T-Cell Trafficking: Methods and Protocols [1 ed.] 160761460X, 9781607614609, 9781607614616

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ME T H O D S

IN

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

TM

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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3.

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

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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

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5.

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

Live Imaging of Leukocyte–Endothelium Interactions

2.2. Static Adhesion Measurements and Staining

1. BCECF-AM (Invitrogen)

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)

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2. HBSS (BioWhittaker) supplemented with 1% BSA 3. Plastic seal

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).

Live Imaging of Leukocyte–Endothelium Interactions

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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.

8.

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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 Wrap, X-ray film (Fuji , Japan) for the development of 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 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 reagent (Invitrogen, UK). 1. TRIzol

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