pipette approximately 5 × 105 cells (in 100 μl) into each FACS tube. 2. Wash cells with 1 ml FACS buffer. Decant the supernatant, leaving the cell pellet. 3. Add the appropriate conjugated antibodies for cell surface staining: concentration and incubation conditions vary. It is recommended that titration of the antibody be performed to give optimum staining signals. Generally, for the above-listed PerCP-conjugated antibodies, incubate for 20 min at room temperature in the dark. For the APC-conjugated antibodies, incubate for 30 min at ◦ 4 C (i.e. fridge or on ice) in the dark. For the FITC-conjugated antibodies, incubate for 30 min at 4◦ C in the dark. 4. Proceed with intracellular staining (Section 3.2.2 ).

3.2.2. Intracellular Staining for TCRζ Expression

NB: All washes are performed at 450×g for 5 min at 4◦ C. 1. Wash surface-stained cells in 1 ml FACS buffer. Discard the supernatant, leaving the cell pellet. 2. Wash in 1 ml of 1× PBS. Decant supernatant, leaving the cell pellet. 3. Fix with 500 μl of 2% paraformaldehyde – incubate for 20 min on ice. Do NOT decant. 4. Wash with 2 ml of 1× PBS (see Note 2). 5. Incubate in 500 μl permeabilisation buffer at room temperature for 10 min. 6. Spin (1,500 rpm for 5 min at 4◦ C) in this 500 μl permeabilisation buffer. 7. Add 50–70 μl permeabilisation buffer to the pellet and resuspend. 8. Add 10 μl PE-conjugated anti-TCRζ antibodies to the pellet (see Note 3). Incubate on ice (or in the fridge at 4◦ C) for 30 min in the dark. 9. Wash with 1 ml permeabilisation buffer. 10. Resuspend cells in 250 μl FACS buffer. 11. Store cells at 4◦ C until ready for flow cytometric analysis (as the cells are fixed, they can be left for a few days).

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3.2.3. Derivation of the TCRζ Expression Index

To quantify TCRζ expression in T cells from healthy donors, a TCRζ expression index can be used which is based on two independent flow cytometric parameters (also see Notes 4–7): 1. Constitutive expression of TCRζ can be determined by calculating the ratio of the mean fluorescence intensity (MFI) of the TCRζ+ population to the MFI of the TCRζ– population, comprising B cells and monocytes (Fig. 16.1a). 2. The ratio of the number of circulating TCRζbright to TCRζdim cells can also be calculated (Fig. 16.1b). These indices can be determined separately for total CD3+ , CD4+ and CD8+ lymphocyte subsets as well as for CD56+ or CD16+ NK-cell subsets. This is based on the acquisition of 10,000 cells using CellQuest or FACSDiva software (Becton Dickinson). A TCRζ MFI index

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Fig. 16.1. FACS-based expression index. This has two components: (a) Constitutive expression of TCRζ was determined by calculating the ratio of the mean fluorescence intensity (MFI) of the TCRζ+ population to the MFI of the TCRζ− population, comprising B cells and monocytes. This is termed the TCRζ MFI index. (b) The ratio of the number of circulating TCRζbright to TCRζdim cells is calculated. This is termed the TCRζ bright/dim ratio. Here, the index is shown in PB from a healthy individual.

3.3. Preparation of Human Umbilical Vein Endothelial Cells (HUVECs)

1. Isolate endothelial cells from human umbilical veins by digestion for 10 min at 37◦ C with 0.25 mg/ml collagenase. 2. Culture cells in RPMI 1640 containing 10% FCS, 10% newborn calf serum (NCS), heparin (90 μg/ml) and endothelial cell growth supplement (20 μg/ml) (see Note 8). 4. HUVECs are then stimulated with 10 ng/ml TNF for 48 h.

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5. Coat six-well (4.67 cm2 ) Transwell plates (Corning) with 1% gelatine and incubate with the supplemented RPMI 1640 at 37◦ C for 24 h. 6. As HUVECs are adherent cells, they require trypsinisation to release them from the flask: add 5 ml trypsin in EDTA and incubate at 37◦ C for 5 min. Cells are then washed in RPMI to remove trypsin. 7. Following trypsinisation, plate 2 × 105 HUVECs in monolayers onto each gelatine-coated Transwell in fresh RPMI supplemented with FCS/NCS. 8. Plated HUVECs are then incubated at 37◦ C for 24 h. 3.4. Transendothelial Migration Assay

1. Twenty-four hours after plating HUVECs on to Transwells, 5 × 106 thawed PBL or fresh elutriated T cells are added to the upper chamber of the Transwell. The Transwell system is illustrated in Fig. 16.2a. 2. A sample of PBL/T cells (5 × 105 cells) is retained for baseline TCRζ staining.

Fig. 16.2. Transmigration of healthy donor PBL stained for TCRζ through activated endothelial cell (HUVEC) monolayers. (a) Diagram of the Transwell system. (b) Representative FACS plots showing the percentage of TCRζbright (right upper quadrant) and TCRζdim (right lower quadrant) of the total TCRζ+ population in the upper and lower chambers. EC = endothelial cell.

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3. Add chemokines (e.g. CXCL-10, 20 ng/ml; CCL5, 10 ng/ml or CXCL-12, 10 ng/ml) to the lower chamber at time 0 to assess their effects on transmigration. 4. The Transwell cell system is then incubated at 37◦ C for 24 h. 5. Harvest cells from both upper and lower chambers of the system in turn by aspiration, taking care to harvest all the cells by repeated pipetting. 6. Count cells using a haemocytometer and confirm viability by Trypan blue exclusion. 7. Stain cells for FACS analysis (for surface staining, see Section 3.2.1; for intracellular staining for TCRζ, see Section 3.2.2). 8. Numbers of migrating TCRζbright or TCRζdim T cells are subsequently determined by flow cytometry. A representative FACS staining is illustrated in Fig. 16.2b (see also Notes 9 and 10).

4. Notes 1. After isolation, PBMCs should be frozen more gradually in a polystyrene box or wrapped in cotton wool in –80◦ C before transferring to liquid nitrogen at least 24 h later. Thawing samples should, conversely, be performed as quickly as possible with immediate transfer to the complete medium. The initial spin should be performed at a slightly lower speed (e.g. 1,200 rpm with the brake off: this increases cell recovery). In comparison to frozen/thawed samples with fresh samples, although more debris and cell death are apparent, there is no obvious deleterious effect on TCRζ staining. Freezing of donor PBMC samples allows for multiple samples to be analysed simultaneously; as there is inherent inter-experimental variability in the process of intracellular staining and in the flow cytometric analysis itself, analysis of samples in batches is preferable. However, to exclude confounding effects from different batches, multiple stepwise linear regression can be employed using the batch as the independent variable; we have shown previously that the batch did not have any significant effect on any of the indices (12). 2. During the process of intracellular staining for TCRζ expression, one can leave the cells overnight in 250 μl FACS buffer following Step 4 (i.e. following washing with 1× PBS). Wash with 1 ml FACS buffer to recover cells the following day. 3. As the TCRζ chain has a short nine-amino acid extracellular domain, monoclonal antibodies that detect the intracellular

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cytoplasmic domain epitopes after fixing and permeabilisation are used to analyse TCRζ expression. Isotype-matched control antibodies are used to establish specificity of expression (Fig. 16.3a). We have compared two clones of antiTCRζ-PE antibody which recognises different intracellular epitopes: 6B10.2 recognises a transmembrane epitope, whereas TIA-2 recognises a cytoplasmic domain epitope. As previously demonstrated in our laboratory, both antibodies recognise phosphorylated as well as unphosphorylated forms of the TCRζ chain, implying that any loss of signal is not due to activation-induced phosphorylation of the ζ chain polypeptide (Dr J Clark, personal communication). As shown in Fig. 16.3b, there is little difference in the resultant staining of the CD3+ cells from the same healthy donor. 6B10.2 can also be used, however, to stain mouse T cells for TCRζ, whereas TIA-2 is specific to human cells. TIA-2 is only manufactured conjugated

Fig. 16.3. TCRζ expression in healthy PB T cells measured by FACS analysis. PBL were obtained from the same healthy donor, stained for TCRζ expression and analysed by FACS. (a) IgG (PE) isotype control; (b) comparison of different TCRζ-PE antibodies using TIA-2 and 6B10.2 clones; (c) TCRζ staining without permeabilisation using TIA-2 (PE) antibody.

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to PE, whereas 6B10.2 may alternatively be conjugated to FITC. However, we have found that the fluorescence of the 6B10.2–PE conjugate is more reliable (as well as giving a stronger fluorescent signal) when compared to that of the FITC-conjugated antibody. Furthermore, we have found that other FITC-conjugated antibodies are often difficult to compensate for when using them in flow cytometric analysis with the anti-TCRζ-PE-conjugated antibodies. To determine the importance of permeabilisation in the process of TCRζ staining (i.e. to ensure that the recognised epitope was predominantly intracellular), we have compared staining of PBL either with or without permeabilisation. As shown in Fig. 16.3c, TCRζ staining was almost completely absent in the non-permeabilised cells. 4. In order to semi-quantitatively and reliably measure TCRζ expression in PBL, we have established a novel FACS-based TCRζ expression index (11, 12). Previously, measurement of TCRζ expression has been limited to simply measuring the MFI of the TCRζ-stained cells of interest using flow cytometry. Attempts have been made to more systematically quantify TCRζ expression using FACS. In patients with cancer, for example, the TCRζ MFI of a patient’s T cells was measured relative to the TCRζ expression of T cells from a normal control. A shift of two standard deviations or more was considered to be significant (13). The index that we have established, however, is able to assess a number of components of TCRζ expression, including the number of circulating cells expressing low levels of TCRζ. Using the index established in our laboratory, markedly lower TCRζ expression was confirmed in a cohort of patients with SLE compared to healthy individuals. In addition, a wide heterogeneity of TCRζ expression was identified in both SLE patients and healthy people (12). However, in direct comparison to the healthy controls, TCRζ expression, using both components of the expression index, has been found to be several orders of magnitude lower. When statistically compared to the healthy controls using the Mann–Whitney U test, both components of the TCRζ expression index were significantly lower in SLE patients (p < 0.0001 for the CD3+ subset; Fig. 16.4). 5. In comparison to the TCRζ staining using this TCRζ expression index, a surprising degree of heterogeneity of TCRζ expression has been found to exist between healthy donors. In the four healthy donors shown in Fig. 16.5, the percentage of TCRζdim cells vary from 4% (A) to 41% (C). In addition, it has been observed that cells expressing high levels of TCRζ were present in both CD56+ and CD16+ populations (Fig. 16.6) (11).

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Fig. 16.4. Heterogeneity of PB T-cell (CD3+ ) TCRζ expression in 60 healthy controls and 30 patients with SLE. (a) CD3+ TCRζbright/dim ratio; (b) CD3+ TCRζ MFI index.

6. TCRζ expression using both components of the TCRζ expression index has been measured longitudinally in CD3+ , CD4+ and CD8+ T-cell subsets at two different time points 10–12 months apart; analysis using the Bland– Altman technique (i.e. linear regression analysis comparing the average TCRζ expression at the two time points with the absolute difference between expression at the two time points) not only suggested that TCRζ expression is relatively stable over time but also suggested that the expression index is robust and reliable (12). 7. We have observed that, during an acute inflammatory response, TCRζdim cells are depleted from the peripheral blood compartment and enriched in inflamed tissue such as the rheumatoid synovial fluid. Furthermore, TNF blockade is observed to reverse this, with an observed increase in TCRζdim cells in the PB following anti-TNF therapy. It has subsequently been shown that TCRζdim T cells accumulated and persisted in the PB of patients who responded well to anti-TNF treatment; this may further explain why rheumatoid disease flares on withdrawing anti-TNF therapy, after which TCRζdim T cells would migrate back to the

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Fig. 16.5. Heterogeneity of TCRζ expression in PBL from four healthy individuals. PBL were isolated from four healthy donors (a, b, c and d) by Ficoll extraction, stained for TCRζ and CD3e and analysed by FACs. The numbers in red represent the percentage of TCRζbright (upper quadrant) and TCRζdim (lower quadrant) cells present. The lines represent the gradients between TCRζ and CD3e expression in the TCRζbright cell subset in each of the different donors. In (a), quadrant [a] represents CD3- TCRζ+ cells (i.e. the NK cell population); quadrant [b] represents TCRζbright T cells; quadrant [c] represents the double negative population of monocytes and B cells that do not express the TCRζ chain; and TCRζdim cells are shown in quadrant [d].

synovial joint compartment and re-initiate a local inflammatory response (11). In general, however, in vitro transmigration studies using PBL from patients with RA have not suggested that anti-TNF therapy reduces the migratory competence of TCRζdim cells. As the endothelial cells remain activated by TNF in this in vitro model, this indicates that, in vivo, the observed reduced migration of TCRζdim cells following anti-TNF therapy may be consequent to reduced endothelial activation. 8. It is recommended that HUVECs should be used in experiments before the fourth passage (14). 9. The main limitation in these studies is the number of migrating cells. Thus, to observe migration in a 24-h period, at least 5 × 106 cells must be added to the upper chamber of the system at time 0. It is necessary to ensure thorough harvesting of cells from upper and lower chambers using multiple rinses with serum-free RPMI. It is also advisable to rinse both surfaces of the Transwell membrane to maximise the cell harvest. It has been found that on exceeding the 24-h incubation period, very little further migration is observed with a significant increase

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Fig. 16.6. TCRζ expression in (a) CD56+ and (b) CD16+ cells. PBL were obtained from two different healthy donors and stained for TCRζ, CD56 and CD16 expression. These FACS plots show that within both CD56+ and CD16+ populations there are significant numbers of cells expressing high levels of TCRζ.

in cell debris. Notably, transmigration and cell viability is maximised if fresh rather than frozen PBMCs are used in the assay. We have demonstrated that, by 24 h, approximately 20–25% of fresh PBMCs from healthy donors migrate across these TNF-stimulated endothelial monolayers cultured in Transwells and that T cells migrating to the lower chamber expressed lower levels of TCRζ (Fig. 16.2b). 10. In the transmigration assay, a control well (i.e. without endothelial cells or Transwell) is required to demonstrate that transmigration does not perturb TCRζ expression (i.e. to show that the proportions of TCRζbright and TCRζdim T cells are similar to those in PB cultured for the same period in the absence of endothelial cells or Transwell). If chemokines are being used in the lower chambers, some wells without chemokines are also required. References 1. Meuer SC, Cooper DA, et al. (1983) “Identification of the receptor for antigen and major histocompatibility complex on human inducer T lymphocytes”. Science 222(4629), 1239–42. 2. Baniyash M. (2004) “TCR zeta-chain downregulation: curtailing an excessive

inflammatory immune response”. Nat Rev Immunol 4(9), 675–87. 3. Irving BA, Weiss A. (1991) “The cytoplasmic domain of the T cell receptor zeta chain is sufficient to couple to receptor-associated signal transduction pathways”. Cell 64(5), 891–901.

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4. Ohno H, Aoe T, et al. (1993) “Developmental and functional impairment of T cells in mice lacking CD3 zeta chains”. EMBO J 12(11), 4357–66. 5. Yamazaki T, Arase H, et al. (1997) “A shift from negative to positive selection of autoreactive T cells by the reduced level of TCR signal in TCR-transgenic CD3 zeta-deficient mice”. J Immunol 158(4), 1634–40. 6. Seitzer U, Kayser K, et al. (2001) “Reduced T-cell receptor CD3zeta-chain protein and sustained CD3epsilon expression at the site of mycobacterial infection”. Immunology 104(3), 269–77. 7. Liossis SN, Ding XZ, et al. (1998) “Altered pattern of TCR/CD3-mediated proteintyrosyl phosphorylation in T cells from patients with systemic lupus erythematosus. Deficient expression of the T cell receptor zeta chain”. J Clin Invest 101(7), 1448–57. 8. Pang M, Setoyama Y, et al. (2002) “Defective expression and tyrosine phosphorylation of the T cell receptor zeta chain in peripheral blood T cells from systemic lupus erythematosus patients”. Clin Exp Immunol 129(1), 160–8. 9. Nambiar MP, Mitchell JP, et al. (2003) “Prevalence of T cell receptor zeta chain deficiency in systemic lupus erythematosus”. Lupus 12(1), 46–51.

10. Kono K, Ichihara F, et al. (1998) “Expression of signal transducing T-cell receptor zeta molecules after adoptive immunotherapy in patients with gastric and colon cancer”. Int J Cancer 78(3), 301–5. 11. Zhang Z, Gorman CL, Vermi AC, Monaco C, Foey A, Owen S, Amjadi P, Vallance A, McClinton C, Marelli-Berg FM, Isomaki P, Russell A, Dazzi F, Vyse TJ, Brennan FM, Cope AP. (2007 May 15) TCR{zeta}dim lymphocytes define populations of circulating effector cells that migrate to inflamed tissues. Blood 109(10), 4328–35. 12. Gorman CL„ Russell AI„ Zhang Z„ Cunninghame Graham D„ Cope AP„ Vyse TJ. (2008 Jan 15) Polymorphisms in the CD3Z gene influence TCRζeta expression in systemic lupus erythematosus patients and healthy controls. J Immunol 180(2), 1060–70. 13. Whiteside TL. (2004) “Down-regulation of zeta-chain expression in T cells: a biomarker of prognosis in cancer?” Cancer Immunol Immunother 53(10), 865–78. 14. Monaco C, Andreakos E, et al. (2002) “T cell-mediated signaling to vascular endothelium: induction of cytokines, chemokines, and tissue factor”. J Leukoc Biol 71(4), 659–68.

Chapter 17 Preclinical Testing of Strategies for Therapeutic Targeting of Human T-Cell Trafficking In Vivo Caroline Coisne and Britta Engelhardt Abstract Naive T cells are migratory cells that continuously recirculate between blood and lymphoid tissues. Antigen-specific stimulation of T cells within the lymph nodes reprograms the trafficking properties of T cells by inducing a specific set of adhesion molecules and chemokine receptors on their surface which allow these activated and effector T cells to effectively and specifically home to extralymphoid organs. The observations of organ-specific homing of T cells initiated the development of therapeutic strategies targeting adhesion receptors for organ-specific inhibition of chronic inflammation. As most adhesion receptors have additional immune functions besides mediating leukocyte trafficking, these drugs may have additional immunomodulatory effects. Therapeutic targeting of T-cell trafficking to the central nervous system is the underlying concept of a novel treatment of relapsing remitting multiple sclerosis with the humanized anti-α-4-integrin antibody natalizumab. In this chapter, we describe a possible preclinical in vivo approach to directly visualize the therapeutic efficacy of a given drug in inhibiting T-cell homing to a certain organ at the example of the potential of natalizumab to inhibit the trafficking of human T cells to the inflamed central nervous system in an animal model of multiple sclerosis. Key words: Intravital microscopy, spinal cord window, human T cells, adhesion molecules, trafficking, endothelium, anesthesia, experimental autoimmune encephalomyelitis, blood–brain barrier, inflammation.

1. Introduction In multiple sclerosis (MS) and in its animal model experimental autoimmune encephalomyelitis (EAE), circulating immune cells get access to the central nervous system (CNS), where they start the molecular events leading to inflammation, edema formation and demyelination, and the development of the disabling clinical picture of the disease. Interaction of circulating immune cells with F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, DOI 10.1007/978-1-60761-461-6_17, © Springer Science+Business Media, LLC 2010

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the endothelial blood–brain barrier (BBB) is thus a critical step in the pathogenesis of EAE and MS. In general, lymphocyte recruitment across the vascular wall is regulated by the sequential interaction of different adhesion or signaling molecules on lymphocytes and endothelial cells lining the vessel wall. An initial transient contact of the circulating leukocyte with the vascular endothelium, generally mediated by adhesion molecules of the selectin family and their respective carbohydrate ligands, slows down the leukocyte in the bloodstream. Subsequently, the leukocyte rolls along the vascular wall with greatly reduced velocity. The rolling leukocyte can receive endothelial signals resulting in its firm adhesion to the endothelial surface. Such signals can be chemokines, the activity of which is transduced via G protein-coupled receptors on the leukocyte surface. Binding of a chemokine to its receptor results in a pertussis toxin-sensitive activation of integrins on the leukocyte surface. Only activated integrins mediate the firm adhesion of the leukocyte to the vascular endothelium by binding to their endothelial ligands, which belong to the immunoglobulin (Ig) superfamily. This, ultimately, leads to the extravasation of the leukocyte. Successful recruitment of circulating leukocytes into the tissue depends on the productive leukocyte/endothelial interaction during each of these sequential steps (1). α4β1-Integrin was identified to mediate T-cell adhesion to inflamed vessels in frozen sections of EAE brains in vitro (2) and antibodies blocking α4-integrins were shown, in a variety of animal models, to prevent the development of EAE [summarized in (3)]. Based on these findings, the humanized monoclonal anti-α4-integrin antibody natalizumab was developed for the treatment of MS with the idea to target α4-integrinmediated T-cell extravasation into the CNS. Natalizumab has proved to be highly beneficial in reducing MS disease activity regarding both clinical parameters and MRI measurements of disease activity. Besides mediating T-cell extravasation, α4-integrins have been suggested to be involved in additional T-cell functions such as T-cell activation and polarization (4) and retention of memory T cells in their niches (5). That natalizumab may exert additional immunomodulatory effects in MS patients is supported by the notion that it does bear the rare risk of progressive multifocal leukoencephalopathy (PML), a very rare opportunistic infection (6) (www.nationalmssociety.org/news/newsdetail/index.aspx?nid=260). Thus, direct visualization of natalizumab-mediated inhibition of the extravasation of human T cells into the CNS in MS patients would improve our understanding of the immunomodulatory effects and the possible risks of this α4-integrin-specific drug. Imaging techniques available to us nowadays do not yet allow the visualization of individual circulating T cells within the CNS

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blood vessels of a patient. In contrast, in animal models allowing deep anesthesia and microsurgery for gaining access to the CNS tissues, intravital epifluorescence videomicroscopy has proven to be a powerful tool to investigate T-cell interaction with the BBB in vivo. In order to gain microscopic access to the CNS microcirculation in vivo, cranial window preparations removing the skull have been developed for rodents which allow observation of the pial and cortical, i.e., CNS gray matter, microcirculation (7, 8). In EAE – the disease setting discussed in this chapter – inflammation is located in the CNS white matter with a preference for the spinal cord. Investigation of the interaction of T lymphocytes with the CNS white matter microvasculature thus requires the preparation of a spinal cord window, which allows direct visualization of CNS white matter microcirculation by intravital fluorescence videomicroscopy (9). Both models require sophisticated microsurgical approaches to visualize the CNS microcirculation but then allow live observation of the interaction of individual T cells with the CNS microcirculation in mice. Due to the short observation times of minutes to 1–2 h required to assess the multistep interaction of T cells with the CNS microvasculature, human T cells can be investigated in immunocompetent mouse strains without any toxic side effects. Certainly though, the emerging varieties of humanized mouse models will greatly improve possibilities to investigate the therapeutic potential of drugs targeting human T-cell interaction with the BBB or other microvascular beds by means of intravital microscopy, in preclinical rodent models in vivo under the influence of physiological shear forces. This chapter describes an experimental approach to investigate the efficacy of a humanized anti-α4-integrin antibody natalizumab, which has been approved for the treatment of relapsing remitting MS based on the concept to target α4-integrinmediated homing of T cells to the inflamed CNS, in blocking the interaction of human T cells with the BBB in vivo.

2. Materials 2.1. Human T-Cell Preparation

1. Pipetboy and sterile (plastic or glass) pipettes: 25-and 10-mL 2. Micropipettes and sterile microtips 3. Sterile 50- and 15-mL Falcon tubes 4. Sterile tissue culture flask 5. Sterile 100-μm nylon mesh R , Invitrogen, Switzerland) 6. RPMI-1640 medium (Gibco R 7. L-Glutamine 100× (Gibco , Invitrogen, Switzerland)

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8. Penicillin/streptomycin, containing 10,000 U/mL each R antibiotic (Gibco , Invitrogen, Switzerland) 9. Phosphate-buffered saline (PBS) without Ca2+ /Mg2+ 10. Wash buffer: Hank’s balanced salt solution (HBSS) with phenol red supplemented with 10% calf serum (CS) and R 25 mM HEPES (Gibco , Invitrogen, Switzerland) R -1077 (Sigma-Aldrich, Switzerland) used at 11. Histopaque room temperature (RT) and stored at 4◦ C

12. Human Pan T Cell Isolation Kit II, MS MACS columns, and MACS separator (Miltenyi Biotec, Germany) 13. Trypan blue (Sigma, Switzerland) 14. Neubauer cell-counting chamber 2.2. Labeling of Human T Cells with Fluorescent Dye

1. Pipetboy and sterile (plastic or glass) 5- and 10-mL pipettes. 2. Micropipettes and sterile microtips. 3. Sterile 50- and 15-mL Falcon tubes. 4. Sterile tissue culture Petri dish (100 mm). 5. RPMI-1640 medium. 6. Phosphate-buffered saline (PBS) without Ca2+ /Mg2+ . 7. Wash buffer: Hank’s balanced salt solution (HBSS) with phenol red supplemented with 10% calf serum (CS) and 25 mM HEPES. 8. Isotonic Nycodenz solution (density 1.09 g/mL at room temperature): to 17g Nycodenz powder (Nycomed, Norway) add 26 mL of RPMI-1641 and 10 mL fetal bovine serum. Fill up to 100 mL with sterile distilled water. Nycodenz solution can be stored at 4◦ C, protected from light for several months. 9. Cell TrackerTM Green (CMFDA; Molecular Probes, Eugene, Oregon, USA): 10 mM stock solution stored at –20◦ C in water-free dimethyl sulfoxide (DMSO).

2.3. Preparation of the Spinal Cord Window in SJL Mice with EAE

Induction of EAE in susceptible mouse strains is not the topic of this chapter but has been described elsewhere in great detail (10, 11). For the experiments described here, 7–12-week-old SJL female mice (Harlan, France) with an approximate body weight of 20 g were used. EAE induction in this strain has been extensively described elsewhere (10, 11). Clinical disease is checked daily and scored as described (10, 11). Mice scored with mild disease, i.e., limp tail to hind limp weakness, are used for intravital fluorescence videomicroscopy (IVM) experiments, with a body weight of at least 15 g. 1. Operating stereomicroscope (Leica M651) with up to 40-fold magnification (Leica, Switzerland)

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2. Heating pad 3. Stereotactic rodent head holder 4. Ketamine, xylazine, and acepromazine 5. Sterile 1-mL disposable syringes 6. NaCl isotonic solution (0.9%) (B. Braun AG, Switzerland) 7. Instruments for surgery: Fine Adson forceps and scissors, jewelry forceps, iris scissors, and needle holder 8. 4-0 silk suture 9. Polyethylene catheter (PE-10) 10. Impermeable transparent membrane (Saran wrap) 2.4. Intravital Fluorescence Videomicroscopy

1. Custom-made Mikron IVM500 epi-fluorescent microscope (Mikron Instruments, San Marcos, CA, USA) coupled with a 50 W mercury lamp (HBO 50 microscope illuminator, Zeiss, Switzerland) and combined with blue (exciter 455DF70, dichroic 515DRLP, emitter 515ALP) and green (exciter 525DF45, dichroic 560DRLP, emitter 565ALP) filter blocks 2. Long-distance objectives (4×, 10×, and 20×) (Zeiss, Switzerland) 3. Low-light-imaging camera VE-1000 silicone-intensified target (SIT) system (DAGE-MTI of MC, Inc., Michigan City, IN, USA) 4. Digital videocassette recorder (DVCAMTM DSR-11; Sony, Switzerland) and video tapes PDV-184 N (Sony, Switzerland) 5. Videotimer (MicroImage Video Systems, Boyertown, PA, USA) R 6. Trinitron color video monitor (Sony, Switzerland)

7. Sterile disposable 1-mL syringes 8. TRITC-conjugated dextran (1%, MW = 155,000; SigmaAldrich) in 0.9% NaCl isotonic solution 2.5. Blocking of Cell Surface Adhesion Molecules on Human T Cells

1. Tissue culture centrifuge 2. Pipetboy with sterile 5- and 10-mL sterile pipettes (either glass or plastic) 3. Micropipettes with sterile microtips 4. Sterile 15-mL Falcon tubes and 5-mL polystyrene roundbottom Falcon tubes 5. NaCl isotonic solution (0.9%) 6. Trypan blue (Sigma, Switzerland) 7. Neubauer cell-counting chamber

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8. Sterile, purified, and endotoxin-free blocking antibodies directed against cell adhesion molecules

3. Methods 3.1. Human T-Cell Purification

The day prior to the IVM experiment, peripheral blood mononuclear cells (PBMCs) are isolated from a buffy coat (see Note 1 below) as follows: 1. Fifty milliliters of buffy coat content is diluted 1:2 with PBS in order to obtain four 50-mL Falcon tubes, each containing 25 mL of blood solution. R 2. In each tube, 13 mL of Histopaque -1077 solution (RT) is carefully underlaid by placing the tip of the 10-mL pipette R containing the Histopaque -1077 solution at the bottom of the tube.

3. Centrifugation for 30 min at 400×g, RT, no break. 4. The top layer of the gradient containing the plasma is discarded. The PBMC layer is transferred into a 50-mL Falcon tube containing 30 mL wash buffer and thoroughly resuspended (see Note 2). 5. Cells are centrifuged for 10 min at 250×g, RT with break. The supernatant is then discarded and this washing step is repeated twice by resuspending the cell pellet in 30 mL wash buffer and centrifuging for 10 min at 250×g, RT with break. 6. PBMCs are counted and plated 1×106 cells/mL in RPMI containing 20% FCS, 1% L-glutamine, 1% penicillin/streptomycin (culture medium) and incubated overnight at 37◦ C in a humidified atmosphere containing 7% CO2 . 7. On the day of the IVM experiment, PBMCs are harvested and proceeded for T-cell purification according to the manufacturer’s recommendations using the human Pan T Cell Isolation Kit II (Miltenyi Biotec, Germany) (see Note 3). 3.2. Labeling of Human T Cells with Fluorescent Dye

Cell TrackerTM Green labeling of T cells was chosen because we found before that it does not alter the phenotype of T lymphocytes, i.e., cell surface expression of adhesion molecules, and reproducibly produces a 100% viable and brightly stained T-cell population stably fluorescing for several days. 1. After purification, 5 × 106 human peripheral blood T cells per milliliter are labeled with 2.5 μM Cell TrackerTM Green (CMFDA) in culture medium for 45 min at 37◦ C in the dark. T cells are subsequently washed by adding fresh complete wash buffer and centrifuged for 10 min at 250×g.

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2. Excess dye is removed by plating 5×106 fluorescently labeled T cells in a 100-mm cell culture Petri dish in 10 mL complete medium for 30 min at 37◦ C. 3. Subsequent removal of dead cells and debris is achieved by pelleting labeled T cells for 10 min at 250×g and resuspending them in 1 mL RPMI-1640. The cell suspension is placed on top of a 3 mL Nycodenz solution overlaid by 5 mL FCS in a 15-mL Falcon tube. The gradient is run for 10 min at RT and 250×g without break. 4. Living cells are collected from the interphase between the Nycodenz solution and FCS and washed three times with 30 mL wash buffer for 10 min at 250×g. 5. Cell TrackerTM Green-labeled T cells can be directly used for IVM or stored in complete medium at 37◦ C and 7% CO2 up to 6 h before use. 6. In coordination with the spinal cord window preparation, 5–6×106 T cells in complete medium are collected and centrifuged for 10 min at 250×g. The cell pellet is resuspended in a small volume of 0.9% NaCl. 7. Cells are counted and the volume of 0.9% NaCl is adjusted to the precise cell number in order to obtain 4×106 T cells/300 μL of 0.9% NaCl. T-cell suspension is filled into a 1 mL-syringe and ready for injection into the right carotid artery of the animal. 3.3. Spinal Cord Window Preparation in the Mouse

1. The mouse with clinical EAE is anesthetized by subcutaneous injection of a mixture of 100 mg/kg ketamine hydrochloride and 5.6 mg/kg xylazine, followed 3–5 min later by a subcutaneous injection of 1.5 mg/mL acepromazine. Throughout the experiment, the duration of anesthesia administered to the animal needs to be carefully observed. If necessary, reinjection of half dosage of ketamine/xylazine mixture is used to maintain stabile anesthesia. Throughout the surgery and the entire duration of the IVM experiment, normal body temperature is controlled and maintained by putting the animal on a thermocontrolled heating pad. 2. For systemic administration of fluorescently labeled T cells and 1% TRITC-conjugated dextran as plasma marker, a polyethylene catheter is inserted into the right common carotid artery into the direction of the aortic arch. 3. The mouse is then turned to the prone position and the animal head is fixed in a specific stereotactic holder. After a midline skin incision of 3–4 cm on the neck, the paravertebral musculature is detached from the cervical spinous processes and retracted laterally, exposing the

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vertebral laminae. Retraction is achieved by the use of 4-0 threads. 4. A laminectomy is carried out from C7 to C2 under the stereomicroscope taking care not to injure the underlying spinal cord. Using higher magnification, the dura on top of the spinal cord tissue is opened using jewelry forceps and iris scissors and removed from the dorsal spinal cord avoiding trauma to the spinal cord microvasculature and parenchyma. 5. Finally, the spinal cord preparation is tightly covered with an impermeable and transparent plastic membrane to prevent any dehydration and any influence of the ambient oxygen on the spinal cord window preparation (see Note 4). 3.4. Intravital Fluorescence Videomicroscopy

1. Remaining within the stereotactic head holder, the animal is transferred to the microscope stage. 2. Intravital fluorescence videomicroscopy is performed by epi-illumination techniques using a modified fluorescence microscope with a 50 W mercury lamp, which is attached to a combined blue and green filter blocks. Observations are made using a 4×, 10×, and 20× long-distance objectives, resulting in 80×, 215×, and 440× magnifications, respectively. The microscopic images are recorded using a lowlight-level SIT camera coupled with a video monitor and a videotimer. For later off-line analysis of the videos, images are recorded using a digital videocassette recorder. 3. The spinal cord microvasculature is first observed within the green light epi-illumination at 4× objective, by slowly injecting 100 μL of pre-warmed fluorescent plasma marker 1% TRITC-conjugated dextran in 0.9% NaCl into the arterial catheter. The spinal cord window is divided into two parts by the middle dorsal vein, which delineates a right and a left side of the spinal cord window. In each side, capillary networks and post-capillary venules which drain the middle dorsal vein are visible. 4. To evaluate the interaction of human T cells with the spinal cord white matter endothelium, 4 × 106 Cell TrackerTM Green-labeled T cells are slowly infused in three aliquots of 100 μL each and directly visualized within the spinal cord microcirculation of the spinal cord window using the (blue): light epi-illumination with the 10× objective (see Notes 5, 6, and 7). Per mouse, 4–6 consecutive fields of view can be defined on each side of the spinal cord window. Cells, injected within the right carotid artery, mainly reach the spinal cord microcirculation on the right side of the spinal cord window, where they initiate contact with the microvascular endothelium. For each injection of 100 μL cell

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suspension, a different field of view is recorded for at least an entire minute, in order to be able to evaluate a sufficient number of initial T cell encounters with the endothelium in numerous post-capillary venules during the later off-line analysis. 5. Right after cell infusion, the arterial catheter is rinsed with a maximum of 100 μL pre-warmed 0.9% NaCl in order to ensure injection of the full amount of cells. 6. At different time points after cell injection (10 min, 30 min, 1 h, and 2 h), all fields of view of the spinal cord window are sequentially scanned to record the number of permanently adhering human T cells within the spinal cord microvessels. 3.5. Therapeutic Targeting of Cell Surface Adhesion Molecules on Human T Cells

3.6. Data Analysis

To study the role of adhesion molecules or their ligands in T-cell recruitment across the spinal cord microvasculature or to determine the ability of a drug to interfere with T cells/CNS microvascular interaction, T-cell blasts or the CNS endothelium can be treated with blocking compounds or antibodies. To block adhesion receptors on T cells, 4 × 106 T cells in 300 μL of 0.9% NaCl are incubated with 120 μg adhesion molecule-blocking monoclonal antibody (mAb) for 20 min prior to their injection into the bloodstream. For human drugs, the dosage used is deduced from the dosage recommended in humans by correlating the blood volume of man and mouse. In any case, working with whole antibodies in vivo requires stringent endotoxin-free antibody preparations and appropriate controls. A condition with nonblocking antibodies from the same isotype, as the blocking mAb tested, ascertains any unspecific side effect mediated by the Fc portions of the immunoglobulin. When possible, control antibodies that specifically bind to molecules expressed on the T-cell surface or on the endothelium, and that do not interfere with T-cell trafficking, are rather recommended than irrelevant isotype control antibodies remaining in the circulation. 1. Hemodynamic parameters The hemodynamic parameters of the blood flow are evaluated for each analyzed post-capillary venule according to the Hagen–Poiseuille law as follows: the mean blood flow velocity (Vblood ) represents (Vblood = Vmax /2– (DL /DV )2 ) (μm/s), where Vmax is the velocity of the fastest noninteracting T cell, DL the T-cell diameter, and DV the vessel diameter (12). To quantify the force acting on rolling or adherent leukocytes, the wall shear rate (γ ) was evaluated for each venule as γ = Vblood × 8/DV (s–1 ). The wall shear stress (τ ), dependent on blood viscosity (assumed to be 0.025 Poise), is calculated as: τ = γ ×0.0025 (dyne/cm) (12, 13).

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2. Quantitative analysis of the initial contact of T cells within post-capillary venules (ø 20–60 μm) of the spinal cord white matter in mice with EAE The percentage of T cells that initiate contact with the endothelium from each observed post-capillary venule is determined at the time point of cell injection. Infusing the total amount of T cells at three different time points and observing three different fields of view allow the analysis of the initiation of T-cell interaction with a sufficient number of spinal cord post-capillary venules per mouse. The number of T cells, counted to roll or to capture per vessel, is correlated with the total number of fluorescently labeled T cells, passing through the vessel during an observation period of 1 min, in order to calculate a rolling fraction and a capture fraction. The initial contact fraction is also evaluated and corresponds to the sum of the rolling and capture fraction. The rolling fraction represents the percentage of cells that roll along the vessel wall, while the capture fraction represents the percentage of T cells that are abruptly arrested without any rolling step on the vessel wall. Both rolling and capture events are evaluated by direct observation of the cell movement and can be further ascertained upon velocity criterion derived from the assumption of a parabolic velocity profile in the microvessel (see Fig. 17.1). Vcrit , the velocity of an idealized cell traveling along but not interacting with the vessel wall, is calculated as Vcrit = Vblood × (DL /DV ) × [2 – (DL /DV )]. Consequently, any

Fig. 17.1. Scheme of the parabolic flow profile within a blood vessel The velocity profile in the bloodstream is characterized by a higher blood flow velocity in the centerline of the blood vessel as compared to lower velocities near the vessel wall. Dv , vessel diameter; DL , leukocyte diameter; V, velocity; Vblood , mean blood flow velocity; Vmax , maximal velocity; Vcrit , critical velocity.

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Fig. 17.2. IVM analysis of the firm adhesion of human T cells within the spinal cord white matter post-capillary venules in SJL mouse with EAE. (a) High-magnification image (80×) of the spinal cord white matter microvasculature after injection of TRITC dextran. The spinal cord window is divided into two parts by the middle dorsal vein, which divides the window into an upper and a lower half. In each half, capillary networks and post-capillary venules, draining the middle dorsal vein, are visualized. For the analysis, the right side of the spinal cord window is divided into consecutive fields of view (FOV). Each FOV is observed at higher magnification (215×) within the green light epi-illumination to image the vasculature (b) and the blue: light epi-illumination to visualize the T cells (c) (see Note 5). White dots in (c) are human T cells firmly adherent to the endothelium of a spinal cord microvessel visible in (b), 10 min after cell injection into the right carotid artery.

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cell traveling below Vcrit is considered as a cell interacting with and therefore rolling along the vessel wall and any cell traveling above Vcrit is regarded as a noninteracting cell (12 and (Veling): 4). 3. Quantitative analysis of the firm adhesion Permanently adherent T cells were identified as cells that are stuck to the vessel wall without moving or detaching from the endothelium. Within the capillary network, plugging T-cell blasts are defined as cells that did not move and obviously blocked the capillary lumen, inducing blood flow stasis in the corresponding vascular segment. Permanent T-cell blast adhesion at 10 min, 30 min, 1 h and 2 h after cell injection is expressed as the number of both adherent and plugging T-cell blasts per field of view (FOV) using 10× objective (see Fig. 17.2). Per mouse, 4–6 fields of view can be defined on the right side of the spinal cord window. All the quantifications of firmly adherent T cells per FOV from different mice are pooled to generate a mean ± standard deviation. Alternatively, another method to quantify firmly adherent T cells within the post-capillary can be used while cells do not plug within the capillaries but adhere only within the post-capillary beds. This method consists of calculating the luminal surface area (SA) of each post-capillary venule (20–60 μm diameter) of the right side of the spinal cord window, considering each venule as a tube and applying the formula SA (mm2 ) = Dv × L × π, where DV represents the vessel diameter (mm) and L the length of the post-capillary venule (mm). The total number of firmly adherent T cells is reported to the pooled SA of all post-capillary venules and expressed as number of cells per square millimeter. All the quantifications of firmly adherent T cells per square millimeter from different mice are pooled to generate a mean ± standard deviation. Statistic tests such as the Mann–Whitney U-test to compare two variables and Kruskal–Wallis test to compare more than two variables are applied.

4. Notes 1. Depending on the amount of PBMCs required for the experiment, either heparinized or EDTA-whole blood or buffy coat can be used. 2. PBMCs isolated from a buffy coat can contain lipid aggregates. They can be easily removed by a filtration step using a sterile nylon mesh.

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3. Due to the relative low proportion of T cells that can be visualized within the total spinal cord window (range of 0–500 firmly adherent cells) versus the total number of injected cells (range of several millions), the purity yield of the cell preparation represents a critical issue and should be determined by FACS analysis. 4. Only preparations showing no bleeding and intact microcirculation are considered acceptable for subsequent examination by IVM. 5. This combination of two fluorescent markers, displaying distinct excitation–emission wavelengths, enables the colocalization of Cell TrackerTM Green-labeled T cells within the TRITC-stained vessel lumen. 6. Pay attention to minimize the exposure of the spinal cord window preparation to light as much as possible; otherwise, phototoxic damage of the microvasculature may occur within seconds, resulting in excessive activation of the endothelium. 7. The angioarchitecture of the spinal cord microvasculature allows the visualization of T cell/endothelial interactions within the spinal cord capillaries and post-capillary venules (20–60 μm diameter), but not within pre-capillary arterioles, which are located deeper within the spinal cord parenchyma and thus not visible by means of IVM.

Acknowledgments We owe special thanks to Heidi Tardent for excellent microsurgical preparations and to Lucienne Tritten for critical comments on this chapter. This work has been supported by the National Multiple Sclerosis Society (NMSS, USA), the Swiss MS Society, and the Bern University Research Foundation. We thank Susan (Goelz): (BiogenIdec, USA) for the gift of Natalizumab. CC has been funded by the Fondation pour la Recherche Médicale (FRM, France), the Association pour la Recherche sur la Sclérose En Plaques (ARSEP, France), and the NMSS. References 1. Ley K, Laudanna C, Cybulsky MI, Nourshargh S. (2007) Getting to the site of inflammation: the leukocyte adhesion cascade updated. Nat Rev Immunol 7, 678–89. 2. Yednock TA, Cannon C, Fritz LC, SanchezMadrid F, Steinman L, Karin N. (1992)

Prevention of experimental autoimmune encephalomyelitis by antibodies against alpha 4 beta 1 integrin. Nature 356, 63–6. 3. Engelhardt B, Ransohoff RM. (2005) The ins and outs of T-lymphocyte trafficking to

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Coisne and Engelhardt the CNS: anatomical sites and molecular mechanisms. Trends Immunol 26, 485–95. Mittelbrunn M, Molina A, Escribese MM, Yanez-Mo M, Escudero E, Ursa A, Tejedor R, Mampaso F, Sanchez-Madrid F. (2004) VLA-4 integrin concentrates at the peripheral supramolecular activation complex of the immune synapse and drives T helper 1 responses. Proc Natl Acad Sci USA 101, 11058–63. Epub 2004 Jul 19. Sixt M, Bauer M, Lammermann T, Fassler R. (2006) Beta1 integrins: zip codes and signaling relay for blood cells. Curr Opin Cell Biol 18, 482–90. Yousry TA, Major EO, Ryschkewitsch C, Fahle G, Fischer S, Hou J, Curfman B, Miszkiel K, Mueller-Lenke N, Sanchez E, Barkhof F, Radue EW, Jager HR, Clifford DB. (2006) Evaluation of patients treated with natalizumab for progressive multifocal leukoencephalopathy. N Engl J Med 354, 924–33. Uhl E, Pickelmann S, Rohrich F, Baethmann A, Schurer L. (1999) Influence of plateletactivating factor on cerebral microcirculation in rats: part 2. Local application. Stroke 30, 880–6. Vajkoczy P, Ullrich A, Menger MD. (2000) Intravital fluorescence videomicroscopy to study tumor angiogenesis and microcirculation. Neoplasia (New York) 2, 53–61.

9. Vajkoczy P, Laschinger M, Engelhardt B. (2001) Alpha4-integrin-VCAM-1 binding mediates G protein-independent capture of encephalitogenic T cell blasts to CNS white matter microvessels. J Clin Invest 108, 557– 65. 10. Engelhardt B, Laschinger M, Schulz M, Samulowitz U, Vestweber D, Hoch G. (1998) The development of experimental autoimmune encephalomyelitis in the mouse requires alpha4-integrin but not alpha4beta7-integrin. J Clin Invest 102, 2096–105. 11. Engelhardt B, Vestweber D, Hallmann R, Schulz M. (1997) E- and P-selectin are not involved in the recruitment of inflammatory cells across the blood–brain barrier in experimental autoimmune encephalomyelitis. Blood 90, 4459–72. 12. Ley K, Gaehtgens P. (1991) Endothelial, not hemodynamic, differences are responsible for preferential leukocyte rolling in rat mesenteric venules. Circ Res 69, 1034–41. 13. von Andrian UH. (1996) Intravital microscopy of the peripheral lymph node microcirculation in mice. Microcirculation 3, 287–300. 14. Von Adrian U.H, M’Rini C. (1998) In situ analysis of lymphocyte migration to lymph nodes. Cell Adhes Commun 6, 85–96.

SUBJECT INDEX Note: The letters ‘f ’ and ‘t’ following the locators refer to figures and tables respectively

R . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 Anti-PE EasySep Anti-TCRζ -PE-conjugated antibodies . . . . . . . . . . . . . . . 262 Anti-TNF therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263–264 APC, see Antigen presenting cells (APC) Atherogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Atherosclerosis and endothelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 atherosclerotic lesions . . . . . . . . . . . . . . . . . . . . . . . . . 33 “native” atherosclerosis . . . . . . . . . . . . . . . . . . . . . . . . 33 or multiple sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 Autoimmune diseases . . . . . . . . . . . . . . . . . 126–127, 194, 231 “Automated” adhesion assay . . . . . . . . . . . . . . . . . . . . . . . . . 250 Automatic cell quantification . . . . . . . . . . . . . . . . . . . . . . . . 208

A Adhesion . . . . . . . . . . . . . . . 5, 7–8, 18–23, 27–28, 31–45, 50, 52, 59–60, 62, 64–66, 69, 73–74, 80–81, 83, 104, 125, 219, 232–235, 239–246, 248–250, 277f, 278 molecules. . . . . .4–6, 8–9, 11, 45, 85, 99–100, 243, 268, 271–272, 275 L-selectin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 MAdCAM-1 or ICAM-1 . . . . . . . . . . . . . . . . . . . . . . 5 P- and E-selectin or VCAM-1 . . . . . . . . . . . . . . . . . . 5 VAP-1/CD44 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 transmigration assay systems . . . . . . . . . . . . . . . . 234–236 cell culture inserts . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 chemoattractant solution . . . . . . . . . . . . . . . . . . . . . 236 collagen solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 cytokine solutions . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 flow-based adhesion assay system . . . . . . . . . . . . . . . . . . . . . . . . 235f, 243–244 measurement of lymphocyte transmigration . . . . . . . . . . . . . . . . . . . . . . . . . . 246f static adhesion assay . . . . . . . . . . . . . . . 244–246, 245f T-cell chemotaxis . . . . . . . . . . . . . . . . . . . . . . . 246–247 VCAM-1 solution . . . . . . . . . . . . . . . . . . . . . . . . . . . 234 Affi-gel Blue Gel (Bio-Rad) . . . . . . . . . . . . . . . . . . . . 163, 172 Anesthesia . . . . . . . . . . . . . . . . . . . . . . 183, 185, 191, 269, 273 Antibody . . . 20, 25, 29, 41, 73–75, 81, 86, 88, 91–92, 101, 103–104, 106, 107f, 108f, 110, 117–118, 129, 135, 136f, 140, 142, 153, 156, 175, 210, 212, 232–234, 240f, 241, 243, 246, 255, 257, 261–262, 268–269, 275 Anti-coagulants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 Antigen . . . . . . . . 3, 5–9, 32, 35, 42, 99, 101, 115, 118, 121, 139–140, 149, 161, 182, 185, 194–195, 202f, 203, 210–212, 217, 220, 232, 239, 253–265 Antigen-experienced effector T cells in vitro and in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253–265 materials flow cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 HUVECs preparation . . . . . . . . . . . . . . . . . . . 255–256 PBMCs by density-gradient centrifugation . . . . . . . . . . . . . . . . . . . . . . . 254–255 transendothelial migration assay . . . . . . . . . . . . . . 256 methods flow cytometry . . . . . . . . . . . . . . . . . . . . . . . . . 257–258 HUVECs preparation . . . . . . . . . . . . . . . . . . . 258–259 PBMCs by density-gradient centrifugation . . . . . . . . . . . . . . . . . . . . . . . 256–257 transendothelial migration assay . . . . . . . . . . 259–260 Antigen presenting cells (APC) . . . 32, 35, 42, 43f, 99–100, 103, 139–140, 232, 255, 257

B BABB, see Benzyl Alcohol and Benzyl Benzoate (BABB) BAC, see Bacterial artificial chromosome (BAC) technology Bacterial artificial chromosome (BAC) technology . . . . . 164 BBB, see Blood-brain barrier (BBB) BEC, see Biliary epithelial cells (BEC) Becton Dickinson (BD) FACSCanto instrument . . . . . . . . . . . . . . . . . . . . . . . . . 134 FACSDiva software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Benzyl Alcohol and Benzyl Benzoate (BABB) . . . . . . . . 164, 175–177 Biliary epithelial cells (BEC) . . . . . . . . . . . . . . . . . . . . . . . . 243 “Biosensor,” 98, 100f, 106, 107f Blood-brain barrier (BBB) . . . . . . . . . . . . . . . . . . 21, 268–269 Brachiocephalic artery . . . . . . . . . . . . . . . . 199–200, 200f, 204 Bright-field or fluorescence microscope . . . . . . . . . . . . . . . 208

C Calcein AM/CellTiter-Glo reagent . . . . . . . . . . . . . . . . . . 130 Calcium signalling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 “Cameleon” calcium probe . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 CAMs high-affinity (integrins) . . . . . . . . . . . . . . . . . . . . . . . . . . 83 low-affinity (L-selectin) . . . . . . . . . . . . . . . . . . . . . . . . . . 83 5(6)-carboxyfluorescein diacetate N -succinimidyl ester (CFSE) . . . . . . . . . . 35, 37, 42–43, 196–197, 204 Cardiogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 Catchment basins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209, 216 CAV, see Chronic allograft vasculopathy (CAV) Cdc42 . . . . . . . 85–86, 89, 93f, 99, 100f, 102, 106–107, 108f “Raichu” biosensor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100f Cells endothelium . . . 5–8, 17–29, 31–33, 43, 53f, 60, 64, 69, 73–74, 76–77, 149, 233, 235f, 241, 243–244, 249–250, 254, 268, 274–276, 277f, 278–279

F.M. Marelli-Berg, S. Nourshargh (eds.), T-Cell Trafficking, Methods in Molecular Biology 616, c Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-60761-461-6, 

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282 Subject Index

Cells (continued) human T cells . . . . . . . . . . . . . . . 268–272, 274–275, 277f leukocyte . . . . . . . . 17–29, 115–123, 231, 233, 239, 242, 244–245, 249, 268, 275, 276f lymphocyte . . . . . . . . . 8, 49–67, 116, 211, 233, 236–239 stromal cells . . . . . . . . . . . . . . . . . . . . . . . . . . . 4, 49–67, 168 surrogate target cell . . . . . . . . . . . . . . . . . . . . 182, 185, 188 Cell–cell junctional remodelling, diapedesis . . . . . . . . . . . . 81 Cell counting equipment counter and haemocytometer. . . . . . . . . . . . . . . . . . . . . .85 phase contrast light microscope . . . . . . . . . . . . . . . . . . . 85 Cell culture . . . . . . . . . 36, 39, 51, 65–66, 70, 72, 85, 88–89, 101–102, 139, 150, 235–236, 243, 246–247, 273 Cell disruption techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 Cell migration . . . . . . . . . . . . . . . . . . 119, 130, 137, 143, 145, 170, 195, 202f, 247 Cell signaling technology . . . . . . . . . . . . . . . . . . . . . . . . . . . 129 CellTiter-Glo dye . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 Central nervous system (CNS) . . . . . . . . . 222, 267–269, 275 gray matter/white matter . . . . . . . . . . . . . . . . . . . . . . . . 269 microvasculature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 CFP, see Cyan fluorescent protein (CFP) CFSE, see 5(6)-Carboxyfluorescein diacetate N -succinimidyl ester (CFSE) “Checkerboard” analysis matrix . . . . . . . . . . . . . . . . . . . . . . 118 Chemokine(s) . . . . . . . . . 4, 6, 9, 18, 27, 115–117, 119–121, 125–126, 129–130, 137, 236, 247, 250–251, 256, 260, 265, 268 receptors . . . . . . 4, 8–11, 84, 98, 125–126, 134–135, 250 Chemokine-dependent chemotaxis . . . . . . . . . . . . . . . . . . 118 Chemotaxis . . . . . . . . . . . . . . . . . . . . . . . 4, 115–123, 125, 130, 137–138, 143–145, 246 R System . . . . . . . . . . . . . . . . . . 129–130, 137, 143 ChemoTx Chronic allograft vasculopathy (CAV) . . . . . . . . . . . . . . . . . 33 CLA, see Cutaneous lymphocyte antigen (CLA) CLIO- . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121, 123 CNS, see Central nervous system (CNS) Co-culture . . . . . . . 40–44, 50, 58–59, 65–66, 70, 73–74, 77 Cohu 4912 monochrome camera . . . . . . . . . . . . . . . . . . . . . 55 Collagenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150, 152 Collagenase (type IA) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 Confocal laser scanning microscopy . . . . . . . . . . . . . . . . . . 176 Confocal microscopy . . . . . . . . . . . . . . . 20f, 22f, 23f, 24f, 41, 70, 75, 104, 122, 167, 173f, 176 Conventional wide-field microscopy . . . . . . . . . . . . . . . . . 167 Cooled benchtop centrifuge R . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 Beckman Coulter Coomassie Blue staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 Coulter counter or haemocytometer . . . . . . . . . . . . . . . . . . . 60 Custom-built perfusion system . . . . . . . . . . . . . . . . . . . . . . 200 Custom-made Mikron IVM500 epi-fluorescent microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Cutaneous lymphocyte antigen (CLA) . . . . . . . . . . . . . . . . . 8 CXCL12-induced chemorepulsion. . . . . . . . . . . . . . . . . . .116 CXCR3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 125–145 ligands CXCL9/CXCL10/CXCL11 . . . . . . . . . . . . . . . . . 126 CXCR3-B and CXCR3-alt, variants . . . . . . . . . . 127 PF4/CXCL4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127 Cyan fluorescent protein (CFP) . . . . . . . . . . . . . . . . . . . 24, 98 Cytokines . . . . . . . . 6, 8, 10, 18, 27, 33, 50, 59, 66, 79, 235f, 243–244, 246, 254 Cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99, 104 Cytotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181–192

Cytotoxic T cell function by (MP-IVM), single-cell analysis of . . . . . . . . . . . . . . . . . . . . . . . . . . 181–192 materials antigen-specific anti-tumor CD8+ T cell response, induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 equipment/procedure of popLN preparation for MP-IVM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184f mouse popLN, intravital microscopy . . . . . . . . . . 183 mouse popLN, MP-IVM . . . . . . . . . . . . . . . 183–185 surgical anesthesia . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 surrogate CTL target cells, preparation/injection . . . . . . . . . . . . . . . . . . . . . 182 surrogate target cell death, image processing/quantitative analysis . . . . . . . . . . . 185 methods antigen-specific anti-tumor CD8+ T cell response, induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 monitoring target cell apoptosis by MP-IVM . 189f mouse popLN, intravital microscopy . . . . . . 186–187 mouse popLN, MP-IVM . . . . . . . . . . . . . . . 187–188 surgical anesthesia . . . . . . . . . . . . . . . . . . . . . . 185–186 surrogate CTL target cells, preparation/injection . . . . . . . . . . . . . . . . . . . . . 185 target cell death, image processing/quantitative analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 188–190 Cytotoxic T lymphocytes (CTL) . . . . . . . . . . . . . . . 182, 185, 188–189, 194

D DAPI, see Diamidino-2-phenylindole (DAPI) Data analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275–278 hemodynamic parameters . . . . . . . . . . . . . . . . . . . . . . . . 275 quantitative analysis of firm adhesion . . . . . . . . . . . . . 278 quantitative analysis of T-cells . . . . . . . . . . . . . . . 276–278 firm adhesion, IVM analysis of. . . . . . . . . . . . . . .277f parabolic flow profile within blood vessel . . . . . . 276f DCs, see Dendritic cells (DCs) Deep-tissue imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Dendritic cells (DCs) . . . . . . . . . . . . . . 5, 116, 193–194, 194, 202f, 232–233, 238–239, 247–248 Diamidino-2-phenylindole (DAPI) . . . . . . . . . . . 35, 41, 45f, 210–213, 217, 218f, 223 Diapedesis, paracellular/transcellular routes discrimination . . . . . . . . . . . . . . . . . . . . . . . . . 69–81 endothelial receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 leucocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79–80 materials cell lines/culture reagents . . . . . . . . . . . . . . . . . . 70–71 immunofluorescence. . . . . . . . . . . . . . . . . . . . . . .71–72 methods EC, cell culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 immunofluorescence assay . . . . . . . . . . . . . . . . . 73–75 T lymphoblasts, cell culture . . . . . . . . . . . . . . . . . . . 72 T-lymphoblast transmigration, see T-lymphoblast transmigration, detection transmigration assay, see Transmigration assay in diapedesis occurrences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 PECAM-1 or CD99, surface receptors . . . . . . . . . . . . 70 thawing endothelial cells . . . . . . . . . . . . . . . . . . . . . . . . . . 79 TNF-α stimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 transmigration process. . . . . . . . . . . . . . . . . . . . . . . . . . . .70 in vitro confocal microscopy studies . . . . . . . . . . . . . . . . 70 in vivo electron microscopy studies . . . . . . . . . . . . . . . . 70 Digital thermometer . . . . . . . . . . . . . . . . . . . . . . . . . . . 185, 187

T-CELL TRAFFICKING Subject Index 283 Dimethyl sulfoxide (DMSO) . . . . . . . . . . . . . . . . . 51, 58, 79, 101, 196, 255, 270 DMSO, see Dimethyl sulfoxide (DMSO) Donkey anti-goat-Cy5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 Donkey anti-mouse-FITC. . . . . . . . . . . . . . . . . . . . . . . .72, 74 Donkey anti-rabbit-Cy5 . . . . . . . . . . . . . . . . . . . . . . . . . . 72, 74 R . . . . . . . . . . . 64, 128, 140, 233–234, 239, 243 Dynabeads

Epigenetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Epi-illumination techniques . . . . . . . . . . . . . . . . . . . . . . . . . 274 Ethylenediaminetetraacetic acid (EDTA) solution . . 34–36, 50–51, 55, 57–58, 71–72, 86, 128, 141, 150, 152–154, 256, 259, 278 Experimental autoimmune encephalomyelitis (EAE) 267–270, 273, 276, 277f Extra cellular matrix (ECM) . . . . . . . . . . . . . . . . . . . . . . . . . 72

E

F

EAE, see Experimental autoimmune encephalomyelitis (EAE) EC, see Endothelial cells (EC) ECGS, see Endothelial cell growth supplement (ECGS) ECM, see Extra cellular matrix (ECM) EC/stromal cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55–56 dispersal of monolayers for passaging . . . . . . . . . . . 57–58 freezing stromal cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 isolation and culture of HUVEC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56 primary fibroblasts . . . . . . . . . . . . . . . . . . . . . . . . 56–57 primary “secretory” SMC from umbilical artery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57 EDTA, see Ethylenediaminetetraacetic acid (EDTA) solution EGFP, see Enhanced green fluorescent protein (EGFP/eGFP) Electron microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70, 220 Embryo dissections, developing lymphoid structures . . . . . . . . . . . . . . . . . . . . . . . . . . . 168–173 explant organ cultures, migration assays in . . . 170–173, 170f–171f sample preparation . . . . . . . . . . . . . . . . . . . . . . . . . 168–169 time-lapse microscopy . . . . . . . . . . . . . . . . . . . . . . 169–170 Embryonic tissue for expression analysis of transcription factors . . . . . . 211 antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 preparation of tissue samples . . . . . . . . . . . . . . . . . 211 sections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 in gene expression studies . . . . . . . . . . . . . . . . . . . 210–211 DAPI staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 LCS software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 optical sections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 secondary antibodies for immunofluorescence . . 210 Emigration process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Endothelial-based transmigration assays . . . . . . . . . . . . . . 250 Endothelial cells (EC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 –stromal cell co-cultures establishing on filters . . . . . . . . . . . . . . . . . . . . . . 58–59 and stromal cell culture for assays cell culture inserts . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 in vitro, uses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33–34 HUVEC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 isolation and purification . . . . . . . . . . . . . . . . . . . . . . 33 Endothelial cell growth supplement (ECGS) . . . . . 71, 151, 256, 258 Endothelial isolation from human liver tissue . . . . . . . . . 234 collagenase solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234 complete endothelial culture medium . . . . . . . . . . . . . 234 nycodenz solution. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .234 trypsin solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234 Endothelium . . . . . . 5–8, 17–29, 31–33, 43, 53f, 60, 64, 69, 73–74, 76–77, 149, 233, 235f, 241, 243–244, 249–250, 254, 268, 274–276, 277f, 278–279 Enhanced green fluorescent protein (EGFP/eGFP) . . . 182

F-actin, see Filamentous actin (F-actin) Fate-reporter systems . . . . . . . . . . . . . . . . . . . . . . . . . . 165, 167 FBS, see Foetal bovine serum (FBS) FCS, see Foetal calf serum (FCS) Fibroblasts . . . . . . . . . . . . . . . . . . . . . . 5, 49–51, 55–59, 65–66 Fibronectin (FN) . . . 8, 34, 36, 44, 65, 71–73, 79, 101, 103, 117, 236, 246, 249 Field of view (FOV) . . . . . . . . . 168, 188, 190, 225, 275, 278 Filamentous actin (F-actin) . . . . . . . . 41, 70, 74, 80–81, 106 FITC, see Fluorescein isothiocyanate (FITC) FLIM, see Fluorescence lifetime imaging microscopy (FLIM) Flow-based assay of lymphocyte adhesion/migration . . . . 51 assembling and connecting flow chamber for fluorescence microscopy . . . . . . . 61 phase contrast flow chamber . . . . . . . . . . . . . . . 61–62 cell behaviour analysis. . . . . . . . . . . . . . . . . . . . . . . . .63–64 circular phase-bright cells . . . . . . . . . . . . . . . . . . . . . 63 phase-bright cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 phase-dark spread cells . . . . . . . . . . . . . . . . . . . . . . . . 63 flow system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54, 54f fluorescence parallel-plate chamber . . . . . . . . . 52–53, 52f image analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 perfusing cellular suspension/recording behaviour . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62–63 phase contrast parallel-plate chamber . . . . . . . 53–54, 53f setting up flow assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 video microscope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54–55 Flow cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . . 255, 257–258 cell surface staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 of PBMCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 derivation of the TCRζ expression index . . . . . . . . . . 258 FACS-based expression index . . . . . . . . . . . . . . . . 258f intracellular staining of PBMCs for TCRζ expression . . . . . . . . . . . . . . 255 for TCRζ expression . . . . . . . . . . . . . . . . . . . . . . . . 257 or fluorescence microscopy . . . . . . . . . . . . . . . . . . . . . . . 167 Fluid shear stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32–33 Fluorescein isothiocyanate (FITC) . . . . . . . . . . . . . . . . . . . 129 FITC-conjugated antibody . . . . . . . . . . . . . . . . . . . . . . 262 Fluorescence lifetime imaging microscopy (FLIM) . . . . . . . 98–99, 100f, 104–105, 107, 109 and confocal microscopy . . . . . . . . . . . . . . . . . . . . . . . . . 104 antibody staining, sample preparation . . . . . . . . . 104 antibody staining, sample preparation that do not require . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Fluorescence microscopy . . . . . . . . . . 43, 50, 52, 59f, 61, 167 Fluorescence parallel-plate chamber . . . . . . . . . . . 52–53, 52f Fluorescent probes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20, 25–26 proteins (FP) . . . 25–26, 29, 98–99, 109, 162, 164–165, 175–176, 182 Fluorescently tagged fusion proteins. . . . . . . . . . . . . . . . . . .19 time-lapse fluorescence confocal microscopy . . . . . . . 24f transfection of endothelial cells . . . . . . . . . . . . . 19, 24–25 transfection of leukocytes . . . . . . . . . . . . . . . . . . . . . 19, 25

T-CELL TRAFFICKING

284 Subject Index

Fluorochrome . . . . . . . . . . . . . . . . . . . . 37, 142, 174, 176, 204 Foetal bovine serum (FBS) . . 18–19, 21, 23–24, 34, 36–37, 40, 70–71, 80, 101, 117, 129, 131, 139, 142, 150, 152, 169, 185 Foetal calf serum (FCS). . . . . . . . .34–35, 51, 58, 65–66, 85, 88, 103, 117, 150, 163, 169, 172, 196–197, 203, 233, 242, 246, 255–256, 258–259, 272–273 F¨orster resonance energy transfer (FRET) . . . . . . . . 98–100, 106–107, 109–110, 192 FOV, see Field of view (FOV) FP, see Fluorescent Fugetaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115–123 Full-width halfmaximum (FWHM) . . . . . . . . . . . . . . . . . 216 FWHM, see Full-width halfmaximum (FWHM)

G GAPs, see GTPase-activating proteins (GAPs) GDP, see Guanosine diphosphate (GDP) GEFs, see Guanosine nucleotide exchange factors (GEFs) Gene transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 GFL, see Glial-derived neurotrophic factor ligand (GFL) GFP, see Green fluorescent protein (GFP) Glial-derived neurotrophic factor ligand (GFL) . . . . . . . 170 ?λυτ ατ ηιoνεttτρανσ φερασ ε (GST) . . . . 85–86, 88–91 GPCR, see G-protein-coupled receptors (GPCR) G-protein-coupled receptors (GPCR) . . . . . . . . . 9, 125–126 R /ImageJ R . . . . . . . . . . . . . . . . . . . . . . . 88 GraphPad Prism Green fluorescent protein (GFP) . . . . . . . . . 98–99, 165, 182 GST, see Glutathione-S−transferase (GST) GTP, see Guanosine triphosphate (GTP) GTPase-activating proteins (GAPs) . . . . . . . . . . . . . . . 84f, 88 Guanosine diphosphate (GDP) . . . . . . . . . . . . . . . . . . . 84–85 Guanosine nucleotide exchange factors (GEFs) . . . . . 84–85 Guanosine triphosphate (GTP) . . . . . . . . . . . 84–85, 88, 93f, 94, 98–99, 100f, 106

H H1077, see Histopaque 1077 (H1077) HA, see Hemagglutinin (HA) Hagen–Poiseuille law . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Hank’s balanced salt solution (HBSS) . . . . . . . . . . . . . . . . 270 Hanks buffered saline solution (HBSS) . . . . . . . . . . . . . . . 254 Harvard Apparatus . . . . . . . . . . . . . . . . . . . . . . . . . . . 19–20, 54 HBSS, see Hank’s balanced salt solution (HBSS); Hanks buffered saline solution (HBSS) HDMVECs, see Human dermal microvascular endothelial cells (HDMVECs) Hemagglutinin (HA) . . . . . . . . . . . . . . . . . . . . . . . . . . . 71, 182 HEV, see High endothelial venules (HEV) High endothelial venules (HEV) . . . . . . . . . . . . . 4, 188, 191 Histopaque 1077 (H1077) . . . . . . . . . . . . . . . . . . . 50, 55, 121 HLA-DR staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 Homing . . . . . . . . . . . . . . . . . . . . . 4, 7–11, 126, 231–251, 269 Homing interactions in human disease . . . . . . . . . . . 231–251 materials adhesion/transmigration assay systems . . . . 234–236 endothelial isolation from human liver tissue . . . 234 lymphocytes/DCs, isolation of . . . . . . . . . . . . . . . . 233 Stamper–Woodruff adhesion assay . . . . . . . 233–234 methods adhesion/transmigration assay systems . . . . 243–247 lymphocytes/DCs, isolation of . . . . . . . . . . . 236–239 Stamper-Woodruff adhesion assay . . . . . . . . 239–242 tissue-specific endothelial adhesion assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 242–243

Hox-induced cell clustering . . . . . . . . . . . . . . . . . . . . . . . . 219f HSVEC, see Human saphenous vein endothelial cells (HSVEC) Human dermal microvascular endothelial cells (HDMVECs), 70–71, 77, 78f Human saphenous vein endothelial cells (HSVEC) . . . 151, 154 Human T cells . . . . . . . . . . . . . . . . . . 268–272, 274–275, 277f Human T lymphocytes, analysis of CXCR3 in . . . 125–145 materials intracellular proteins, phosphorylation levels of . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129–130 PCR primers, CXCR3 variants . . . . . . . . . . . . . . 129t receptor internalisation by flow cytometry. . . . . .130 RNA extraction/reverse transcription/PCR analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128 T-cell isolation/activation/ex vivo expansion . . . . . . . . . . . . . . . . . . . . . . . . . . . 127–128 T-cell markers and receptors, expression of . . . . . 129 in vitro cell migration assay . . . . . . . . . . . . . . . . . . . 130 methods agonist-induced receptor internalisation by flow cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . 135–137 flow cytometry analysis . . . . . . . . . . . . . . . . . . 133–134 internalisation of CXCR3 . . . . . . . . . . . . . . . . . . . 136f intracellular S6 ribosomal phosphorylation . . . . . . . . . . . . . . . . . . . . . 134–135 isolation/ex vivo activation/expansion . . . . . 130–131 migratory responses . . . . . . . . . . . . . . . . . . . . . . . . . 138f omniscript RT Kit. . . . . . . . . . . . . . . . . . . . . . . . . . .132 PBMC separation . . . . . . . . . . . . . . . . . . . . . . . . . . 131f PCR analysis of mRNA expression . . . . . . . . . . . 133f RNA extraction/reverse transcription/PCR analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131–133 in vitro directional cell migration assays . . . 137–139 Human umbilical vein endothelial cells (HUVEC) . . . . . 34, 44–45, 70 preparation of . . . . . . . . . . . . . . . . . . . . 255–256, 258–259 HUVEC, see Human umbilical vein endothelial cells (HUVEC)

I ICAM-1 . . . . . . . . . . . . . . . . 5, 18, 21, 26–27, 69–71, 73–75, 77, 78f, 79–81 Fluorescein isothiocyanate (FITC) . . . . . . . . . . . . . . . . 75 IDO, see Indoleamine 2,3 dioxygenase (IDO) IDO in primary ECs . . . . . . . . . . . . . . . . . . . . . . . . . . 149–155 materials cell culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150–151 IDO activity assay . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 isolation/purification of HSVEC . . . . . . . . . . . . . 150 transfection of ECs . . . . . . . . . . . . . . . . . . . . 151, 154f methods IDO activity assay . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 isolation of HSVEC . . . . . . . . . . . . . . . . . . . . 152–154 transfection of ECs using electroporation . . . . . . . . . . . . . . . . . . . . . . 154–155 IFNtγ , see Interferon-γ g(IFNtγ p Image-Pro software (Media Cybernetics) . . . . . . . . . . . . . . 55 Image segmentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 Imaging confocal laser scanning microscopy . . . . . . . . . . . . . . . 176 confocal microscopy . . . . . . 20f, 22f, 23f, 24f, 41, 70, 75, 104, 122, 167, 173f, 176 electron microscopy . . . . . . . . . . . . . . . . . . . . . . . . . 70, 220 fluorescence microscopy . . . . . . . 43, 50, 52, 59f, 61, 167

T-CELL TRAFFICKING Subject Index 285 image segmentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 intravital microscopy . . . . . . . . . . . . . . . . . . 118, 121, 181, 183, 186–187, 269 live cell imaging. . . . . . . . . . . . . . . . . . .164–165, 167–168 multiphoton intravital microscopy . . . . . . . 181, 183, 187 phase contrast microscopy . . . . . . 40, 43, 50, 53, 57, 244 stereo fluorescent microscopy . . . . . . . . . . . . . . . . . . . . 176 time lapse microscopy . . . . . . . . . . . . . . . . . . . . . . . 99, 169 watershed segmentation . . . . . . . . . . . . . . . . 209, 213, 216 IMDM, see Iscove’s Modified Dulbecco’s Medium (IMDM) Immune/cardiovascular systems by multiphoton microscopy, 193–203 materials CFSE/CMTPX stock . . . . . . . . . . . . . . . . . . . . . . . 196 media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196 mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195–196 other items . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196 methods adoptive transfer, preparation for . . . . . . . . . . . . . . 197 analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201–203 brachiocephalic artery for imaging preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199 dissection of brachiocephalic artery . . . . . . . . . . . 200f ex vivo imaging of LN, procedure for . . . . . . . . . 198f LN for imaging preparation. . . . . . . . . . . . . .197–199 lymph nodes removal . . . . . . . . . . . . . . . . . . . . 196–197 microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 perfusion system . . . . . . . . . . . . . . . . . . . . . . . . . . . . 200 T cell migration/interactions in LN, analysis of . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202f Immune-receptor tyrosine-based activation motifs (ITAMs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 Immunofluorescence assay (triple staining) . . . . . . . . . 73–75 βtcatenin staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 double fluorescence staining. . . . . . . . . . . . . . . . . . . . . . .73 fluorophore-conjugated phalloidin staining . . . . . . . . . 74 ICAM-1 staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 paracellular from transcellular, parameters . . . . . . . 74–75 Immunofluorescence procedure . . . . . . . . . . . . . . . . . . . . . . . 73 Immunohistochemistry . . . . . . . . . . . . . . 35, 40–41, 163, 167 Immunomagnetic cell selection techniques . . . . . . . . . . . . 185 Imprinting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9–11, 239 Incubation system (La-con GBr Pe-con GmbH) . . . . 20, 26 Indoleamine 2,3 dioxygenase (IDO) . . . . . . . . . . 6, 149–155, 154–156 Inflammation . . . . . . . . . . 5–8, 32, 73, 79–80, 123, 126, 137, 149–150, 232, 236, 247, 254, 267, 269 Inflammatory chemokines . . . . . . . . . . . . . . . . . . . . . . . . . . 126 Inflammatory or autoimmune diseases, lymphocytes in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 monoclonal antibody therapies . . . . . . . . . . . . . . . . . . . 232 T-cell and dendritic cell populations . . . . . . . . . . . . . . 232 tissue damage, consequences . . . . . . . . . . . . . . . . . . . . . 231 A4 β7 integrin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7–8, 232 Interactions cell:cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108, 194 protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 Intercellular adhesion molecule-1 (ICAM-1) . . . . 5, 18, 21, 26–27, 69–71, 73–75, 77, 78f, 79–81 Interferon-γ g(IFNtγ p, 6, 10, 51, 66, 79, 126, 210, 212, 235, 243–244 Interleukin-2 (IL-2) 18, 21, 32, 71–72, 128, 131, 133, 140, 143, 248 Intravital fluorescence videomicroscopy (IVM) experiments 269–271, 274

Intravital microscopy . . . . 118, 121, 181, 183, 186–187, 269 in vivo imaging of immunity in atherosclerosis, see Immune/cardiovascular systems by multiphoton microscopy IPLab software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Iscove’s Modified Dulbecco’s Medium (IMDM) . . . . . . 163, 168–169, 174 ITAMs, see Immune-receptor tyrosine-based activation motifs (ITAMs) IVM, see Intravital fluorescence videomicroscopy (IVM) experiments

J Junctions . . . . . . . . . . . . . . . . . . . . . . . . . 18, 70, 74, 79–81, 249

K Kruskal–Wallis test . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278

L Laminectomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274 Laminin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101, 103, 117 LCR, see Locus control region (LCR) Leucocyte . . 5–6, 31–45, 49–50, 54f, 55, 65, 69–70, 72–74, 79–81, 85, 115–123, 137, 149 trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 transmigration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 Leucocyte adhesion under haemodynamic flow conditions apparatus, uses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42–43 analysis of leucocyte adhesion . . . . . . . . . . . . . . . . . . 43 interposition of step barrier creates disturbed flow downstream . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44f silicone steps to simulate separated flows . . . . 43–44 atherosclerosis and endothelium . . . . . . . . . . . . . . . . . . . 33 EC in vitro, uses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33–34 endothelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31–32 HUVEC culture in static . . . . . . . . . . . . . . . . . . . . . . . . 45f materials flow loop . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 HUVEC culture on glass slides . . . . . . . . . . . . . . . . 34 immunohistochemistry . . . . . . . . . . . . . . . . . . . . . . . . 35 PHA proliferation assay . . . . . . . . . . . . . . . . . . . 35–36 T-cell alloproliferation assay . . . . . . . . . . . . . . . . . . . 35 T-cell purification . . . . . . . . . . . . . . . . . . . . . . . . . 34–35 methods HUVEC culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36 T-cell purification . . . . . . . . . . . . . . . . . . . . . . . . . 36–37 use of parallel-plate flow chamber, see Parallel-plate flow chamber parallel-plate flow chamber, uses. . . . . . . . . . . . . . . . . . .34 shear stress. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .32–33 flow patterns for laminar/disturbed flow . . . . . . . . 32f Leukocyte . . . . . . . . . . . 17–29, 115–123, 231, 233, 239, 242, 244–245, 249, 268, 275, 276f Leukocyte chemotaxis and fugetaxis, quantitation methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115–123 materials intraperitoneal inflammatory infiltration of mice118 linear and complex gradients of chemokines . . . . 117 T cells to cell-secreted CXCL12, migrational response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 transmigration assay . . . . . . . . . . . . . . . . . . . . . . . . . 117 methods intraperitoneal inflammatory infiltration of mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

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Leukocyte chemotaxis (continued) intravital microscopy of rat mesentery . . . . . 121–122 linear and complex gradients of chemokines . . . . . . . . . . . . . . . . . . . . . . . . . 119–120 MRI tracking of antigen-specific T cells . . . . . . . 121 T cells to cell-secreted CXCL12, migrational response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 transmigration assay . . . . . . . . . . . . . . . . . . . . . . . . . 118 Leukocyte-endothelium interactions, live imaging of . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17–29 extravasation process, adhesion steps . . . . . . . . . . . 17–18 leukocyte transendothelial migration . . . . . . . . . . . . . . . 17 materials cell models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 detachment experiments . . . . . . . . . . . . . . . . . . . . . . 19 under flow conditions . . . . . . . . . . . . . . . . . . . . . . . . . 19 fluorescently tagged fusion proteins . . . . . . . . . . . . 19 staining of living cells . . . . . . . . . . . . . . . . . . . . . . . . . 20 static adhesion measurements and staining . . . . . . 19 time-lapse fluorescence microscopy . . . . . . . . . . . . . 20 methods cell models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 detachment experiments . . . . . . . . . . . . . . . . . . . 22–23 under flow conditions . . . . . . . . . . . . . . . . . . . . . 23–24 fluorescently tagged fusion proteins . . . . . . . . . . . . 24 staining of living cells . . . . . . . . . . . . . . . . . . . . . . . . . 25 static adhesion measurements and staining . . . . . . 21 time-lapse fluorescence microscopy . . . . . . . . . . . . . 26 Ligand-dependent activation assay for live imaging Jurkat-Raji conjugate formation . . . . . . . . . . . . . . . . . . 103 YTS:721.221 conjugate formation . . . . . . . . . . . . . . . . 104 Live cell imaging . . . . . . . . . . . . . . . . . . . . . 164–165, 167–168 Liver . . . . . . 7, 161–162, 165, 232–234, 236, 237f, 238, 240, 242–243, 247–248 LMSW, see Local maximum seeded watershed (LMSW) LN, see Lymph node (LN) Local maximum detection Illustration . . . . . . . . . . . . . . . 215f Local maximum seeded watershed (LMSW) . . . . . . . . . 209 approaches and manual, comparison . . . . . . . . . . . . . 218f procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214f Locus control region (LCR) . . . . . . . . . . . . . . . . . . . . 164–165 L-selectin . . . . . . . . . . . . . . . . . . . . . 4, 7–8, 83–86, 88–89, 93f LTis, see Lymphoid tissue “inducer” cells (LTis) L-tryptophan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149, 155, 157 Lymph node (LN) . . . . . . . . . . . . 5, 7–8, 108, 126, 162, 172, 182–183, 186–187, 194–195, 203, 238 atherogenic T cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 DC:T cell interactions . . . . . . . . . . . . . . . . . . . . . . . . . . 194 imaging immune cell behaviour . . . . . . . . . . . . . . . . . . 195 See also Immune/cardiovascular systems in vivo by multiphoton microscopy Lymphocyte adhesion/migration on EC . . . . . . . . . . . 49–67 materials blood cell isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 culture of EC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 culture of stromal cells . . . . . . . . . . . . . . . . . . . . . . . . 51 endothelial/stromal cell culture for assays . . . . . . . 51 flow-based adhesion assay . . . . . . . . . . . . . . . . . . 52–55 methods co-cultures on 3.0-μm pore filters under static conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59–60 endothelial–stromal cell co-cultures . . . . . . . . . 58–59 flow-based assay, see Flow-based assay of lymphocyte adhesion/migration for isolating lymphocytes . . . . . . . . . . . . . . . . . . 64–67

isolation and culture of EC/stromal cells, see EC/stromal cells leucocyte isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Lymphocytes/DCs, isolation of . . . . . . . . . . . . 233, 236–239 cell isolation from liver homogenate . . . . . . . . . . . . . . 238 culture medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 downstream use of DCs . . . . . . . . . . . . . . . . . . . . . . . . . 239 isolation of DCs from liver homogenate . . . . . . . . . . . . . . . . . . . . . . . 238 from lymph node . . . . . . . . . . . . . . . . . . . . . . . 238–239 from skin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238 from whole liver tissue . . . . . . . . . . . . . . . . . . . . . . . 238 isolation of lymphocytes from human liver specimens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237f OptiprepTM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 purification of DCs negative selection by depletion of CD3 lymphocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 positive selection of CD11c+ DCs . . . . . . . . . . . . 239 sample collection and preparation . . . . . . . . . . . . . . . . 236 tissue storage medium . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 wash buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Lymphocyte trafficking . . . . . . . . . . . . . . . . . . . . . . 8, 116, 211 Lymphoid organ development, visualisation of . . . 161–178 live cell imaging embryo dissections . . . . . . . . . . . . . . . . . . . . . . 168–173 microscope set-ups . . . . . . . . . . . . . . . . . . . . . . 167–168 materials cell/organ culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 immunohistochemistry . . . . . . . . . . . . . . . . . . 163–164 reporter mouse strains cell fate reporter mouse models . . . . . . . . . . . . . . . 167 direct expression of FP . . . . . . . . 165–166, 166f–167f whole-mount staining image acquisition . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176 sample optical clearing . . . . . . . . . . . . . . . . . . 175–176 tissue preparation and sectioning . . . . . . . . . . . . . . 174 Lymphoid organogenesis . . . . . 162, 164–165, 167–170, 173 Lymphoid system. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .161–162 haematopoietic inducer cells . . . . . . . . . . . . . . . . . . . . . 162 categories . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 LTis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 primary lymphoid organs haematopoietic cells . . . . . . . . . . . . . . . . . . . . . . . . . 162 NCCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 primary lymphoid organs . . . . . . . . . . . . . . . . 161–162 T and B cell differentiation . . . . . . . . . . . . . . . . . . . 161 secondary lymphoid organs . . . . . . . . . . . . . . . . . . . . . . 162 foetal lymphoid organogenesis . . . . . . . . . . . . . . . . 162 LN and PP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 Lymphoid tissue “inducer” cells (LTis) . . . . . . . . . . . . . . . 162 Lymphoprep . . . . . . . . . . . . . . . . . . . . 127, 130–131, 139, 254

M Major histocompatibility (MHC) . . . 32, 45, 101, 139–140, 182, 210, 212 Mann–Whitney U -test . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 Manual counting. . . . . . . . . . . . . . . . . . . . . . . . . . 208, 217–218 MATLAB 13 software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 MCIs, see Mean chemotropic indices (MCIs) Mean chemotropic indices (MCIs) . . . . . . . . . . . . . . . . . . 119 Memory cells . . . . . . . 5, 7–8, 10, 32, 66, 126, 210–211, 268 Memory T-cell migration . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 IFN-γ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210

T-CELL TRAFFICKING Subject Index 287 labelling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 peritoneal membranes or frozen tissue sections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 MetaMorph 4.5 software . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 MHC, see Major histocompatibility (MHC) Microtubule organizing centre (MTOC) . . . . . . . . . . . . . 100 Migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4, 8, 15–276 Migration of T cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3–12 antigen, effects of . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6–7 T and B cells, antigen-induced trapping . . . . . . . 6–7 transgenic DTH model . . . . . . . . . . . . . . . . . . . . . . . . 7 disposal of combatants . . . . . . . . . . . . . . . . . . . . . . . . . . 5–6 adhesion molecules, expression of . . . . . . . . . . . . . . . 5 antigen-specific T cells . . . . . . . . . . . . . . . . . . . . . . . 5–6 immune system, role of . . . . . . . . . . . . . . . . . . . . . . . . . 6 LPS, TLR signals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 multi-step transmigration process . . . . . . . . . . . . . . . 6 regulatory T cells, types of . . . . . . . . . . . . . . . . . . . . . . 6 resident leucocytes, stress signals . . . . . . . . . . . . . . . . 5 TNF/IFNγ , cytokines . . . . . . . . . . . . . . . . . . . . . . . . . 6 migratory pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 cytokine-like mediators . . . . . . . . . . . . . . . . . . . . . . . . 9 G-protein-coupled receptors . . . . . . . . . . . . . . . . . . . . 9 organ-specific homing, paradigm. . . . . . . . . . . . . . .7–9, 8 α4 β7 -integrin/MAdCAM and α4 βx -integrin . . . . 8 “central”/“effectormemory” T cells . . . . . . . . . . . . . . 8 CLA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 E-selectin/P-selectin . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 “homing,” description . . . . . . . . . . . . . . . . . . . . . . . . . . 7 L-selectin, CCR7, 8 peripheral lymph node homing . . . . . . . . . . . . . . . . . 7 selective homing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 “Stamper-Woodruff ” assay . . . . . . . . . . . . . . . . . . . . . 7 VAP-1/CD44, adhesion molecules . . . . . . . . . . . . . . 8 recirculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3–4 blood flow and cell motility . . . . . . . . . . . . . . . . . . . . . 4 chemokine receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 LFA-1/α4 -integrins, components of transmigration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 L-selectin, homing-related molecule . . . . . . . . . . . . . 4 lymphocytes, role of . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 migratory mechanisms . . . . . . . . . . . . . . . . . . . . . . . 3–4 within tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4–5 chemokine receptor CCR7, 4–5 chemotactic mechanisms . . . . . . . . . . . . . . . . . . . . . . . 4 FTY 720 drug . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 process of emigration . . . . . . . . . . . . . . . . . . . . . . . . . . 5 prostaglandins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 S1P, chemotactic agent . . . . . . . . . . . . . . . . . . . . . . . . . 4 topographical memories . . . . . . . . . . . . . . . . . . . . . . . . 9–11 chemokine receptor CCR9, 10 epigenetic imprinting mechanisms . . . . . . . . . . . . . 11 GATA-3 and “histone code,” 11 homing receptors, induction/expression . . . . . . . . . . 9 mucosal homing receptor . . . . . . . . . . . . . . . . . . . . . . 10 organ-specific homing . . . . . . . . . . . . . . . . . . . . . . . . . 9 selectin ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Th1 or Th2 cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Molecules. . . . . . . . .4–6, 8–9, 18, 25, 45, 79, 81, 85, 97–99, 101, 119, 125, 135, 139, 149, 170, 172, 194, 210, 243, 253, 268, 271–272, 275 Monomeric red fluorescent protein 1 (mRFP1) . . . . . . . . 99, 100f, 107f MP-IVM, see Multiphoton intravital microscopy (MP-IVM)

MPLSM, see Multiphoton laser-scanning microscopy (MPLSM) MRFP1, see Monomeric red fluorescent protein 1 (mRFP1) MS, see Multiple sclerosis (MS) MTOC, see Microtubule organizing centre (MTOC) Multi-channel registration and co-localization, illustrations of . . . . . . . . . . . . . . . 223f Multicolour intravital two-photon microscopy . . . . . . . . 108 Multiphoton confocal microscopy . . . . . . . . . . . . . . . . . . . 167 Multiphoton intravital microscopy (MP-IVM) . . 181, 183, 187 Multiphoton laser-scanning microscopy (MPLSM) 194–195, 201 Multiphoton microscopy . . . . . . . . . . . . . . . . . . . 168, 181–204 Multiple fluorescence labelling . . . . . . . . . . . . . . . . . . . . . . 209 Multiple sclerosis (MS) . . . . . . . . . . . . . . . . . . . . . . . 8, 70, 267

N National Institutes of Health AIDS Research and Reference Reagent program . . . . . . . . . . . . . . . . 18 Natural killer (NK) cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 NCCs, see Neural crest cells (NCCs) NCS, see Newborn calf serum (NCS) Near infra-red (NIR) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Neural crest cells (NCCs) . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 Newborn calf serum (NCS) . . . . . . . . . . . . . . . . 256, 258–259 Nikon Eclipse TE2000-S microscope . . . . . . . . . . . 117, 120 NIR, see Near infra-red (NIR) NK, see Natural killer (NK) cells Nonlinear multi-photon microscopy . . . . . . . . . . . . . 108–109 R . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Novex Sharp R  . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 NuPAGE

O Olympus UCMAD3 QICAH . . . . . . . . . . . . . . . . . . . . . . . . 55 Omniscript Reverse Transcriptase Kit . . . . . . . . . . . . . . . . 128 Openlab (Improvision) . . . . . . . . . . . . . . . . . . . . 169–170, 173 Optical microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109, 220 R medium . . . . . . . . . . . . . . . . . . . . . . . . . . 19, 25 Opti-MEM Organogenesis study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 168 multiphoton microscopes . . . . . . . . . . . . . . . . . . . . . . . . 168 stereo microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 168 OVA, see Ovalbumin (OVA)-specific Ovalbumin (OVA)-specific . . . . . . . 117–118, 120, 122, 195

P Panasonic AG-6730, 55 Paracellular . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69–81 Paracellular/transcellular diapedesis, markers in VE-cadherin/PECAM- 1/caveolin-1, 79 Paraformaldehyde (PFA) . . . 19, 22, 71, 101, 104, 106, 163, 174–177, 210–213, 255, 257 Parajunctional PECAM-1, 81 Parallel-plate flow chamber . 22, 34, 37–39, 44f, 52–53, 54f, 58 cone and plate viscometer . . . . . . . . . . . . . . . . . . . . . . . . . 34 “flow chambers,” two dimensional . . . . . . . . . . . . . . . . . 34 for laminar flow experiments with HUVEC alloproliferation . . . . . . . . . . . . . . . . . . . . . . . . . . . 41–42 apparatus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39–41 CD4 T-cell alloproliferation . . . . . . . . . . . . . . . . . . 42f CD4 T-cell proliferation to PHA . . . . . . . . . . . . . 43f

T-CELL TRAFFICKING

288 Subject Index

Parallel-plate (continued) flow loop apparatus, diagram of . . . . . . . . . . . . . . . 37f immunohistochemistry . . . . . . . . . . . . . . . . . . . . . . . . 41 parallel flow chamber . . . . . . . . . . . . . . . . . . . . . . . . 38f PHA proliferation assay . . . . . . . . . . . . . . . . . . . 42–43 Parallel-plate recirculating flow loop system . . . . . . . . . . . . 37 PBL, see Peripheral blood lymphocytes (PBL) PBMCs, see Peripheral blood mononuclear cells (PBMCs) PBS, see Phosphate-buffered saline (PBS) PBSA, see Phosphate-buffered saline (PBSA) PCR, see Polymerase chain reaction (PCR) PDMS, see Polydimethylsiloxane (PDMS) PE, see Phycoerythrin (PE) PECAM-1 or CD99, 70 Penicillin/streptomycin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 Peripheral blood lymphocytes (PBL) . . . . . . . 22, 55, 59–60, 66, 89, 241, 245f, 254, 259f, 261f, 262, 264f, 265f Peripheral blood mononuclear cells (PBMCs) . . . . 254, 272 by density-gradient centrifugation . . . . . . . . . . . 254–255 Peritoneal infiltration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212 Peyer’s Patches (PP) . . . . . . . . . . . . . . . . . . . . . . . . . 4, 162, 165 PF4, see Platelet factor 4 (PF4) PFA, see Paraformaldehyde (PFA) PHA, see Phytohaemagglutinin (PHA) Phalloidin-TRITC (P1971) . . . . . . . . . . . . . . . . . . . . . . . . . . 72 Phase contrast microscopy . . . . . . . . . 40, 43, 50, 53, 57, 244 Phase contrast parallel-plate chamber . . . . . . . . . . 53–54, 53f Phenylmethanesulphonylfluoride (PMSF) . . . . . . . . . . . . . 86 PHOGEMON, see Phosphorylation and guanine-nucleotide exchange monitor (PHOGEMON) probes Phosphate-buffered saline (PBS) . . . . . . . . . . . 19, 22, 34–37, 40–41, 50, 56, 60, 71–72, 74–75, 101–104, 117–118, 120, 127–130, 129, 134–135, 139, 142–143, 150, 152–154, 163–164, 174, 185, 199, 233–234, 236, 242–244, 246, 255–257, 260, 270, 272 Phosphate-buffered saline (PBSA) . . . . . . . . . . . . . . . . . . . . 50 Phosphoinositide 3-kinase (PI3K) . . . . . . . . . . . . . . 126, 134 Phosphorylation and guanine-nucleotide exchange monitor (PHOGEMON) probes . . . . . . . . . . . . . . . . . . . 98 Phycoerythrin (PE) . . . . . . . . . . . . . . . . . . . 20, 129, 134–135, 217, 239, 255, 257, 261–262, 271 Phycoll procedure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 Phytohaemagglutinin (PHA) . . . . . . . . . . . . . . . . . 35, 71, 128 proliferation assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35–36 PI3K, see Phosphoinositide 3-kinase (PI3K) Platelet factor 4 (PF4) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127 PML, see Progressive multifocal leukoencephalopathy (PML) PMSF, see Phenylmethanesulphonylfluoride (PMSF) Polydimethylsiloxane (PDMS) . . . . . . . . . . . . . . . . . . . . . . 119 Polymerase chain reaction (PCR) . . . . . . . . . . . 40, 128–129, 131, 133, 140–141, 151 Polyvinylidene fluoride (PVDF) . . . . . . . . . . . . . . . . . . . 87, 92 Popliteal lymph node (popLN) . . . . . . . . 182–183, 186–187 PopLN, see Popliteal lymph node (popLN) “Population biochemistry,” 99 Portex Blue Line Manometer . . . . . . . . . . . . . . . . . . . . . 61–62 PP, see Peyer’s Patches (PP) Progressive multifocal leukoencephalopathy (PML) . . . . . . . . . . . . . . . 211–212, 221–222, 268 Prokaryotic protein expression plasmids (pGEX – GE Healthcare). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .85

Proliferation . . . . . . . . . . . . . . . 5, 32, 35, 41–42, 43f, 72, 125, 130, 150, 239 Protein interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 PVDF, see Polyvinylidene fluoride (PVDF)

R Rac1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85–86, 89, 106, 107f Ras homology (Rho) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Real-time polymerase chain reaction (RT-PCR) . . . . . . . . . . . . . . . . . . . . . . . . . . 140, 151 Receptor–arrestin complex . . . . . . . . . . . . . . . . . . . . . . . . . . 126 Receptor tyrosine kinases (RTK). . . . . . . . . . . . . . . . . . . . . .98 Recirculation . . . . . . . . . . . . . . . 3, 5, 8–9, 37–38, 43, 44f, 211 Regulatory T cells, types of Foxp3+ Tregs and IL-10, 6 Retinoic acid-related orphan receptors (RORs). . . . . . . .165 Reverse transcriptase (RT) enzyme . . . . . . . . . 128, 132, 140 Rheumatoid arthritis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50, 194 RhoA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85–86, 88–89, 99 RhoGDIs, see RhoGTPase dissociation inhibitors (RhoGDIs) RhoGTPase activity in lymphocytes . . . . . . . . . . . . . . . 83–94 materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Cdc42 activity decreases in response to stimulation of WT L-selectin . . . . . . . . . . . . . . . . . . . . . . . . 93f cell culture and cell lines . . . . . . . . . . . . . . . . . . . . . . 85 cell stimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 GST-fused effector domain bound to glutathione sepharose beads . . . . . . . . . . . . . . . . . . . . . . . . . . 90f quantification and statistical analysis . . . . . . . . . . . 88 RhoA/Rac1/Cdc42, protein expression and purification . . . . . . . . . . . . . . . . . . . . . . . . . . . 85–86 RhoGTPase activation assays . . . . . . . . . . . . . . . 86–87 SDS-PAGE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 western blotting . . . . . . . . . . . . . . . . . . . . . . . . . . 87–88 methods cell disruption techniques . . . . . . . . . . . . . . . . . . . . . 88 cell stimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 maintenance of cell culture . . . . . . . . . . . . . . . . . 88–89 quantification and analysis . . . . . . . . . . . . . . . . . . . . . 94 RhoA/Rac1/Cdc42, protein expression and purification . . . . . . . . . . . . . . . . . . . . . . . . . . . 89–90 SDS-PAGE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92 western blotting . . . . . . . . . . . . . . . . . . . . . . . . . . 92–93 RhoGTPase cycling between active and inactive states . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84f RhoGTPase dissociation inhibitors (RhoGDIs) . . . . . . . . 84 RNA polymerase II . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 sample processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 “Rolling adhesion,” 20, 244 RORs, see Retinoic acid-related orphan receptors (RORs) Roswell Park Memorial Institute (RPMI) . . . . . . . . . 18–19, 21, 35, 51, 56, 71–72, 85, 88, 91, 101–104, 127–128, 130–131, 133–134, 137, 139, 143, 163, 169–170, 172–173, 196–197, 200, 233, 236, 246, 255–256, 258–259, 264, 269–270, 272–273 RPMI, see Roswell Park Memorial Institute (RPMI) RT, see Reverse transcriptase (RT) enzyme RTK, see Receptor tyrosine kinases (RTK) RT-PCR, see Real-time polymerase chain reaction (RT-PCR)

T-CELL TRAFFICKING Subject Index 289 S Scaffolding protein caveolin-1 staining . . . . . . . . . . . . . . . . 81 SDS-PAGE, see SDS-polyacrylamide gel electrophoresis (SDS-PAGE) SDS-polyacrylamide gel electrophoresis (SDS-PAGE) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 SEB, see Staphylococcal enterotoxin B (SEB) “Secretory” smooth muscles . . . . . . . . . . . . . . . . . . . . . . . . . . 65 SEE, see Superantigen enterotoxin E (SEE) Shear stress . . . . . . . . . . . . . . . . . . . 23, 32–33, 37, 40, 54f, 62, 67, 244, 249, 275 disturbed flow/low fluid shear stress (atherosclerotic lesions) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 fluid shear stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 high shear stress (atheroprotective) . . . . . . . . . . . . . . . . 33 Signalling . . . . . . . . . . . . . . . . . . . . 10, 79–80, 84, 88, 97–110, 125–145, 162, 222, 254 Silicone-intensified target (SIT) system . . . . . . . . . . 271, 274 Single-photon confocal microscopy . . . . . . . . . . . . . . . . . . 167 SIT, see Silicone-intensified target (SIT) system Smooth muscle cells (SMC) . . . . . . . . . . . . 33, 49, 55–58, 65 S1P, see Sphingosine-1-phosphate (S1P) Sphingosine-1-phosphate (S1P) . . . . . . . . . . . . . . . . . . . 4, 116 Spinal cord window . . . . . . . . . . . . . . . . . . 269–270, 273–275, 277f, 278–279 Spinning disc confocal microscopy . . . . . . . . . . . . . . . . . . . 167 S6 ribosomal protein, phosphorylation of . . . . . . . . . . . . . 134 Staining of living cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20 cell labeling with fluorescent probes . . . . . . . . . 20, 25–26 fluorescently labeled antibodies . . . . . . . . . . . . . . . . 20, 25 Stamper–Woodruff adhesion assay . . . . . . . . . . . . . . 233–234 adherent cells/endothelium/quantification . . . . 241–242 adhesion assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241 medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 analysis of lymphocyte binding to tissue sections . . 240f detection antibody . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 preparation of lymphocytes for adhesion assay . . . . . . . . . . . . . . . 241 tissue specimens as adhesive substrates . . . . 240–241 secondary and tertiary antibodies for APAAP staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 tris buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 water-soluble mountant . . . . . . . . . . . . . . . . . . . . . . . . . 234 Staphylococcal enterotoxin B (SEB) . . . . . . . . . . . . . . . . . 127 “Static” adhesion assays . . . . . . . . . . . . . . . . . . . . . . . . 243, 249 Static adhesion measurements and staining . . . . . . . . . . . . 19 confocal microscopy, visualization by . . . . . . . . . . . . . . 20f static adhesion measurements . . . . . . . . . . . . . . . . . . 19, 21 static adhesion staining . . . . . . . . . . . . . . . . . . . . . . . 19, 22 Stereo fluorescent microscopy . . . . . . . . . . . . . . . . . . . . . . . 176 “Stomacher bag,” 236 Stromal cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4, 49–67, 168 culture of . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 fibroblast complete medium . . . . . . . . . . . . . . . . . . . 51 promocell SMC medium . . . . . . . . . . . . . . . . . . . . . . 51 Superantigen enterotoxin E (SEE) . . . . . . . . . . . . . . 101, 103 Surrogate target cells . . . . . . . . . . . . . . . . . . . . . . 182, 185, 188

T TAE, see 50X Tris-acetate-EDTA (TAE) buffer TBS, see Tris-buffered saline (TBS) TCA, see Trichloroacetic acid (TCA) T cell receptor . . . . . . . . . . . . . . . . . . . . . 100, 139–140, 182, 195 receptor zeta

T-cell alloproliferation assay . . . . . . . . . . . . . . . . . . . . . . . . . . 35 T-cell-based cancer immunotherapy . . . . . . . . . . . . . . . . . 116 T-cell migration/embryonic development . . . . . . . . 207–226 materials embryonic sections in gene expression studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210–211 embryonic tissue for expression analysis of transcription factors . . . . . . . . . . . . . . . . . . . . . . 211 memory T-cell migration . . . . . . . . . . . . . . . . . . . . 210 RNA polymerase II . . . . . . . . . . . . . . . . . . . . . . . . . 211 non-lymphoid tissues, quantification of . . . . . . . . . . 218f sample preparation/imaging . . . . . . . . . . . . . . . . . 211–213 embryonic tissue for expression analysis of transcription factors . . . . . . . . . . . . . . . . . . . . . . 213 TEM imaging of RNA polymerase II . . . . . . . . . 212 tissue-infiltrating memory T cells . . . . . . . . 211–212 transcription factors, analysis of . . . . . . . . . . . . . . . 212 tissue cell quantification, watershed-based algorithm . . . . . . . . . . . . . . . . . . . . . . . . . . . 213–216 algorithm implementation . . . . . . . . . . . . . . . 216–217 application to TEM images . . . . . . . . 220–222, 221f general description . . . . . . . . . . . . . . . . . . . . . . . . . . 213 image pre-processing . . . . . . . . . . . . . . . . . . . . . . . . 216 implementation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 local maximum detection in target identification . . . . . . . . . . . . . . . . . . 213–216, 215f quantification of Hox-induced cell clustering . . . . . . . . . . . . . . . . . . . . . . . . . . . 219–220 T cell in non-lymphoid tissues . . . . . . . . . . . 217–218 transcription factors in embryonic development . . . . . . . . . . . . . . . . . . . . . . . . 222–224 T cell receptor (TCR) . . . . . . . . . . . . . . . . . . . . . . . . . . 139, 182 TCR complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 α and β chains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 CD3 complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 heterogeneity of TCRζ expression in PBL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264f ITAMs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 TCRζ (CD247) functions . . . . . . . . . . . . . . . . . . . 254 TCRζ expression in CD56+ and CD16+ cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265f TCRζ expression in healthy PB T cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261f, 263f TCRζ low-expressing cells or TCRζ dim T cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 TCR, see T cell receptor (TCR) TCRζ bright and TCRζ dim T cells . . . . . . . . . . . . . . . . . . 265 TCSPC, see Time-correlated single photon counting (TCSPC) TEM, see Transendothelial migration (TEM) Therapeutic targeting of human T-cell trafficking in vivo, strategies . . . . . . . . . . . . . . . . . . . . . . . . . . . 267–279 materials cell surface adhesion molecules, blocking of . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271–272 human T-cell preparation . . . . . . . . . . . . . . . . 267–270 intravital fluorescence videomicroscopy . . . . . . . . 271 labeling of human T cells with fluorescent dye . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 spinal cord window in SJL mice with EAE, preparation . . . . . . . . . . . . . . . . . . . . . . . . . 270–271 methods cell surface adhesion molecules . . . . . . . . . . . . . . . 275 data analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275–278 human T-cell purification . . . . . . . . . . . . . . . . . . . . 272 intravital fluorescence videomicroscopy . . . 274–275

T-CELL TRAFFICKING

290 Subject Index

Therapeutic targeting (continued) labeling of human T cells with fluorescent dye 272–273 spinal cord window preparation in mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273–274 Time-correlated single photon counting (TCSPC) . . . . . . . . . . . . . . . . . . . . . 105f, 106, 110 Time-lapse confocal microscopy . . . . . . . . . . . . . . . . . . . . . 23f Time-lapse fluorescence confocal microscopy . . . . . . . . . . 24f Time-lapse fluorescence microscopy . . . . . . . . . . . . . . . 20, 26 Time-lapse microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . 99, 169 Tissue cell quantification . . . . . . . . . . . . . . . . . . . . . . . 207–226 See also T-cell migration/embryonic development Tissue entry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4, 7 Tissue exit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4–5, 7 Tissue-specific endothelial adhesion assays . . . . . . . 242–243 EC isolation from liver homogenate . . . . . . . . . . . . . . 243 sample collection and preparation . . . . . . . . . . . . . . . . 242 T-lymphoblast transmigration, detection . . . . . . . . . . . 75–77 confocal software . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 detail of paracellular diapedesis in HUVECs . . . . . . . 76f diapedesis in HUVECs . . . . . . . . . . . . . . . . . . . . . . . . . . 75f epifluorescence, analysis by . . . . . . . . . . . . . . . . . . . . . . . 75 micrographs of co-cultures . . . . . . . . . . . . . . . . . . . . . . . . 77 paracellular vs. transcellular diapedesis . . . . . . . . . . 77–79 T-lymphoblasts diapedesis . . . . . . . . . . . . . . . . . . . . . . . . 77 paracellular/transcellular diapedesis, markers in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 transcellular diapedesis in HDMVECs . . . . . . . . 78f T lymphocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . 40, 71–72, 99, 125–145, 182, 242–243, 269 T lymphopoiesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 TNF-α, see Tumour necrosis factor alpha (TNF-αp TNF-α stimulation, media for . . . . . . . . . . . . . . . . . . . . . . . . 80 Toxin Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Trafficking . . . . . . . . . . . . . . . . . . . . . . . . 5, 8, 70, 79, 116, 137, 211, 267–279 Transcellular . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18, 69–81 Transendothelial migration assay . . . . . . . . . . . 256, 259–260 transmigration of healthy donor PBL stained for TCRζ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259f Transendothelial migration (TEM) . . . . . . . . . . . . . . . . 69–81 See also Diapedesis, paracellular/transcellular routes discrimination Transfection . . . . . 19, 24–25, 28, 29, 79–80, 101–102, 110, 149–156 “Transmigrated cells,” 244 Transmigration . . . . . . . . 4–6, 20, 22–23, 27–28, 59f, 66, 70, 72–74, 75f, 76–77, 78f, 79–81, 117–118, 122, 149, 234, 235f, 243, 246, 250–251, 254, 259f, 260, 264–265 Transmigration assay in diapedesis mimicking long-term inflammation. . . . . . . . . . . . . . . . 73 stimulation of HDMVEC monolayers . . . . . . . . . . 73 stimulation of HUVECs monolayers . . . . . . . . . . . 73 preparation of cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 transmigration assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Transplantation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Transwell assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59f Trichloroacetic acid (TCA) . . . . . . . . . . . . . . . . . . . . . 151, 155 Tris-buffered saline (TBS) . . . . . . 19, 22, 27–28, 71, 74, 87, 92–93, 104, 233, 241 R reagent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35, 40, TRIzol 45, 128, 132, 133f, 140 Tumor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182, 185, 191 Tumour necrosis factor alpha (TNF-αp . . . . . . . . . . . 50–51

U University of Birmingham . . . . . . . . . . . . . . . . . . . . . . . . 52, 54 University of Strathclyde . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201

V Visualisation of signalling in immune cells . . . . . . . . 97–111 accuracy/specificity of assays, improvement . . . 106–107 bi-exponential fluorescence decay model . . . . . . . 106 Raichu biosensors . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Raichu-Cdc42 FLIM data . . . . . . . . . . . . . . . . . . . 107 Raichu-Rac-expressing Jurkat T cells . . . . 106, 107f YTS NK cell surveying a potential target cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108f future development . . . . . . . . . . . . . . . . . . . . . . . . . 108–110 deep-tissue imaging . . . . . . . . . . . . . . . . . . . . . . . . . 109 “deformable mirrors,” 109 eGFP with red fluorescent protein mKeima, combination of . . . . . . . . . . . . . . . . . . . . . . . . . . 109 multicolour intravital two-photon microscopy . . 108 nonlinear multi-photon microscopy . . . . . . . 108–109 optical microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 “surgically accessible” targets . . . . . . . . . . . . . . . . . 109 materials cell culture/transfection by electroporation . . . . . 101 cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100–101 conjugate formation . . . . . . . . . . . . . . . . . . . . . 101–102 ligand-dependent activation assay for jurkat T cells, 101 methods cell culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 confocal or multi-photon excitation TCSPC FLIM set-up, 105f electroporation . . . . . . . . . . . . . . . . . . . . . . . . . 102–103 FLIM and confocal microscopy, see Fluorescence lifetime imaging microscopy (FLIM) fluorescence lifetime imaging of fixed samples 105–106 integrin ligand and antibody stimulation assays . 103 ligand-dependent activation assay, see Ligand-dependent activation assay for live imaging Vitronectin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101, 103 Volocity (Improvision). . . . . . . . . . . .169–170, 173, 185, 201

W “Walk-out” method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248 “Wash reservoir” and “sample reservoir,” 61 Watershed segmentation . . . . . . . . . . . . . . . . . . . 209, 213, 216 Western LightningTM Chemiluminescence reagent . . . . 88 Western-type diet (WTD) . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Wide-field fluorescent stereo microscopy . . . . . . . . . . . . . 167 Wnt1-Cre/Foxn1-Cre/Tie2-Cre transgenic mice. . . . . .167 Wolfson Applied Technology Laboratory . . . . . . . . . . 52, 54 WTD, see Western-type diet (WTD)

X Xcell II blot module (Invitrogen) . . . . . . . . . . . . . . . . . . . . . 87 R gel electrophoresis tank (Invitrogen) . 87 Xcell SureLock 50X Tris-acetate-EDTA (TAE) buffer . . . . . . . . . . . . . . . 128

Y Yellow fluorescent protein (YFP) . . . . . . . . . . 24, 98–99, 167 YFP, see Yellow fluorescent protein (YFP)