Suppression and Regulation of Immune Responses: Methods and Protocols [1 ed.] 1607618680, 9781607618683, 9781607618690

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METHODS

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MOLECULAR BIOLOGY™

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

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Suppression and Regulation of Immune Responses Methods and Protocols

Edited by

Maria Cristina Cuturi INSERM UMR 643, Nantes, France and

Ignacio Anegon INSERM UMR 643, Nantes, France

Editors Dr. Maria Cristina Cuturi INSERM UMR 643, Nantes, France [email protected]

Ignacio Anegon, MD INSERM UMR 643, Nantes, France [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-868-3 e-ISBN 978-1-60761-869-0 DOI 10.1007/978-1-60761-869-0 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010936185 © Springer Science+Business Media, LLC 2011 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. Cover Illustration Caption: The Yin/Yang image. In this case, the black represents immune response, and the white represents immune regulation. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Dedication To my wonderful, unique and beloved family

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Acknowledgments We would like to thank Mrs. Brenda Evans for her very efficient editorial assistance. We would also like to thank the support of RISET European program (http://www.risetfp6. org/), the program IMBIO (http://www.imbio.fr/accueil.html) from the Région Pays de la Loire, the Fondation Progreffe (http://www.progreffe.com/), and the Fondation Centaure (http://www.fondation-centaure.fr/).

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Preface “Le savant n’est pas l’homme qui fournit les vraies réponses, c’est celui qui pose les vraies questions.” Claude Lévi-Strauss

The immune system is able to mount an effective immune response to virtually any foreign antigen but does not react against self-components. One of the mechanisms for doing that is to create the largest repertoire of B and T cells able to recognize the most antigen candidates imaginable. This can also in some cases represent a problem as the immune system may recognize selfantigen as foreign antigens and mount an immune response leading to the destruction of self and developing autoimmune diseases. In order to avoid that, most autoreactive cells are eliminated or neutralized by a mechanism called self-tolerance. There are two components of self-tolerance: central and peripheral. Central tolerance occurs in the primary sites of lymphocytic maturation (thymus and bone marrow) where deletion of cells that react strongly with self-antigens (clonal deletion) takes place in the thymus (T cells) or bone marrow (B cells). These cells will undergo apoptosis and die (negative selection). The cells that have moderate self-recognition will survive and migrate to the periphery (positive selection). The lymphocytes that escape central tolerance come to the periphery where they can meet and also interact with self-antigen, but in the periphery there are also regulatory mechanisms which balance the activation of an immune response (immunity against pathogens) and its suppression to control the magnitude of the immune response and prevent unwanted damage or its regulation toward self-antigen (tolerance). Moreover, there are some special cases in which the immune system has to tolerate foreign antigens while requiring the capacity to respond to infections, as in the case of pregnancy. One of the goals of immunological research is to understand the mechanisms which regulate the immune system. We can define immunoregulation as a balance between activation and suppression/regulation of the immune response in order to achieve an efficient response without damaging the host. The characterization of the mechanisms of immunoregulation would not only allow us to make predictions about the outcome of a particular immune response, but also to define the ways of controlling this response either by amplification or inhibition. This would have practical consequences for certain clinical situations, e.g., the inhibition of the immune response in allergic, transplantation, or autoimmune diseases or the enhancement of the immune response in the case of infectious diseases or cancer. Over the past several years, a high diversity of regulatory cells and suppressive molecules has taken center stage in the field of immunoregulation. Several types of regulatory cells have been described on the basis of their origin, generation, and mechanisms of action. T-regulatory cells (naturally occurring and inducible ones) were the most studied, and several subsets were identified (CD4+, CD8+, and CD4−CD8−). This book will highlight recent advances in the identification, characterization, and generation of regulatory cells not only of the T-cell lineage but also of other origins such as B, NK, myeloid, and

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dendritic cells. We will also approach the role of several suppressive molecules in immunoregulation. Certain physiological situations where immunoregulation plays a central role as in pregnancy will be treated separately. Particular emphasis will be put on the characterization of the molecular mechanisms and the therapeutic applications of regulatory cells and molecules in human diseases. Nantes, France

Maria Cristina Cuturi Ignacio Anegon

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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PART I: REGULATORY CELLS 1 Natural and Induced T CD4+CD25+FOXP3+ Regulatory T Cells . . . . . . . . . . . . Lucienne Chatenoud 2 Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells. . . . . . . . . . . . . Petra Hoffmann, Ruediger Eder, and Matthias Edinger 3 Methods for In Vitro Generation of Human Type 1 Regulatory T Cells . . . . . . . . Silvia Gregori, Maria Grazia Roncarolo, and Rosa Bacchetta 4 Ex Vivo Generation of Regulatory T Cells: Characterization and Therapeutic Evaluation in a Model of Chronic Colitis . . . . . . . . . . . . . . . . . . Fridrik Karlsson, Sherry A. Robinson-Jackson, Laura Gray, Songlin Zhang, and Matthew B. Grisham 5 Phenotypic and Functional Characterization of CD8+ T Regulatory Cells . . . . . . . Séverine Ménoret, Carole Guillonneau, Séverine Bezié, Lise Caron, Ignacio Anegon, and Xian-Liang Li 6 Regulatory CD4– CD8– Double Negative T Cells . . . . . . . . . . . . . . . . . . . . . . . . . Edward Y. Kim, Stephen C. Juvet, and Li Zhang 7 Identifying Regulatory B Cells (B10 Cells) that Produce IL-10 in Mice . . . . . . . . Takashi Matsushita and Thomas F. Tedder 8 DCs in Immune Tolerance in Steady-State Conditions . . . . . . . . . . . . . . . . . . . . . Tomohiro Fukaya, Hideaki Takagi, Honami Taya, and Katsuaki Sato 9 Plasmacytoid Dendritic Cells in Tolerance. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eric Gehrie, William Van der Touw, Jonathan S. Bromberg, and Jordi C. Ochando 10 In Vitro-Generated DC with Tolerogenic Functions: Perspectives for In Vivo Cellular Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cees van Kooten and Kyra A. Gelderman 11 Preparation of Mouse Bone Marrow-Derived Dendritic Cells with Immunoregulatory Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mercedes Segovia, Maria Cristina Cuturi, and Marcelo Hill 12 Myeloid-Derived Suppressor Cells: Characterization and Expansion in Models of Endotoxemia and Transplantation . . . . . . . . . . . . . . . . . . . . . . . . . . Nicolas Van Rompaey and Alain Le Moine 13 Human Regulatory Macrophages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . James A. Hutchinson, Paloma Riquelme, Edward K. Geissler, and Fred Fändrich 14 NKT and Tolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julien Diana, Lucie Beaudoin, Anne-Sophie Gautron, and Agnès Lehuen

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15 Antiinflammatory and Immunosuppressive Functions of Mast Cells . . . . . . . . . . . 207 Janet Kalesnikoff and Stephen J. Galli 16 A Mesenchymal Stem Cell Potency Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Joy Jiao, Jack M. Milwid, Martin L. Yarmush, and Biju Parekkadan 17 In Vitro Analyses of the Immunosuppressive Properties of Neural Stem/Progenitor Cells Using Anti-CD3/CD28-Activated T Cells . . . . . . . . . . . . 233 Virginie Bonnamain, Isabelle Neveu, and Philippe Naveilhan

PART II: REGULATORY MOLECULES 18 Immunoregulatory Properties of Heme Oxygenase-1 . . . . . . . . . . . . . . . . . . . . . . Philippe Blancou, Virginie Tardif, Thomas Simon, Séverine Rémy, Leandro Carreño, Alexis Kalergis, and Ignacio Anegon 19 Indoleamine 2,3-Dioxygenase and Regulatory Function: Tryptophan Starvation and Beyond . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ciriana Orabona and Ursula Grohmann 20 Regulatory T Cell Enrichment by IFN-g Conditioning . . . . . . . . . . . . . . . . . . . . . Gang Feng, Kathryn J. Wood, and Andrew Bushell 21 Transforming Growth Factor-Beta: Recent Advances on Its Role in Immune Tolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pierre-Yves Mantel and Carsten B. Schmidt-Weber 22 Induction of Tolerogenic Dendritic Cells by NF-kB Blockade and Fcg Receptor Modulation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Leandro J. Carreño, Claudia A. Riedel, and Alexis M. Kalergis 23 Fine-Tuning Antitumor Responses Through the Control of Galectin–Glycan Interactions: An Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . Mariana Salatino and Gabriel A. Rabinovich 24 Regulation of Lymphocytes by Nitric Oxide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christian Bogdan

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PART III: PHYSIOPATHOLOGICAL SITUATIONS 25 Pregnancy: Tolerance and Suppression of Immune Responses . . . . . . . . . . . . . . . . Anne Leber, Maria Laura Zenclussen, Ana Teles, Nadja Brachwitz, Pablo Casalis, Tarek El-Mousleh, Federico Jensen, Katja Woidacki, and Ana Claudia Zenclussen 26 Organ Transplantation: Modulation of T-Cell Activation Pathways Initiated by Cell Surface Receptors to Suppress Graft Rejection. . . . . . . . . . . . . . . Kathleen Weatherly and Michel Y. Braun 27 Immunosuppressive Mechanisms During Viral Infectious Diseases . . . . . . . . . . . . Ghanashyam Sarikonda and Matthias G. von Herrath 28 Ocular Immune Privilege Sites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sharmila Masli and Jose L. Vega 29 Immunoprivileged Sites: The Testis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monika Fijak, Sudhanshu Bhushan, and Andreas Meinhardt Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors IGNACIO ANEGON • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France ROSA BACCHETTA • Department of Regenerative Medicine, Stem Cells, and Gene Therapy, San Raffaele Telethon Institute for Gene Therapy (HSR-TIGET), Milan, Italy LUCIE BEAUDOIN • INSERM U986, Hôpital Cochin/St Vincent de Paul, Paris, France; Université Paris Descartes, Paris, France SÉVERINE BEZIÉ • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France SUDHANSHU BHUSHAN • Department of Anatomy and Cell Biology, Justus-Liebig-University of Giessen, Giessen, Germany PHILIPPE BLANCOU • ONIRIS, Nantes, France; INRA, UMR_A 707, Nantes, France CHRISTIAN BOGDAN • Medical Microbiology and Immunology of Infectious Diseases, Microbiology Institute – Clinical Microbiology, Immunology and Hygiene, Friedrich-Alexander-University Erlangen-Nuremberg and University Clinic of Erlangen, Erlangen, Germany VIRGINIE BONNAMAIN • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France NADJA BRACHWITZ • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany MICHEL Y. BRAUN • Institute for Medical Immunology, Université Libre de Bruxelles (ULB), Gosselies, Belgium JONATHAN S. BROMBERG • Department of Gene and Cell Medicine, Mount Sinai School of Medicine, New York, NY, USA ANDREW BUSHELL • Transplantation Research Immunology Group, Nuffield Department of Surgery, John Radcliffe Hospital, University of Oxford, Oxford, UK LISE CARON • Faculté de Médecine, INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Université de Nantes, Nantes, France LEANDRO CARREÑO • Millenium Nucleus of Immunology and Immunotherapy, Departamento de Genética Molecular y Microbiología, Facultad de Ciencias Biológicas, Pontificia Universidad Catolica de Chile, Santiago, Chile PABLO CASALIS • Department for Neurosurgery, Charite, Medical University of Berlin, Berlin, Germany LUCIENNE CHATENOUD • INSERM U1013, Faculté Paris Descartes, Hôpital Necker-Enfants Malades, Paris, France

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MARIA CRISTINA CUTURI • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France JULIEN DIANA • INSERM U986, Hôpital Cochin/St Vincent de Paul, Paris, France; Université Paris Descartes, Paris, France RUEDIGER EDER • Department of Hematology and Oncology, University Hospital Regensburg, Regensburg, Germany MATTHIAS EDINGER • Department of Hematology and Oncology, University Hospital Regensburg, Regensburg, Germany TAREK EL-MOUSLEH • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany FRED FÄNDRICH • Clinic for Applied Cell Therapy, University Hospital of Schleswig-Holstein, Kiel, Germany GANG FENG • Transplantation Research Immunology Group, Nuffield Department of Surgery, John Radcliffe Hospital, University of Oxford, Oxford, UK MONIKA FIJAK • Department of Anatomy and Cell Biology, Justus-Liebig-University of Giessen, Giessen, Germany TOMOHIRO FUKAYA • Laboratory for Dendritic Cell Immunobiology, RIKEN Research Center for Allergy and Immunology, Kanagawa, Japan STEPHEN J. GALLI • Department of Pathology, Stanford University School of Medicine, Stanford CA, USA ANNE-SOPHIE GAUTRON • INSERM U986, Hôpital Cochin/St Vincent de Paul, Paris, France; Université Paris Descartes, Paris, France ERIC GEHRIE • Department of Gene and Cell Medicine, Mount Sinai School of Medicine, New York, NY, USA EDWARD K. GEISSLER • Laboratory for Transplantation Research, Department of Surgery, University Hospital Regensburg, Regensburg, Germany KYRA A. GELDERMAN • Department of Nephrology, Leiden University Medical Center, Leiden, The Netherlands LAURA GRAY • Immunology and Inflammation Research Group, Department of Molecular and Cellular Physiology, LSU Health Sciences Center, Shreveport LA, USA SILVIA GREGORI • Department of Regenerative Medicine, Stem Cells, and Gene Therapy, San Raffaele Telethon Institute for Gene Therapy (HSR-TIGET), Milan, Italy MATTHEW B. GRISHAM • Immunology and Inflammation Research Group, Department of Molecular and Cellular Physiology, LSU Health Sciences Center, Shreveport LA, USA URSULA GROHMANN • Department of Experimental Medicine and Biochemical Sciences, University of Perugia, Perugia, Italy CAROLE GUILLONNEAU • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France MARCELO HILL • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France

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PETRA HOFFMANN • Department of Hematology and Oncology, University Hospital Regensburg, Regensburg, Germany JAMES A. HUTCHINSON • Laboratory for Transplantation Research, Department of Surgery, University Hospital Regensburg, Regensburg, Germany SHERRY A. ROBINSON-JACKSON • Immunology and Inflammation Research Group, Department of Molecular and Cellular Physiology, LSU Health Sciences Center, Shreveport LA, USA FEDERICO JENSEN • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany JOY JIAO • Center for Engineering in Medicine and Surgical Services, Massachusetts General Hospital, Harvard Medical School, Boston MA, USA; Department of Chemical Engineering, Massachusetts Institute of Technology, Cambridge MA, USA STEPHEN C. JUVET • Toronto General Hospital Research Institute, Toronto ON, Canada ALEXIS KALERGIS • Millenium Nucleus of Immunology and Immunotherapy, Departamento de Genética Molecular y Microbiología, Facultad de Ciencias Biológicas, Departamento de Reumatología, Facultad de Medicina, Pontificia Universidad Catolica de Chile, Santiago, Chile JANET KALESNIKOFF • Department of Pathology, Stanford University School of Medicine, Stanford CA, USA FRIDRIK KARLSSON • Immunology and Inflammation Research Group, Department of Molecular and Cellular Physiology, LSU Health Sciences Center, Shreveport LA, USA EDWARD Y. KIM • Toronto General Hospital Research Institute, Toronto ON, Canada CEES VAN KOOTEN • Department of Nephrology, Leiden University Medical Center, Leiden, The Netherlands ALAIN LE MOINE • Institute for Medical Immunology, Université Libre de Bruxelles, Gosselies, Belgium AGNÈS LEHUEN • INSERM U986, Hôpital Cochin/St Vincent de Paul, Paris, France; Université Paris Descartes, Paris, France XIAN-LIANG LI • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France PIERRE-YVES MANTEL • Department of Immunology and Infectious Diseases, Harvard School of Public Health, Harvard University, Boston MA, USA SHARMILA MASLI • Department of Ophthalmology, Harvard Medical School, Schepens Eye Research Institute, Boston MA, USA TAKASHI MATSUSHITA • Department of Immunology, Duke University Medical Center, Durham NC, USA ANDREAS MEINHARDT • Department of Anatomy and Cell Biology, Justus-Liebig-University of Giessen, Giessen, Germany SÉVERINE MÉNORET • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France; Platform Rat Transgenic Nantes IBiSA-CNRS, Nantes, France

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JACK M. MILWID • Center for Engineering in Medicine and Surgical Services, Massachusetts General Hospital, Harvard Medical School, BostonMA, USA; Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge MA, USA PHILIPPE NAVEILHAN • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France ISABELLE NEVEU • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France JORDI C. OCHANDO • Department of Gene and Cell Medicine, Mount Sinai School of Medicine, New York, NY, USA; Laboratorio de Immunología de Trasplantes, Centro Nacional de Microbiología, Instituto de Salud Carlos III, Madrid, Spain CIRIANA ORABONA • Department of Experimental Medicine and Biochemical Sciences, University of Perugia, Perugia, Italy BIJU PAREKKADAN • Center for Engineering in Medicine and Surgical Services, Massachusetts General Hospital, Harvard Medical School, Boston MA, USA GABRIEL A. RABINOVICH • Laboratorio de Inmunopatología, Instituto de Biología y Medicina Experimental, Consejo Nacional de Investigaciones Científicas y Técnicas, Buenos Aires, Argentina; Departamento de Química Biológica, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Buenos Aires, Argentina SÉVERINE RÉMY • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France CLAUDIA A. RIEDEL • Millenium Nucleus of Immunology and Immunotherapy, Departamento de Genética Molecular y Microbiología, Laboratorio de Biología Celular y Farmacología, Departamento de Ciencias Biológicas, Facultad de Ciencias Biológicas, Pontificia Universidad Catolica de Chile, Santiago, Chile PALOMA RIQUELME • Laboratory for Transplantation Research, Department of Surgery, University Hospital Regensburg, Regensburg, Germany MARIA GRAZIA RONCAROLO • Department of Regenerative Medicine, Stem Cells, and Gene Therapy, San Raffaele Telethon Institute for Gene Therapy (HSR-TIGET), Milan, Italy; Universita’ Vita-Salute San Raffaele, Milan, Italy MARIANA SALATINO • Laboratorio de Inmunopatología, Instituto de Biología y Medicina Experimental, Consejo Nacional de Investigaciones Científicas y Técnicas, Buenos Aires, Argentina GHANASHYAM SARIKONDA • Department of Developmental Immunology-3, Diabetes Center of San Diego, La Jolla Institute for Allergy and Immunology, La Jolla CA, USA KATSUAKI SATO • Laboratory for Dendritic Cell Immunobiology, RIKEN Research Center for Allergy and Immunology, Kanagawa, Japan CARSTEN B. SCHMIDT-WEBER • Allergy and Clinical Immunology, National Heart and Lung Institute, Imperial College, London, UK

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ANNE SCHUMACHER • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany MERCEDES SEGOVIA • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France THOMAS SIMON • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France HIDEAKI TAKAGI • Laboratory for Dendritic Cell Immunobiology, RIKEN Research Center for Allergy and Immunology, Kanagawa, Japan VIRGINIE TARDIF • INSERM UMR 643, Nantes, France; CHU Nantes, Institut de Transplantation et de Recherche en Transplantation (ITERT), Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France HONAMI TAYA • Laboratory for Dendritic Cell Immunobiology, RIKEN Research Center for Allergy and Immunology, Kanagawa, Japan THOMAS F. TEDDER • Department of Immunology, Duke University Medical Center, DurhamNC, USA ANA TELES • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany WILLIAM VAN DER TOUW • Department of Gene and Cell Medicine, Mount Sinai School of Medicine, New York, NY, USA NICOLAS VAN ROMPAEY • Institute for Medical Immunology, Université Libre de Bruxelles, Gosselies, Belgium JOSE L. VEGA • Department of Ophthalmology, Harvard Medical School, Schepens Eye Research Institute, Boston, MA, USA; Department of Neurology, Columbia University Medical Center, Columbia University College of Physicians and Surgeons, New York, NY, USA MATTHIAS G. VON HERRATH • Department of Developmental Immunology (D1-3), Diabetes Center of San Diego, La Jolla Institute for Allergy and Immunology, La Jolla CA, USA KATHLEEN WEATHERLY • Institute for Medical Immunology, Université Libre de Bruxelles (ULB), Gosselies, Belgium KATJA WOIDACKI • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany KATHRYN J. WOOD • Transplantation Research Immunology Group, Nuffield Department of Surgery, John Radcliffe Hospital, University of Oxford, Oxford, UK MARTIN L. YARMUSH • Center for Engineering in Medicine and Surgical Services, Massachusetts General Hospital, Harvard Medical School, Boston MA, USA; Harvard-MIT Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge MA, USA ANA CLAUDIA ZENCLUSSEN • Division of Reproductive Immunology, Department for Experimental Obstetrics and Gynaecology, University of Magdeburg, Magdeburg, Germany

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MARIA LAURA ZENCLUSSEN • Department for Neurosurgery, Charite, Medical University of Berlin, Berlin, Germany LI ZHANG • Toronto General Hospital Research Institute, Toronto ON, Canada SONGLIN ZHANG • Immunology and Inflammation Research Group, Department of Pathology, LSU Health Sciences Center, Shreveport LA, USA

Part I Regulatory Cells

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Chapter 1 Natural and Induced T CD4+CD25+FOXP3+ Regulatory T Cells Lucienne Chatenoud Abstract Evidence has been accumulated to show that the forkhead/winged-helix transcription factor Foxp3 is a good marker for specialized CD4+ T cells that regulate immune responses to self as well as to a variety of foreign antigens including infectious or tumor antigens, alloantigens, allergens, and commensal antigens. It is now well established that CD4+CD25+Foxp3+ regulatory T cells encompass two categories of lymphocytes that are distinct in their origin, antigen specificity, as well as the stimuli driving their differentiation and homeostasis. Natural CD4+CD25+Foxp3+ regulatory T cells are an independent lineage generated in the thymus through major histocompatibility class II molecules-dependent MHC class high avidity interactions with their T cell receptor. They are specific for self-antigens. Adaptive or induced CD4+CD25+Foxp3+ regulatory T cells stem from mature CD4+CD25-Foxp3-precursors at the periphery following adequate stimulation. They have been shown to develop in vivo following suboptimal antigen stimulation, in situations characterized by chronic inflammation (autoimmunity, allergy, immune responses to tumors and transplants) and also as physiological actors of the mucosal immune system. Although major progress has been accomplished over the last years in our understanding of the central role of CD4+CD25+Foxp3+ regulatory T cells in the control of immune responses, major issues are still elusive. In particular, there are still no reliable phenotypic or functional markers that make it possible to distinguish between natural and induced CD25+Foxp3+ regulatory T cells. Key words: Natural regulatory T cells, Induced regulatory T cells, Foxp3, Immune regulation

1. Introduction As all biological systems the immune system is redundant and finely tuned. Immune regulatory mechanisms make it possible to face extreme situations in which discrete signals must be amplified or conversely the intensity of some immune responses, which could be the source of “damage,” must be tempered. These immune regulatory mechanisms are complex; some are antigen-specific Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_1, © Springer Science+Business Media, LLC 2011

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and others not. Major players in mediating immune regulation are specialized subsets of T lymphocytes some of which exert helper functions, thus amplifying effector B and T cell responses, and others which operate as suppressors. The concept of a distinct subset of lymphocytes mediating suppressive or regulatory functions was initially proposed by Gershon and Kondo in the 1970s, and they coined the term “infectious tolerance” (1). For many years, the tools to approach suppressive or regulatory T cells remained more than elusive, which explains the difficult situation of the whole field for about 20 years until, in the 1990s, important technical steps were accomplished that drove the revival of the field of regulatory T cells based on robust molecular grounds. It is now well established that depending on the experimental model or the clinical situation, distinct cell types, including subsets of CD4+ and CD8+ T cells and invariant NK T cells, which all may be presented in detail in the following chapters, may afford downregulation of immune responses. Our aim, here, is to focus on CD4+ Foxp3-expressing regulatory T cells, also termed Tregs, which are effective at containing immune responses to self and foreign antigens as well as to commensal microorganisms. According to the present state of the art, one may distinguish essentially two sets of CD4+Foxp3+ Tregs: “natural” CD4+ Foxp3+ Tregs (nTregs) emerging from the thymus as a distinct lineage (2–4) and “adaptive” or “induced” CD4+CD25+ regulatory T cells (iTregs) that develop outside the thymus, from mature CD4+CD25− T cell precursors, under particular conditions, i.e., antigenic stimulation and cytokine environment (5–9). It would be unproductive and tedious to draw the long list of methodological details that allow purification and characterization of nTregs and iTregs in the many in vitro and in vivo studies, which have been reported over the last 15 years. In addition, it is fair to admit that we are still in the quest of reliable phenotypic and/or functional markers to distinguish nTregs from iTregs. Therefore, we shall address in parallel the salient steps that lead to the characterization of these two categories of Tregs, which in fact were closely related to the methodological advances gained though the multiplicity of experimental approaches used, and that involved the study of conventional and genetically modified murine models.

2. General Considerations There is no single method that affords characterizing in a reliable fashion nTregs and iTregs, neither in mice nor in humans. Therefore, the usage of a combination of tests is needed to assess in parallel the phenotypic pattern as well as the functional activity

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of the selected T-cell subset and, importantly, to interpret the data based on the physiological or pathological situation under study. The phenotypic characterization is possible using whole mononuclear cell or lymphocyte populations, recovered mostly from peripheral blood in humans and from various lymphoid organs in mice, as it is feasible by fluorescence-activated cell analysis using combinations of membrane markers to gate on individual T lymphocyte subsets (e.g., CD4+CD25+ T cells) looking for the expression of other key markers (Foxp3, CTLA-4, GITR within gated cells (10–13). At variance with cell phenotyping, the analysis of the functional capacities of putative Tregs populations requires the purification of the given subset prior to testing by means of in vitro culture techniques or, in the case of experimental studies, in vivo models. Cell purification is performed by classical techniques using monoclonal antibodies to trace specific subsets; labeled cells are then separated from the unlabelled ones by magnetic bead cell sorting or fluorescence-activated cells sorting (FACS) (14–17). The latter method allows a much better purification of the selected subset especially when the marker used is expressed at variable density in the population of interest. This is, for instance, the case for CD4+CD25+ nTregs, which express a high density of CD25 (the a chain of the IL-2 receptor).

3. In Vitro Functional Analysis

Both in humans and in experimental models the method that has been and is widely used for the analysis of the in vitro functional capacity of Tregs to suppress is the so-called coculture assay. The principle is simple and consists in culturing purified “responder” T cells together with putative Tregs, also purified, to assess the capacity of the latter population to refrain the response of the former. The most commonly used marker of response is cell proliferation although the production of a given cytokine has also been adopted. For cell proliferation analysis, the two methods commonly used are conventional incorporation of tritiated thymidine. The pulse labeling of cells with radioactive thymidine provides a means to determine the distribution of times of entry into the first cell division. Alternatively, staining with 5-,6-carboxyfluorescein diacetate succinimidyl ester (CFSE) of the responder T cell subset is also used. CFSE is a vital dye whose intracellular concentration will decrease proportionally with cell division and serves visualizing (by FACS) and classifying proliferating cell populations into groups according to the number of divisions each cell has undergone.

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In practice, as originally described the coculture test used as responder T cells purified polyclonal CD4+CD25− T cells (or when needed purified CD8+ T cells) stimulated either with antigen and low numbers of antigen-presenting cells (APCs) or anti-T-cell receptor (mostly anti-CD3) antibody. When needed, in order to analyze what are the mechanisms underlying the effect observed, namely, cell–cell contact interactions and/or soluble mediators or both, transwell devices are used which physically separate the two cell subsets. This method was used in the case of both mouse and human cells. Although it is obvious that in vitro systems will never fully recapitulate the in vivo situation, important properties of Tregs were approached with this method. It could thus be established that first Tregs suppress the proliferation and interleukin-2 (IL-2) production of responder cells in a dose-dependent fashion; second, both CD4+ and CD8+ cells could be suppressed; third, the suppressive activity of Tregs could be bypassed following addition of exogenous interleukin (IL)-2 or by triggering costimulatory pathways (e.g., CD28:B7); and fourth, depending on the situations, the suppressive ability of Tregs was or was not impaired through blockade of given immune regulatory cytokines such as TGF-b or IL-10. Although exceptions may be found in the literature, the consensus is that thymic-derived nTregs exert their in vitro function in a cell–cell contact-dependent but cytokine-independent way (18), while iTregs use both pathways.

4. In Vivo Functional Analysis

4.1. Neonatal Thymectomy Experiments

These methods are, for obvious reasons, mostly restricted to mouse experimental models. Among in vivo manipulations that allow testing for the suppressive effect of Tregs one may cite: (1) in vivo depletion experiments whereby elimination of Tregs, following thymectomy on day 3 after birth or upon administration of monoclonal antibodies that deplete the target population (antiCD25) (19), an exacerbation of given immune responses is observed and, (2) adoptive transfer experiments, mostly through i.v. injection, of purified subsets of putative Tregs to analyze their capacity to downregulate in vivo immune responses that develop in the recipients. Most commonly, immune-mediated inflammatory, autoimmune, allergic, or tumor responses were targeted. These adoptive transfers may be performed either in immunecompetent or immune-deficient recipients (mice bearing the scid, nu/nu, or Rag−/− mutations and therefore devoid of T cells). The seminal experiment showing that cells expressing suppressive functions emerged from the thymus was that performed in the late 1960s by the group of Nishizuka and Sakakura (20).

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These authors showed that neonatal thymectomy of female mice resulted in developmental arrest of the ovary leading to sterility. The ovaries of thymectomized mice were small, completely lacked follicles and corpora lutea, a histopathological pattern now recognized as autoimmune oophoritis (20). Disease was observed when mice were thymectomized at 3 days of age, but not at 7 days or later, and it was fully prevented by thymus grafting. As shown later, disease was also prevented by adoptive transfer of syngeneic T splenocytes from adult normal donors. In addition, more detailed analysis showed that the day 3 thymectomy induced in fact a more generalized autoimmune syndrome, also including a high proportion of mice showing gastritis and thyroïditis. Then started the long quest for cell markers which could afford characterizing among mature T lymphocytes the subset endowed with the regulatory properties; cells were negatively selected based on given surface molecules and then transferred to neonatally thymectomized mice to test their potential to prevent the autoimmune syndrome. CD5 was the first marker identified; removing CD5+ cells from donor splenocytes reduced the capacity of peripheral T cells to protect day 3 thymectomized mice from autoimmune oophoritis and gastritis. As a mirror image, CD5-depleted splenocytes adoptively transferred into immunodeficient/lymphopenic mice induced multiorgan autoimmunity (21). A major finding in the mid-1990s was that CD25, the high-affinity subunit of the IL-2 receptor (IL2Ra), was another important marker of the thymic-derived suppressive cells. Almost undetectable levels of peripheral CD4+CD25+ T cells were present a week after day 3 thymectomy, and tenfold less splenic CD4+CD25+ T cells were detected in thymectomized adult mice compared to controls (22). Importantly, CD4+CD25+ regulatory T cells were capable of preventing autoimmunity not only in neonatally thymectomized mice but also in lymphopenic animals infused with pathogenic effector T lymphocytes (2, 23). This was demonstrated in models whereby transferring CD25+ T-cell-depleted splenocytes into lymphopenic hosts induced a multiorgan autoimmunity syndrome that resembled the one observed in neonatally thymectomized mice (2). Similarly, transfer of purified CD4+CD45RBhigh cells into immunodeficient hosts induced severe inflammatory colitis leading to wasting syndrome (24, 25). The major breakthrough that gave molecular “credit” to the concept that thymic-derived suppressor T cells constituted a bona fide individual lineage of thymocytes was the description in 2003, by three independent laboratories (26–28), that the Foxp3 transcription factor was highly expressed constitutively by CD4+CD25+ regulator y T cells, while CD4+CD25− T cells did not upregulate Foxp3 when they acquired CD25 expression following TRC stimulation. Foxp3 is a forkhead/winged-helix transcription factor family member.

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Mutations in the Foxp3 encoding gene had been identified as responsible for the Immune dysregulation, Polyendocrinopathy, Enteropathy, X-linked (IPEX) syndrome (29), described in 1982 characterized by a cohort of autoimmune manifestations that develop during early infancy in affected males. A similar X-linked pathology was described back in 1959 in the scurfy mouse which was later on also attributed to a mutation in Foxp3 gene. Using bone marrow chimeras and also using Foxp3 reporter mice (see below), it was ascertained that FoxP3 was mandatory for nTreg development in the thymus and constituted a valuable marker for this independent lineage of T cells. In fact, a significant proportion of developing T cells likely commit to the regulatory T-cell lineage within the thymus, as CD4+CD25+ with suppressive activity are detected in neonatal mice prior to peripheral generation of activated CD25+ cells in the spleen (3). In keeping with this conclusion were also the data showing that adoptive transfer of nTregs isolated from normal wild-type mice significantly prevented disease and related mortality in Foxp3 mutant mice (3). 4.2. Generation of FoxP3 Reporter Mice

Once Foxp3 was found to represent the lineage-specific marker for Tregs, a major effort was devoted to create a model which could allow purification of T cells based on Foxp3 expression. In fact, Foxp3 being a transcription factor, staining of Foxp3+ cells in conventional mice necessitated cell fixation, which certainly allowed cell phenotyping but totally excluded in vitro or in vivo manipulations for which living cells were needed. This was the rationale for creating reporter Foxp3GFP mice in which the green fluorescent protein-coding sequence was inserted in-frame within exon2 of Foxp3. In these reporter mice, approximately 96% of Foxp3+ cells resided within the CD4+ T-cell compartment; 60–88% also expressing high levels of CD25, depending on anatomical location (3). Interestingly, the analysis of these mice directly validated the reports describing that Tregs were also included within the CD4+CD25− or CD4+CD25low compartments. Comparison of the gene “signatures” of Foxp3+CD4+CD25high and Foxp3+CD4+CD25low/− T cells showed a greater similarity between these two subsets than between Foxp3+CD4+CD25high and Foxp3−CD4+CD25− T cells. Additionally, low yet detectable numbers of Foxp3+ cells were detected among a small TCRab+ that stained CD8+. Finally, it is important to mention that thymic differentiation of nTregs is CD28-dependent (30).

4.3. Regulatory T Cells Recognize Self-Determinants

Since the neonatal thymectomy-induced autoimmunity model was described, nTregs were hypothesized to recognize self-antigens. As mentioned above, transferring total splenocytes from normal syngeneic donors as a source of Tregs protected thymectomized neonates from the autoimmune syndrome. In contrast, adoptive transfer of splenocytes from donors in which a target tissue was

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removed (in the case for instance of splenic preparations from athyroid donors) prevented all autoimmune manifestations but thyroiditis (31). The conclusion proposed was that the presence of the target antigens was mandatory to maintain nTregs. Concerning their thymic development, compelling evidence has been accumulated to show that the TCR affinity threshold that induces nTregs was placed immediately after or during signal strengths that negatively select self-reactive thymocytes. Experimental models used mice that coexpress a transgenic TCR and its transgene encoded cognate ligand in the thymus (32). In these mice, cognate ligand-induced engagement of TCR on developing thymocytes resulted in significant thymic deletion, but remaining thymocytes contained significantly higher percentages of nTregs. 4.4. Extrathymic Regulatory T-Cell Differentiation

Studies using polyclonal populations (7) and suggesting that extrathymic generation of Tregs was possible were challenged by the description of Foxp3-expressing CD4+CD25− cells (3) that could become Foxp3-expressing CD25+ cells. The initial experimental evidence in support of the generation of regulatory T cells from peripheral CD4+CD25-Foxp3− T cells came from studies of adoptive transfer of T cells from TCR transgenic recombinase (RAG)-deficient mice that are devoid of regulatory T cells, which are not generated in the absence of endogenous TCR gene rearrangements. Upon transfer of these TCR transgenic T cells into hosts harboring the cognate antigen, intense proliferation was observed followed by apoptosis/ contraction of most dividing T cells; surviving cells expressed Foxp3 and suppressor functions. The group of A. Abbas transferred naïve DO11 RAG−/− T-cells (anti-ovalbumin (OVA)) into recipients in whom the cognate antigen is expressed ubiquitously as a secreted soluble protein (33). The transferred T cells expanded and differentiated into pathogenic effectors causing a transient graft-versus-host disease. In the surviving mice, once the syndrome resolved, OVA-specific CD25+Foxp3+ Tregs were detected (33). Using nonlymphopenic recipients, the group of H. van Boehmer demonstrated that regulatory T-cell differentiation is favored when stimulating T cells under poorly immunogenic conditions. Thus, a subset of CD25+Foxp3+ Tregs could be established in TCR-hemagglutinin (HA) Tg RAG−/− mice in which the cognate antigen was delivered by an osmotic pump (34). Similarly, hemagglutinin (HA)-specific CD25− T cells transferred into nonlymphopenic hosts exposed to variable doses of antigen coupled to the dendritic cell-specific anti-DEC205 monoclonal antibody promoted the appearance of CD25+FoxP3+ Tregs, the highest proportion of Tregs being observed when using low doses of antigen (35). Moreover, the in vivo Foxp3 induction

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is significantly impaired in T cells insensitive to TGF-b signaling (expressing a dominant-negative TGF-b receptor) (35). Differentiation of iTregs has also been described in a variety of other situations including: induction of oral tolerance (36), autoimmunity (14, 37), allergic inflammation (38), tumor responses (39), and transplant responses (9). Moreover, they appear as an essential “physiological” component of the gut immune system generated by the response to microbiota and food antigens (40, 41). At this point, it is important to mention that it was shown quite early that CD4+CD25+ Tregs, which subsequently were identified as Foxp3+, could be expanded in vitro from naïve conventional T cells following TCR-mediated stimulation in presence of TGF-b (5). These iTregs were shown to acquire functional suppressive capacities in vitro, in coculture, and also in vivo in an allergic asthma model. The mechanisms that mediate the capacity of TGF-b to induce Foxp3 have now been dissected at the molecular level and involve the transcription factors STAT 5 and NF-AT (42). At this point, it is interesting to mention the work of the group of H. Weiner showing that following distinct stimulations regulatory CD4+Foxp3+ cells expressing membrane TGF-b (LAP+ T cells) that are derived from peripheral precursors, especially from T cells in the gut, may be induced (43, 44). Another cytokine that is important for iTreg differentiation and/ or survival is IL-2. IL-2 is essential for the in vitro generation of TGF-b-induced iTregs. Aside from dedicated cytokines, additional microenvironmental factors have been reported as important to drive the differentiation of iTregs. This is the case for retinoic acid. Dendritic cells expressing the integrin CD103 present in the lamina propria and mesenteric lymph nodes that produce both TGF-b and retinoic acid are highly effective at mediating conversion of naïve T cells into Foxp3+ iTregs (40).

5. Conclusions In spite of a long controversy, it is now well accepted that CD4+CD25+Foxp3+ play an essential role in immune regulation. It is also well established that these cells come in two flavors, thymically-derived nTregs and iTregs emerging from peripheral precursors. Their precise respective roles in the physiological and pathological settings must still be fully elucidated and new tools must be generated to approach a more precise characterization of the two subsets. This will be important not only from the fundamental point of view but also in terms of clinical translation as great hope

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is focused (as discussed in detail in other chapters of this volume) on the use of expanded Tregs as therapeutic tools in various pathological conditions (45). It is important to add before closing that here we exclusively concentrated on Foxp3+iTregs. There is, however, in the literature compelling evidence to show the existence of subsets of CD4+ T cells mediating effective regulation that are induced at the periphery following adequate antigenic stimulation and lack Foxp3. These include, in particular, Tr1 cells (described in more detail elsewhere in this volume) that have been characterized by the group of M.G. Roncarolo and that play a major role in the regulation of transplantation responses and autoimmunity (46–48). References 1. Gershon, R.K., and K. Kondo. 1971. Infectious immunological tolerance. Immunology 21:903–914. 2. Sakaguchi, S., N. Sakaguchi, M. Asano, M. Itoh, and M. Toda. 1995. Immunologic self-tolerance maintained by activated T cells expressing IL-2 receptor alpha-chains (CD25). Breakdown of a single mechanism of selftolerance causes various autoimmune diseases. J Immunol 155:1151–1164. 3. Kim, J.M., and A. Rudensky. 2006. The role of the transcription factor Foxp3 in the development of regulatory T cells. Immunol Rev 212:86–98. 4. Fontenot, J.D., J.P. Rasmussen, L.M. Williams, J.L. Dooley, A.G. Farr, and A.Y. Rudensky. 2005. Regulatory T cell lineage specification by the forkhead transcription factor foxp3. Immunity 22:329–341. 5. Chen, W., W. Jin, N. Hardegen, K.J. Lei, L. Li, N. Marinos, G. McGrady, and S.M. Wahl. 2003. Conversion of peripheral CD4+CD25naive T cells to CD4+CD25+ regulatory T cells by TGF-beta induction of transcription factor Foxp3. J Exp Med 198:1875–1886. 6. Liang, S., P. Alard, Y. Zhao, S. Parnell, S.L. Clark, and M.M. Kosiewicz. 2005. Conversion of CD4+ CD25- cells into CD4+ CD25+ regulatory T cells in vivo requires B7 costimulation, but not the thymus. J Exp Med 201:127–137. 7. Curotto de Lafaille, M.A., A.C. Lino, N. Kutchukhidze, and J.J. Lafaille. 2004. CD25- T cells generate CD25+Foxp3+ regulatory T cells by peripheral expansion. J Immunol 173:7259–7268. 8. Karim, M., C.I. Kingsley, A.R. Bushell, B.S. Sawitzki, and K.J. Wood. 2004.

9.

10.

11.

12.

13.

14.

Alloantigen-induced CD25+CD4+ regulatory T cells can develop in vivo from CD25-CD4+ precursors in a thymus-independent process. J Immunol 172:923–928. Cobbold, S.P., R. Castejon, E. Adams, D. Zelenika, L. Graca, S. Humm, and H. Waldmann. 2004. Induction of foxP3+ regulatory T cells in the periphery of T cell receptor transgenic mice tolerized to transplants. J Immunol 172: 6003–6010. Takahashi, T., T. Tagami, S. Yamazaki, T. Uede, J. Shimizu, N. Sakaguchi, T.W. Mak, and S. Sakaguchi. 2000. Immunologic self-tolerance maintained by CD25(+)CD4(+) regulatory T cells constitutively expressing cytotoxic T lymphocyte-associated antigen 4. J Exp Med 192:303–310. Shimizu, J., S. Yamazaki, T. Takahashi, Y. Ishida, and S. Sakaguchi. 2002. Stimulation of CD25(+)CD4(+) regulatory T cells through GITR breaks immunological self-tolerance. Nat Immunol 3:135–142. Read, S., V. Malmstrom, and F. Powrie. 2000. Cytotoxic T lymphocyte-associated antigen 4 plays an essential role in the function of CD25(+)CD4(+) regulatory cells that control intestinal inflammation. J Exp Med 192: 295–302. Tang, Q., and J.A. Bluestone. 2006. Regulatory T-cell physiology and application to treat autoimmunity. Immunol Rev 212:217–237. You, S., B. Leforban, C. Garcia, J.F. Bach, J.A. Bluestone, and L. Chatenoud. 2007. Adaptive TGF-{beta}-dependent regulatory T cells control autoimmune diabetes and are a privileged target of anti-CD3 antibody treatment. Proc Natl Acad Sci USA 104:6335–6340.

12

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15. Mchugh, R.S., M.J. Whitters, C.A. Piccirillo, D.A. Young, E.M. Shevach, M. Collins, and M.C. Byrne. 2002. CD4(+)CD25(+) immunoregulatory T cells: gene expression analysis reveals a functional role for the glucocorticoidinduced TNF receptor. Immunity 16:311–323. 16. Piccirillo, C.A., J.J. Letterio, A.M. Thornton, R.S. Mchugh, M. Mamura, H. Mizuhara, and E.M. Shevach. 2002. CD4(+)CD25(+) regulatory T cells can mediate suppressor function in the absence of transforming growth factor beta1 production and responsiveness. J Exp Med 196:237–246. 17. You, S., M. Belghith, P. Cobbold, M.A. Alyanakian, C. Gouarin, S. Barriot, C. Garcia, H. Waldmann, L. Chatenoud, and J.F. Bach. 2005. Autoimmune diabetes onset results from qualitative rather than quantitative agedependent changes in pathogenic T cells. Diabetes 54(5):1415–1422. 18. Shevach, E.M., R.A. DiPaolo, J. Andersson, D.M. Zhao, G.L. Stephens, and A.M. Thornton. 2006. The lifestyle of naturally occurring CD4+ CD25+ Foxp3+ regulatory T cells. Immunol Rev 212:60–73. 19. Billiard, F., E. Litvinova, D. Saadoun, F. Djelti, D. Klatzmann, J.L. Cohen, G. Marodon, and B.L. Salomon. 2006. Regulatory and effector T cell activation levels are prime determinants of in vivo immune regulation. J Immunol 177:2167–2174. 20. Nishizuka, Y., and T. Sakakura. 1969. Thymus and reproduction: sex-linked dysgenesia of the gonad after neonatal thymectomy in mice. Science 166:753–755. 21. Sakaguchi, S., K. Fukuma, K. Kuribayashi, and T. Masuda. 1985. Organ-specific autoimmune diseases induced in mice by elimination of T cell subset. I. Evidence for the active participation of T cells in natural self-tolerance; deficit of a T cell subset as a possible cause of autoimmune disease. J Exp Med 161:72–87. 22. Asano, M., M. Toda, N. Sakaguchi, and S. Sakaguchi. 1996. Autoimmune disease as a consequence of developmental abnormality of a T cell subpopulation. J Exp Med 184:387–396. 23. Suri-payer, E., A.Z. Amar, A.M. Thornton, and E.M. Shevach. 1998. CD4+CD25+ T cells inhibit both the induction and effector function of autoreactive T cells and represent a unique lineage of immunoregulatory cells. J Immunol 160:1212–1218. 24. Powrie, F., R. Correa-oliveira, S. Mauze, and R.L. Coffman. 1994. Regulatory interactions between CD45R Bhigh and CD45 RBlow CD4+ T cells are important for the balance between protective and pathogenic cell-mediated immunity. J Exp Med 179: 589–600.

25. Stephens, L.A., C. Mottet, D. Mason, and F. Powrie. 2001. Human CD4(+)CD25(+) thymocytes and peripheral T cells have immune suppressive activity in vitro. Eur J Immunol 31:1247–1254. 26. Fontenot, J.D., M.A. Gavin, and A.Y. Rudensky. 2003. Foxp3 programs the development and function of CD4+CD25+ regulatory T cells. Nat Immunol 4:330–336. 27. Hori, S., T. Nomura, and S. Sakaguchi. 2003. Control of regulatory T cell development by the transcription factor Foxp3. Science 299: 1057–1061. 28. Khattri, R., T. Cox, S.A. Yasayko, and F. Ramsdell. 2003. An essential role for Scurfin in CD4+CD25+ T regulatory cells. Nat Immunol 4:337–342. 29. Wildin, R.S., S. Smyk-Pearson, and A.H. Filipovich. 2002. Clinical and molecular features of the immunodysregulation, polyendocrinopathy, enteropathy, X linked (IPEX) syndrome. J Med Genet 39:537–545. 30. Salomon, B., D.J. Lenschow, L. Rhee, N. Ashourian, B. Singh, A. Sharpe, and J.A. Bluestone. 2000. B7/CD28 Costimulation is essential for the homeostasis of the CD4+CD25+ immunoregulatory T cells that control autoimmune diabetes. Immunity 12:431–440. 31. Seddon, B., and D. Mason. 1999. Peripheral autoantigen induces regulatory T cells that prevent autoimmunity. J Exp Med 189: 877–882. 32. Jordan, M.S., A. Boesteanu, A.J. Reed, A.L. Petrone, A.E. Holenbeck, M.A. Lerman, A. Naji, and A.J. Caton. 2001. Thymic selection of CD4(+)CD25(+) regulatory T cells induced by an agonist self-peptide. Nat Immunol 2:301–306. 33. Knoechel, B., J. Lohr, E. Kahn, J.A. Bluestone, and A.K. Abbas. 2005. Sequential development of interleukin 2-dependent effector and regulatory T cells in response to endogenous systemic antigen. J Exp Med 202: 1375–1386. 34. Apostolou, I., and H. von Boehmer. 2004. In vivo instruction of suppressor commitment in naive T cells. J Exp Med 199:1401–1408. 35. Kretschmer, K., I. Apostolou, D. Hawiger, K. Khazaie, M.C. Nussenzweig, and H. von Boehmer. 2005. Inducing and expanding regulatory T cell populations by foreign antigen. Nat Immunol 6:1219–1227. 36. Mucida, D., N. Kutchukhidze, A. Erazo, M. Russo, J.J. Lafaille, and M.A. Curotto de Lafaille. 2005. Oral tolerance in the absence of naturally occurring Tregs. J Clin Invest 115:1923–1933.

Natural and Induced T CD4+CD25+FOXP3+ Regulatory T Cells 37. You, S., N. Thieblemont, M.A. Alyanakian, J.F. Bach, and L. Chatenoud. 2006. Transforming growth factor-beta and T-cell-mediated immunoregulation in the control of autoimmune diabetes. Immunol Rev 212:185–202. 38. Curotto de Lafaille, M.A., N. Kutchukhidze, S. Shen, Y. Ding, H. Yee, and J.J. Lafaille. 2008. Adaptive Foxp3+ regulatory T cell-dependent and -independent control of allergic inflammation. Immunity 29:114–126. 39. Liu, V.C., L.Y. Wong, T. Jang, A.H. Shah, I. Park, X. Yang, Q. Zhang, S. Lonning, B.A. Teicher, and C. Lee. 2007. Tumor evasion of the immune system by converting CD4+ CD25- T cells into CD4+CD25+ T regulatory cells: role of tumor-derived TGF-beta. J Immunol 178:2883–2892. 40. Sun, C.M., J.A. Hall, R.B. Blank, N. Bouladoux, M. Oukka, J.R. Mora, and Y. Belkaid. 2007. Small intestine lamina propria dendritic cells promote de novo generation of Foxp3 T reg cells via retinoic acid. J Exp Med 204:1775–1785. 41. Barnes, M.J., and F. Powrie. 2009. Regulatory T cells reinforce intestinal homeostasis. Immunity 31:401–411. 42. Josefowicz, S.Z., and A. Rudensky. 2009. Control of regulatory T cell lineage commitment and maintenance. Immunity 30:616–625. 43. Ochi, H., M. Abraham, H. Ishikawa, D. Frenkel, K. Yang, A.S. Basso, H. Wu, M.L. Chen, R. Gandhi, A. Miller, R. Maron, and

44.

45.

46.

47.

48.

13

H.L. Weiner. 2006. Oral CD3-specific antibody suppresses autoimmune encephalomyelitis by inducing CD4(+)CD25(-)LAP(+) T cells. Nat Med 12:627–635. Chen, M.L., B.S. Yan, Y. Bando, V.K. Kuchroo, and H.L. Weiner. 2008. Latencyassociated peptide identifies a novel CD4+ CD25+ regulatory T cell subset with TGFbetamediated function and enhanced suppression of experimental autoimmune encephalomyelitis. J Immunol 180:7327–7337. Tang, Q., K.J. Henriksen, M. Bi, E.B. Finger, G. Szot, J. Ye, E.L. Masteller, H. McDevitt, M. Bonyhadi, and J.A. Bluestone. 2004. In vitro-expanded antigen-specific regulatory T cells suppress autoimmune diabetes. J Exp Med 199:1455–1465. Battaglia, M., A. Stabilini, E. Draghici, B. Migliavacca, S. Gregori, E. Bonifacio, and M.G. Roncarolo. 2006. Induction of tolerance in type 1 diabetes via both CD4+CD25+ T regulatory cells and T regulatory type 1 cells. Diabetes 55:1571–1580. Groux, H., A. O’garra, M. Bigler, M. Rouleau, S. Antonenko, J.E. De Vries, and M.G. Roncarolo. 1997. A CD4+ T-cell subset inhibits antigen-specific T-cell responses and prevents colitis. Nature 389:737–742. Battaglia, M., S. Gregori, R. Bacchetta, and M.G. Roncarolo. 2006. Tr1 cells: from discovery to their clinical application. Semin Immunol 18:120–127.

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Chapter 2 Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells Petra Hoffmann, Ruediger Eder, and Matthias Edinger Abstract Based on results from experimental animal models, the adoptive transfer of CD4+CD25+FOXP3+ regulatory T cells (Treg) is expected to be efficacious in treating autoimmune and inflammatory diseases, as well as in preventing alloresponses after solid organ or stem-cell transplantation. For potential clinical applications, large numbers of Treg cells in maximum purity will be required to avoid the risk of disease exacerbation by contaminating effector T cells. We have recently described methods for the efficient in vitro expansion of human Treg cells and identified CD4+CD25highCD45RA+ T cells as the ideal starting population for the generation of homogeneous and stable Treg cell products. Here, we provide detailed instructions for their identification, isolation, expansion, and functional characterization. Key words: Immunotherapy, Tolerance, Suppressor T cell, Transplantation, T-cell therapy

1. Introduction Thymus-derived CD4+CD25+Foxp3+ regulatory T cells play a crucial role in the maintenance of peripheral self-tolerance, as loss-of-function mutations of the foxp3 gene abrogate their suppressive activity and cause fatal autoimmunity in mice and humans (1). Similarly, the deletion of Foxp3+ Treg cells causes autoimmune syndromes in rodents but strengthens immune responses to tumors (2) or to microbial infections (3). Inversely, their adoptive transfer protects from autoimmunity and exacerbated inflammatory responses in various disease models (4). In experimental colitis, transferred Treg cells even cure ongoing disease (5). These findings highlight the relevance of Treg cells for physiological immune reactions and raised interest in their therapeutic manipulation. Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_2, © Springer Science+Business Media, LLC 2011

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Treg-based immune modulation is particularly attractive in the context of allogeneic transplantation. Several groups could show that they contribute to tolerance induction after solid organ transplantation (6) and the coinfusion of donor-derived Treg cells protects from lethal graft-versus-host disease (GVHD) after allogeneic stem-cell transplantation (SCT) (7). As allogeneic SCT is per se a cellular therapy, both in terms of replacement of the myeloid compartment and with regard to the beneficial graftversus leukemia effect (GVL) mediated by donor T cells, this treatment modality seems ideally suited for the exploration and exploitation of cellular immune suppression. In murine SCT experiments, the coinfusion of large numbers of freshly isolated donor Treg cells [at a 1:1 ratio with conventional T cells (Tconv)] prevented GVHD without abrogating GVL effects (8, 9). Clinical trials exploring this strategy are now ongoing at our and several other institutions and good manufacturing practice (GMP)compatible isolations strategies have been developed for that purpose (10). Yet, it has to be emphasized that such isolation strategies were specifically designed for the prevention of GVHD in SCT, where the cotransplantation of Tconv cells is intended for the prevention of opportunistic infections and the promotion of GVL effects. Thus, these technologies purposely enrich Tregs to only 50–60% purity (10) [and unpublished data] but were explicitly not developed for the treatment of GVHD or any other diseases, as a contamination of the transferred cell product with effector T cells might aggravate the respective condition and endanger patients (11). In our opinion, a therapeutic Treg cell product has to fulfil several criteria: (1) Clinically relevant cell numbers have to be achieved reliably; therefore, efficient in vitro expansion of Treg cells is required. (2) Maximum purity of Treg cells, as determined by current knowledge and technology, has to be enforced (currently evaluated by the level of FOXP3 expression). (3) No proinflammatory cytokines should be secreted, as this might contribute to disease progression. To address these issues, we previously described efficient in vitro expansion protocols that permit the 2–3 log expansion of CD4+CD25high Tregs within 2–3 weeks. Although human and murine Treg cells were described to be anergic and therefore unsuited for large-scale expansion, we showed that the supplementation of high-dose IL-2, combined with TCR- and CD28-mediated stimulation, was sufficient to promote their in vitro proliferation (12). Cross-linking of antiCD3 and anti-CD28 antibodies either coupled to magnetic beads or presented by Fc-receptor expressing fibroblasts turned out to be important for their expansion as was supplementation of sufficient IL-2, due to their inability to produce this vital cytokine by themselves (13). However, once FOXP3 staining reagents became available, we discovered that only a portion of CD4+CD25high

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T cells (50–70%) maintained expression of this lineage-defining transcription factor during in vitro culture (14). When we further examined this subpopulation, we found that those cells also constitutively expressed CD62L and CCR7, suggesting that they originated from either naïve or central memory-type cells (15). Using CD45RA as a discriminating marker, we were able to identify the CD45RA+ subpopulation of Treg cells as their precursors (14). Treg cell cultures derived from this starting population homogeneously maintained FOXP3 expression, showed maximum suppressive activity in vitro, and did not secrete any proinflammatory cytokines. In contrast, the CD45RA– subpopulation of CD4+CD25high Treg cells lost FOXP3 expression over time, started to secrete cytokines, and showed diminished in vitro suppressive activity (14). In mouse, the stability of FOXP3 expression is at least in part determined by epigenetic mechanisms (16). In particular, DNA demethylation of a conserved noncoding region with known enhancer function in the FOXP3 locus is required for stable FOXP3 expression (termed TSDR for Tregspecific demethylated region) (17). We have recently confirmed that human Treg cells are also demethylated in this TSDR (18). However, whereas DNA methylation in this region increases in CD45RA– Treg cells after repetitive stimulation, CD45RA+ Treg cells maintain the demethylated state of their TSDR during in vitro culture (19). Thus, based on phenotypic, functional, molecular, and epigenetic data, CD45RA+ Treg cells seem to be the most reliable source for the generation of a homogeneous Treg cell product described to date. Even recently published alternative isolation strategies, such as depletion of CD127+ cells (20, 21) or selection of CD49d− cells (22) (and unpublished results), do not ensure the same quality of a Treg cell product after prolonged in vitro expansion (23). Thus, until we possess specific pharmacological agents that exclusively target Treg cells to enhance their suppressive activity in vivo, the isolation and expansion of CD45RA+ Treg cells seem the most promising strategy for the generation of safe and homogeneous Treg cell products for therapeutic purposes. Unfortunately, appropriate technologies for the reliable and GMP-compatible isolation of these rare Treg cell subpopulations are not yet available for clinical trials. However, several groups now collaborate with biotech and pharmaceutical companies for the implementation of such treatment strategies. Until then, the difficulties during the development of appropriate isolation and expansion techniques should not seduce us to employ insufficient technologies, since the health and safety of patients remains the prime priority of our efforts. With those principles in mind, we provide below detailed instructions for the identification, isolation, expansion, and functional characterization of Treg cell populations.

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2. Materials 2.1. MACSand FACS-Based Isolation of T Cell Subpopulations

1. Anti-PE and anti-CD4 magnetic beads (Miltenyi Biotec, Bergisch Gladbach, Germany). 2. MACS buffer: PBS containing 1% FCS, 2 mM EDTA. 3. MACS Midi magnets and LS columns. 4. MACS Pre-Separation Filters (Mesh size: 30 mM; Miltenyi Biotec). 5. FACS staining buffer: PBS containing 2% FCS. 6. Propidium iodide (SIGMA-Aldrich, Steinheim, Germany; final concentration: 2.5 mg/mL) for exclusion of dead cells in flow cytometry; kept as sterile 10× solution in PBS at 4°C; added immediately before sorting or analysis. 7. FACS Aria® high-speed cell sorter (BD Biosciences). 8. Antibodies used for cell isolation: Antibody

Fluorescent dye

Clone

Source

CD 4

FITC; PE-Cy7

SK3

BD Biosciences

CD 25

PE; PE-Cy7; APC

2A3

BD Biosciences

CD 45 RA

APC; Pacific Blue

MEM-56

Invitrogen

CD 45 RA

FITC

HI100

BD Pharmingen

CD 127

PE

hIL-7R-M21

BD Pharmingen

The color combination used routinely for isolation of naïve CD4+CD25+CD45RA+ Treg cells is indicated in bold (see also Note 1). 2.2. Cell Culture and Quality Control Assays

1. RPMI 1640 (without l-glutamine), supplemented with 2 mM L-glutamine, 10% FCS, 1% MEM vitamins, 1 mM sodium pyruvate, 1% MEM NEAA, 10 mM HEPES, 100 U/mL penicillin, 100 mg/mL streptomycin, 50 mM 2-mercaptoethanol. This medium is used for the maintenance of L929 feeder cells and for suppression assays. cRPMI for Treg cell expansion cultures is further supplemented with 300 U/mL human IL-2 (PROLEUKIN S, Novartis Pharma GmbH, Nürnberg, Germany). 2. 96-well and 24-well flat bottom plates (BD Falcon™, BD Biosciences, Heidelberg, Germany). 3. 96-well round bottom plates and cell culture flasks (25 and 75 cm2; Costar®, Corning, New York, NY). 4. PBS with 0.05%Trypsin, 0.02% EDTA (PAN, Aidenbach, Germany).

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5. PBS (w/o Ca2+ and Mg2+; PAA, Pasching, Germany). 6. NA/LE purified antibodies for cell culture: Anti-CD3: OKT3 (JANSSEN-CILAG GmbH, Neuss, Germany). Anti-CD28: CD28.2 (BD Pharmingen). 7. Magnetic beads used for in vitro expansion of T-cell subpopulations: Dynabeads® Human T-Activator CD3/CD28 (111.31D; Invitrogen). 8. Dynal magnets for bead removal (e.g., MPC™-L or MPC™-15). 9. Phorbol 12-myristate 13-acetate (PMA; SIGMA-Aldrich) is dissolved at 2 mg/mL in DMSO and stored in aliquots at −20°C. 10. Ionomycin (Alexis, Lausen, Switzerland) is stored as 1 mM solution in DMSO at −20°C. 11. CFSE (carboxyfluorescein succinimidyl ester; SIGMA-Aldrich Cat No.:21888) is stored in aliquots as 5 mM stock solution in DMSO at −20°C. 12. FOXP3 staining kit (includes fixation/permeabilization buffers and PE-, APC-, or Pacific Blue-labeled anti-FOXP3 antibidy; clone:PCH101 [eBioscience, San Diego, CA]).

3. Methods 3.1. Isolation of Human CD4+CD25 high Treg Cells

1. PBMC are isolated from leukapheresis products of healthy volunteers (after their informed consent and in accordance with protocols approved by the local authorities) by density gradient centrifugation over Ficoll/Hypaque (Biocoll, Biochrom AG, Berlin, Germany). 2. Cells are stained with PE-anti-CD25 (10 × 106 cells/5 mL PE-anti-CD25/100 mL FACS buffer) for 20 min at 4°C, followed by incubation with anti-PE magnetic beads (Miltenyi Biotech) in MACS buffer and positive selection of CD25+ cells on LS columns according to the manufacturer’s instructions. 3. Isolated cells are washed once in FACS buffer and stained with FITC-anti-CD4 (4 × 106 cells/10 mL FITC-antiCD4/100 mL FACS buffer) for 20 min at 4°C. 4. Stained cells are washed once, filtered through a 30 mm mesh filter, and resuspended in FACS buffer at a cell density of 10–20 × 106 cells/mL.

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5. Cells are sorted on a FACS Aria® high-speed cell sorter into 5-mL- or 15-mL tubes filled with 200 mL or 500 mL FCS, respectively. Gating strategy see Fig. 2. Sterile PI is added immediately before sorting. 6. Sorted cells are centrifuged at 350 × g for 10 min, resuspended in cRPMI, counted, and can be used either directly or after short-term expansion in functional assays (see Note 2). 3.2. Isolation of Human CD4+CD25+CD127− Treg Cells

1. As in Subheading 3.1. 2. Cells are incubated with anti-CD4 magnetic beads (Miltenyi Biotec) in MACS buffer and CD4+ cells are positively selected on LS columns according to the manufacturer’s instructions. 3. Isolated cells are washed once in FACS buffer and stained with FITC-anti-CD4, PE-anti-CD127, and APC-anti-CD25 (4 × 106 cells/10 mL FITC-anti-CD4/5 mL PE-CD127/2 mL APC-anti-CD25/100 mL FACS buffer) for 20 min at 4°C. 4. Steps 4–6: As in Subheading 3.1. For Gating strategy see Fig. 1 (see Note 3).

3.3. Isolation of Human CD4+CD25highCD45RA+ Naïve Treg Cells

1. As in Subheading 3.1. 2. As in Subheading 3.1. 3. Isolated cells are washed once in FACS buffer and stained with FITC-anti-CD4 and APC-anti-CD45RA (4 × 106 cells/ 10 mL FITC-anti-CD4/1.5 mL APC-anti-CD45RA/100 mL FACS buffer) for 20 min at 4°C. 4. Steps 4–6: As in Subheading 3.1. For Gating strategy see Fig. 2 below.

3.4. In Vitro Expansion of Human CD4+CD25+ Treg Using Murine L929 Fibroblasts 3.4.1. Maintenance of the HuCD32+ Murine Fibroblast Cell Line L929

1. Cells of the L 929-derived murine Ltk− cell line stably transfected with human FcgRII (CD32) (24) are grown in 25 or 75 cm2 flasks in cRPMI to subconfluent monolayers. 2. To harvest the cells, medium is discarded; trypsin/EDTA is added (1–2 mL for 25 cm2 flask, 3–3.5 mL to 75 cm2 flask) and incubated at 37°C (incubator) for 5 min. 3. Cells are detached by thorough tapping against the side wall of the flask. 4. 10 mL of cRPMI (5 mL for 25 cm2 flask) are added and cells are thoroughly mixed. 5. Cells are transferred to a 50-ml cell culture tube (BD Falcon™); the tube is filled with cRPMI and centrifuged at 300 × g for 10 min. 6. Pelleted cells are resuspended in cRPMI, counted, and new flasks are set up with 0.25 to 1 × 106 cells/20 mL cRPMI/ 75 cm2 flask.

FSc

94.6

76.1

99.3

99.7

99.4

50.4

CD25

CD4

PI

SSc

CD127

94.5

8.3

FOXP3

93.7

7.3

Fig. 1. Isolation of CD4+CD25+CD127– Treg cells (a) Gating strategy for, and FOXP3 expression of, CD4+CD25+CD127– Treg cells within human PBMC. (b) Reanalysis of CD4+CD25+CD127– Treg cells, isolated according to the protocol detailed in Subheading 3.2.

b

a

Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells 21

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CD4+Cells

All Cells

a 1.8

4.9 50.6

PBMC

33.5

All Cells

b 98.2

98.4 0.9

Sorted + RA Treg

98.3

c 99.8

97.1 0.0

CD25

99.4

Sorted RA− ฀ Treg

CD4

CD45RA

FOXP3

Fig. 2. Isolation of CD4+CD25+CD45RA+ and CD4+CD25+CD45RA– Treg cells. (a) MNC from a leukapheresis product are stained with FITC-anti-CD4, PE-anti-CD25, and APC-anti-CD45RA. To isolate the entire CD4+CD25high Treg population (as mentioned in Subheading 3.1), the gate is set on the population with a CD4 expression level slightly lower than that of the CD4+CD25int T cell population (representing recently activated Tconv cells) and with a CD25 expression level above that of the main CD4–CD25+ population (representing mostly activated B cells). To further separate Treg cells into a naive (CD45RA+) and a memory-type (CD45RA–) subpopulation, gates are set as shown in the middle panel. (b, c) Reanalysis of sorted CD4+CD25+CD45RA+ (b) and CD4+CD25+CD45RA– Treg cells (c). In parallel, MNC as well as aliquots of the sorted populations are stained with FITC-anti-CD4, PE-anti-CD25, APC-anti-CD45RA, and Pacific Blue-anti-FOXP3 to identify Treg cells (and verify successful sorting) via FOXP3 expression (see Notes 4 and 5). 3.4.2. Preparation of HuCD32+ L929 Feeder Cells

1. HuCD32+ L929 cells are harvested, washed once, counted, and cell number is adjusted to 5 × 106 cells/mL in cRPMI 2. Cells are irradiated with 70 Gy. 3. Immediately after irradiation, cells are diluted to appropriate cell concentration in cRPMI and seeded to cell culture plates: 15,000 cells/100 mL in 96-well flat bottom plates. 80,000 cells/400 mL in 24-well plates. 380,000 cells/2.5 mL in 6-well plates.

Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells

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4. Cells are allowed to adhere (approx. 60 min at 37°C) before T cells are added. Feeder cells can be prepared 1 day prior to T-cell culture setup. 1. 96-well flat bottom plates with adherent feeder cells in cRPMI are prepared (see Subheading 3.4.2, step 2).

3.4.3. Setup and Maintenance of Treg Cell Cultures on L929 Feeder Cells

2. Sorted CD4+CD25+ Treg cells or subpopulations thereof are seeded at 10,000 cells/well in 100 mL cRPMI, supplemented with 600 U/mL IL-2, 200 ng/mL OKT-3, and 200 ng/mL anti-CD28 (resulting in final concentrations for IL-2, OKT3, and anti-CD28 of 300 U/mL, 100 ng/mL, and 100 ng/ mL, respectively). 3. Cells are incubated at 37°C and 5% CO2 in humidified atmosphere. 4. On day 4, 100 mL/well is removed and replaced by fresh cRPMI with 300 U/mL IL-2. 5. On day 6, cells are harvested; cells from parallel cultures are united, washed once in cRPMI, counted, and reseeded to either 24-well or 6-well plates at 100,000 cells and 500,000 cells/well in a final volume of 500 mL and 3 mL, respectively. IL-2, OKT-3, and anti-CD28 are added to result in final concentrations as stated in step 2. 6. Timeline for the maintenance of long-term Treg/feeder cell cultures:

start

feeding

restimulation

feeding

restimulation

d0

d4

d6

d8 / d9

d11

feeding

d15 or d22 or d29…

restimulation

d18 or d25 or d32…

7. Routinely obtained expansion rates for FACS-purified human CD4+CD25+CD45RA+ Treg cells kept on L929 feeder cells for up to 25 days are shown in Fig. 3. 3.5. In Vitro Expansion of Human Treg Cells Using AntibodyCoated Bead Stimulation

1. Sorted Treg cells in cRPMI/300 U/mL IL-2 are mixed with Dynabeads® Human T-Activator CD3/CD28 to result in a bead:cell ratio of 4:1. 2. 10,000 cells/100 mL/well are seeded in 96-well round bottom plates. The plates are kept at 4°C for approx. 1 h before culture setup to minimize adherence of the seeded cells to the walls of the well and thereby facilitate cell accumulation at the bottom of the well.

Hoffmann, Eder, and Edinger 10000

1000

x-fold Expansion

24

100

10

1 0

7

14

21

28

Culture period (d) Fig. 3. Expansion of CD4+CD25+CD45RA+ Treg cells on huCD32+ L929 feeder cells. CD4+CD25+CD45RA+ Treg cells were sorted from 11 different leukapheresis products and cultured for up to 25 days on huCD32+ L929 feeder cells as detailed in Subheading 3.4. Cells were harvested and counted on indicated days using Trypan Blue for dead cell exclusion. Cumulative expansion rates were calculated. Data represent mean ± SD from 11 individual cultures.

3. Cells are incubated at 37°C and 5%CO2 in humidified atmosphere. 4. On day 4, 100 mL cRPMI with 300 U/mL IL-2 are added to each well. 5. On day 7, cells are resuspended within the wells (cells during the first week of in vitro culture tend to stick together tightly and can only be singularized while still in the well), cells from parallel wells are united in a 5-mL or 15-mL cell culture tube, thoroughly resuspended, and placed in a magnet holder for 2–3 min. 6. Cells are removed from the tubes, transferred to a new tube, counted, washed once with cRPMI, and resuspended at 5 × 105/mL in cRPMI/300 U/mL IL-2 with stimulatory beads added to result in a bead:cell ratio of 1:1. 7. 1 mL/well of the cell and bead mixture is added to the wells of a 24-well plate. 8. On day 11, 1 mL cRPMI/300 U/mL IL-2 is added to each well. If necessary, wells can be split into two and refilled to 1 mL final volume with cRPMI/300 U/mL IL-2. 9. On day 14, cells are harvested and restimulated as detailed in step 6 (see Note 6). 10. Timeline for the maintenance of long-term Treg/Bead cultures

Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells

start

feeding

restimulation

feeding

restimulation

d0

d4

d7

d11

d14

1000

feeding

d18 or d25

25

restimulation

d21 or d28

10000

a

b

x-fold Expansion

1000 100

100 10 10

1

1 0

7

14

21

0

7

14

21

Culture period (d) Fig. 4. Expansion of CD4+CD25+CD45RA+ Treg using antibody-coated magnetic beads. CD4+CD25+CD45RA+ Treg cells were sorted from four different leukapheresis products and cultured for up to 21 with anti-CD3/anti-CD28-coated paramagnetic beads as detailed in Subheading 3.5. Cells were harvested and counted on indicated days with Trypan Blue for exclusion of dead cells. Cumulative expansion rates were calculated. Data represent mean ± SD from four individual cultures.

11. Routinely achieved expansion rates for FACS-purified CD4+CD25+CD45RA+ Treg cells cultured with anti-CD3/ anti-CD28 coated stimulatory beads are shown in Fig. 4. 3.6. Quality Control During and at End of Culture 3.6.1. Staining of In Vitro Expanded Treg Cells for FOXP3 with or without Simultaneous Staining for Proinflammatory Cytokines

FOXP3 expression is still the most widely used and generally accepted quality control criterion for in vitro-expanded Treg cells. However, since FOXP3 can also be transiently expressed by activated Tconv cells, a short resting period before staining for FOXP3 is recommended. Alternatively, expanded cells can be simultaneously stained for FOXP3 and proinflammatory cytokines to reveal any cells converting from a Treg to a Tconv phenotype during in vitro culture.

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1. Expanded cells are harvested (in case expansion was done with stimulatory beads, the beads are removed), washed once in cRPMI, and reseeded at 1 × 106 cells/mL in cRPMI/300 U/ mL IL-2 in 6-well plates (5 mL/well) or 24-well plates (1 mL/well) for 2–4 days (see Note 7). 2. Variation 1. Cells are stained directly for FOXP3 (can be combined with surface staining for CD4 and/or CD25) following the manufacturer’s instructions. Variation 2. Cells are washed once with cRPMI, reseeded at 1 × 106 cells/mL in cRPMI in 24-well plates (1 mL/well), and stimulated with PMA (20 ng/mL)/ionomycin (1 mM) in the presence of monensin (GolgiStop; BD Biosciences; 0.67 mL/mL) for 5 h. Cells are washed once and are now ready for combined FOXP3/intracellular cytokine staining. A common combination would be CD4 (surface staining performed before fixation/permeabilization), FOXP3, IL-2 and/or IFN-g (anti-cytokine and anti-FOXP3 antibodies are added simultaneously; the protocol for FOXP3 staining should be strictly applied). Typical results with both quality control staining protocols are shown in Fig. 5. 3.6.2. Functional Analysis of In Vitro Expanded Treg Cells: Suppression of Autologous Tconv Cells After Polyclonal Stimulation

Quality control of in vitro expanded Treg cells can also be carried out by determination of their suppressive activity after polyclonal activation. Commonly, proliferation of freshly isolated or cryopreserved autologous Tconv cells is used as readout system. Since in vitro expanded Treg cells show a higher tendency than freshly isolated Treg cells to proliferate under normal assay conditions, a CFSE dilution assay is preferable to detection of 3H-thymidine incorporation for quantification of the proliferative capacity of cocultured Tconv cells. 1. Cryopreserved PBMC are thawed and CD4+CD25− Tconv cells are isolated by either MACS- or FACS-based techniques as detailed above. 2. All CFSE labeling steps should be done with the lights in the laminar flow turned off. 3. 107 Tconv cells in a 15-mL conical tube are washed twice with PBS (w/o FCS!) and resuspended in 500 mL PBS (see Note 8). 4. 500 mL of a 2× CFSE solution (4 mM in PBS) is added and cells are incubated for 4 min. at room temperature in the dark. 5. Cells are washed twice with PBS/10% FCS or cRPMI, counted, and adjusted to 1 × 106/mL in cRPMI. 6. In vitro expanded and 2–4 days-rested Treg cells are washed once and adjusted to 1 × 106 cells/mL in cRPMI. If Treg cells are to be titrated, 1:2 serial dilutions are prepared.

Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells −

RA+ Treg

a

RA Treg

104

104

103

FOXP3

27

103

99.3

102

102

101

101

100

58.4

100 0

200

400

600

800

1000

0

200

400

600

800

1000

FSc b

104

104

FOXP3

94.4

0.6

103

103

102

102

101

101

4.8

100 100

0.2 101

102

103

100 104

100

65.2

4.9

14.3

15.6 101

102

103

104

IL-2 Fig. 5. FOXP3 and intracellular cytokine expression in in vitro expanded CD45RA+ and CD45RA– Treg cells. (a) CD4+CD25+CD45RA+ and CD4+CD25+CD45RA– Treg cells were isolated by MACS/FACS, cultured in vitro on L929 feeder cells for 2 weeks and rested for 4 days in cRPMI/300 U/mL before staining for FOXP3. (b) Cells prepared and treated as in (a) were stained for FOXP3 and intracellular IL-2 after an additional 5 h-stimulation period with PMA/ionomycin in the presence of monensin.

7. Antigen-presenting cells (APC) are prepared from CD4depleted PBMC (step 1) by incubation with anti-CD2 beads (e.g., Dynabeads® CD2 Pan T; Invitrogen) following the manufacturer’s instructions. CD2-depleted cells are washed once in cRPMI, irradiated with 30 Gy, and adjusted to 2 × 106 cells/mL in cRPMI.

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8. Cultures are set up in 96-well round bottom plates in the following order (see also Notes 9–11): − 50 mL APC (→ 100,000 cells/well). − 50 mL OKT3 (400 mg/mL → 100 mg/mL final conc.). − 50 mL Tconv cells (→ 50,000 cells/well). − 50 mL Treg cells (→ a maximum of 50,000 cells/well). 9. Cultures are kept at 37°C, 5% CO2 in a humidified atmosphere for 3 days. 10. Cells are harvested, those from parallel wells are united, washed once in FACS staining buffer, stained, for example, with CD4 and CD25, and analyzed by FACS.

4. Notes 1. Staining for CD25 should not be performed with FITClabeled antibodies, as discrimination of CD25high and CD25int CD4 T cell populations is difficult due to low resolution. Also, we obtained better results with clone 2A3 as compared to clone M-A251 (both BD Biosciences). 2. As detailed in the introduction, bulk CD4+CD25high Treg cells comprise CD45RA+ (naïve) as well as CD45RA– (memory-type) cells. Both represent Treg cell populations, as demonstrated by phenotypic and epigenetic analysis. However, since they differ considerably in Treg phenotype stability during in vitro culture, Treg cells isolated solely on the basis of CD4 and CD25 expression should only be used directly or after short-term in vitro activation/expansion. 3. Since the CD4+CD25+CD127– Treg cell population also comprises naïve as well as memory-type cells, the same caution should be exercised in applying these cells as detailed above for bulk CD4+CD25+ Treg cells. 4. Another characteristic of Treg cells that can be used for their identification and flow-cytometric isolation in combination with staining for surface markers is their smaller size (hence the lower CD4 expression mentioned in Fig. 2), resulting in a lower forward scatter signal. 5. CD45RA and CD45RO expression levels are inversely correlated, with most CD4 T cells expressing both isoforms of the common leukocyte antigen (CD45) to a certain degree. However, only CD4+ Treg cells with a high CD45RA expression level are also homogeneously positive for the two additional developmental markers CCR7 and CD62L, which are usually associated with a naïve phenotype and have been

Polyclonal Expansion of Human CD4+CD25+ Regulatory T Cells

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shown by us before to identify those Treg cells with a stable suppressor/regulatory phenotype during in vitro cultivation. 6. The bead:cell ratio of 4:1 in the first week and then 1:1 for all subsequent restimulation rounds is optimized for the needs of Treg cells, but are too strong for the expansion of conventional (CD4+CD25–) T cells (Tconv). For those cells, a 1:1 ratio for the first week and a 1:10 ratio thereafter turned out to be more successful. 7. In vitro expanded T cells with a stable Treg phenotype (e.g., CD4+CD25+CD45RA+ Treg cells) do not form clusters when kept under these conditions and show only very little proliferation. In contrast, in vitro-expanded Tconv cells tend to aggregate in small clusters and retain considerable proliferative capacity even after removal of the TCR stimulus (e.g., antibody-coated beads). 8. Volume can be adjusted to actual cell number but should be no less than 100 mL. 9. Several negative and positive controls should be included in CFSE assays: (a) cultures with APC and CFSE-labeled Tconv w/o OKT3 (→ useful for exact identification of undiluted CFSE label intensity as most of the Tconv cells will survive but not proliferate under these conditions), (b) in vitroexpanded CD4+CD25– T cells should be added instead of Treg cells to discriminate Treg-cell-specific suppression from unspecific effects of in vitro culture-adapted cells. 10. CD8+ instead of CD4+ T cells can also be used as responders in Treg suppression assays. This facilitates discrimination of strongly proliferating (and thus CFSE-losing) responder cells from unlabeled Treg cells. 11. A minimum of five parallel wells/intended FACS staining combination in recommended. References 1. Josefowicz SZ, and Rudensky A (2009) Control of regulatory T cell lineage commitment and maintenance. Immunity 30:616–25. 2. Zou W (2006) Regulatory T cells, tumour immunity and immunotherapy. Nat Rev Immunol 6:295–307. 3. Belkaid Y, and Tarbell K (2009) Regulatory T cells in the control of host–microorganism interactions. Annu Rev Immunol 27:551–89. 4. Sakaguchi S, Yamaguchi T, Nomura T, and Ono M (2008) Regulatory T cells and immune tolerance. Cell 133:775–87. 5. Mottet C, Uhlig HH, and Powrie F (2003) Cutting edge: cure of colitis by CD4+CD25+ regulatory T cells. J Immunol 170:3939–43.

6. Feng G, Chan T, Wood KJ, and Bushell A (2009) Donor reactive regulatory T cells. Curr Opin Organ Transplant 14:432–8. 7. Hoffmann P, and Edinger M (2006) CD4+CD25+ regulatory T cells and graftversus-host disease. Semin Hematol 43:62–9. 8. Edinger M, Hoffmann P, Ermann J, Drago K, Fathman CG, Strober S, and Negrin RS (2003) CD4(+)CD25(+) regulatory T cells preserve graft-versus-tumor activity while inhibiting graft-versus-host disease after bone marrow transplantation. Nat Med 9:1144–50. 9. Hoffmann P, Ermann J, Edinger M, Fathman CG, and Strober S (2002) Donor-type

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11.

12.

13.

14.

15.

16.

17.

Hoffmann, Eder, and Edinger CD4(+)CD25(+) regulatory T cells suppress lethal acute graft-versus-host disease after allogeneic bone marrow transplantation. J Exp Med 196:389–99. Hoffmann P, Boeld TJ, Eder R, Albrecht J, Doser K, Piseshka B, Dada A, Niemand C, Assenmacher M, Orso E, Andreesen R, Holler E, and Edinger M (2006) Isolation of CD4(+) CD25(+) regulatory T cells for clinical trials. Biol Blood Marrow Transplant 12:267–74. Trzonkowski P, Bieniaszewska M, Juscinska J, Dobyszuk A, Krzystyniak A, Marek N, Mysliwska J, and Hellmann A (2009) First-inman clinical results of the treatment of patients with graft versus host disease with human ex vivo expanded CD4+CD25+CD127- T regulatory cells. Clin Immunol 133:22–6. Hoffmann P, Eder R, Kunz-Schughart LA, Andreesen R, and Edinger M (2004) Largescale in vitro expansion of polyclonal human CD4(+)CD25high regulatory T cells. Blood 104:895–903. Thornton AM, and Shevach EM (1998) CD4+CD25+ immunoregulatory T cells suppress polyclonal T cell activation in vitro by inhibiting interleukin 2 production. J Exp Med 188:287–96. Hoffmann P, Eder R, Boeld TJ, Doser K, Piseshka B, Andreesen R, and Edinger M (2006) Only the CD45RA+ subpopulation of CD4+CD25high T cells gives rise to homogeneous regulatory T-cell lines upon in vitro expansion. Blood 108:4260–7. Sallusto F, Geginat J, and Lanzavecchia A (2004) Central memory and effector memory T cell subsets: function, generation and maintenance. Annu Rev Immunol 22: 745–63. Kim HP, and Leonard WJ (2007) CREB/ ATF-dependent T cell receptor-induced FoxP3 gene expression: a role for DNA methylation. J Exp Med 204:1543–51. Floess S, Freyer J, Siewert C, Baron U, Olek S, Polansky J, Schlawe K, Chang HD, Bopp T, Schmitt E, Klein-Hessling S, Serfling E, Hamann A, and Huehn J (2007) Epigenetic control of the foxp3 locus in regulatory T cells. PLoS Biol 5:e38.

18. Baron U, Floess S, Wieczorek G, Baumann K, Grutzkau A, Dong J, Thiel A, Boeld TJ, Hoffmann P, Edinger M, Turbachova I, Hamann A, Olek S, and Huehn J (2007) DNA demethylation in the human FOXP3 locus discriminates regulatory T cells from activated FOXP3(+) conventional T cells. Eur J Immunol 37:2378–89. 19. Schmidl C, Klug M, Boeld TJ, Andreesen R, Hoffmann P, Edinger M, and Rehli M (2009) Lineage-specific DNA methylation in T cells correlates with histone methylation and enhancer activity. Genome Res 19:1165–74. 20. Liu W, Putnam AL, Xu-Yu Z, Szot GL, Lee MR, Zhu S, Gottlieb PA, Kapranov P, Gingeras TR, de St Groth BF, Clayberger C, Soper DM, Ziegler SF, and Bluestone JA (2006) CD127 expression inversely correlates with FoxP3 and suppressive function of human CD4+ T reg cells. J Exp Med 203:1701–11. 21. Seddiki N, Santner-Nanan B, Martinson J, Zaunders J, Sasson S, Landay A, Solomon M, Selby W, Alexander SI, Nanan R, Kelleher A, and Fazekas de St Groth B (2006) Expression of interleukin (IL)-2 and IL-7 receptors discriminates between human regulatory and activated T cells. J Exp Med 203:1693–700. 22. Kleinewietfeld M, Starke M, Di Mitri D, Borsellino G, Battistini L, Rotzschke O, and Falk K (2009) CD49d provides access to “untouched” human Foxp3+ Treg free of contaminating effector cells. Blood 113: 827–36. 23. Hoffmann P, Boeld TJ, Eder R, Huehn J, Floess S, Wieczorek G, Olek S, Dietmaier W, Andreesen R, and Edinger M (2009) Loss of FOXP3 expression in natural human CD4+CD25+ regulatory T cells upon repetitive in vitro stimulation. Eur J Immunol 39:1088–97. 24. Peltz GA, Trounstine ML, and Moore KW (1988) Cloned and expressed human Fc receptor for IgG mediates anti-CD3-dependent lymphoproliferation. J Immunol 141:1891–6.

Chapter 3 Methods for In Vitro Generation of Human Type 1 Regulatory T Cells Silvia Gregori, Maria Grazia Roncarolo, and Rosa Bacchetta Abstract Type 1 regulatory T (Tr1) cells are adaptive regulatory T cells that are induced in the periphery upon chronic exposure to antigen (Ag) in a tolerogenic environment containing interleukin (IL)-10. Tr1 cells are Ag-specific; they produce high levels of IL-10 and TGF-b in the absence of IL-4 and suppress T-cell responses via a cytokine-dependent mechanism. During the last decade, several protocols have been developed to generate Tr1 cell lines in vitro. In this chapter, we outline protocols to generate non-Ag- and Ag-specific Tr1 cell lines and assays used to characterize Tr1 cell phenotype and functions. Key words: Type 1 regulatory T cells, Dendritic cells, Interleukin-10

1. Introduction Regulatory T (Tr) cells are a specialized subset of T cells critically involved in promoting and maintaining peripheral tolerance. Differentiation of Tr cells occurs both in the thymus and in the periphery. Thymus-derived Tr cells are defined as naturally occurring Tr (nTr) cells and strictly depend on FOXP3 expression for their function (1, 2). Adaptive Tr cells, generated in the periphery in tolerogenic microenvironments, comprise different Tr subsets and suppress T-cell responses via immunomodulatory cytokines (3). Among them, the better characterized are the adaptive type 1 regulatory T (Tr1) cells that depend on IL-10 for their generation and functions (4). Tr1 cells are defined by the ability to produce high levels of IL-10 and TGF-b, low amounts of IFN-g and IL-2, and detectable levels of IL-5, in the absence of IL-4 (5–7). Unlike nTreg cells, Tr1 cells are independent from FOXP3 expression for generation Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_3, © Springer Science+Business Media, LLC 2011

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((8–10), and Passerini et al., manuscript submitted). Similar to other Treg cells, Tr1 cells are anergic, but their proliferation can be restored by IL-2 and IL-15, independently from TCRmediated activation (7). A variety of transcription factors and membrane-bound markers have been described to identify Tr1 cells. In mouse models, a comparative study between Tr1 and nTr cells demonstrated the selective expression of repressor of GATA-3 (ROG) by Tr1 cells (11). Moreover, Tr1 cells induced in vitro in the presence of IL-27 and TGF-b express c-Maf (12). Human IL-10-producing T cells, resembling Tr1 cells, have been shown to selectively express coinhibitory molecule ICOS (12, 13) or adhesion molecule CD18 (14). However, none of these markers has been confirmed in different experimental systems; therefore, so far no molecular marker that specifies Tr1 cells has been found. Tr1 cells are induced in an Ag-specific manner; therefore, in contrast to other Treg cells, they are Ag specific. Once activated through their specific TCR, Tr1 cells secrete IL-10 and TGF-b that directly inhibit effector T cells proliferation and expression of MHC class II and costimulatory molecules on antigen-presenting cells (APC), which indirectly suppress effector T cells activation. It has been shown that IL-10-producing Tr1-like cells suppress T-cell responses via perforin and granzyme B (15). We have recently demonstrated that Tr1 cell lines and cell clones express and secrete high levels of granzyme B and can mediate suppression of T cell responses by selectively lysed myeloid cells (Magnani et al., manuscript under preparation). In vitro repetitive stimulations via TCR of naïve CD4+ T cells in the presence of IL-10 can be used to generate Tr1 cells (6). We demonstrated that IFN-a, but not TGF-b, synergizes with IL-10 in promoting generation of human polyclonal Tr1 cells (16). In alternative to exogenous IL-10, other factors have been shown to promote the induction of polyclonal human IL-10producing T cells resembling Tr1 cells: TCR triggering of naïve T cells in the presence of vitamin D3 and dexamethasone (8, 17), and activation of CD4+ T cells with CD3/CD46 cross-linking (18). Notably, induction of IL-10-producing T cells using the former methods is dependent on IL-10. However, it has been recently demonstrated that IL-27 can be an alternative to IL-10 for promoting human IL-10-producing Tr1-like cells differentiation (19). It remains to be determined whether IL-10-producing Treg cells generated with different methods are the same cells. Antigen(Ag)-specific human Tr1 cells can be isolated in vitro by Ag restimulation of CD4+ T cells from peripheral blood of immunized subjects, after in vivo rechallenge (7). Alternatively, allergen-specific Tr1 cells can be generated in vitro by stimulating human T cells with autologous tolerogenic dendritic cells (DC) pulsed with allergen (20). In addition, allo-Ag-specific Tr1 cells can be induced in vitro by stimulating CD4+ T cells with

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allogeneic APC in the presence of exogenous IL-10 ((7, 21), and 22). Although exogenous IL-10, in the presence of the specific Ag, is critical for Tr1 cell induction, it is not sufficient to de novo differentiate them in high numbers, in the absence of APC (16). Several reports indicate that in vitro T-cell priming by immature DC, or DC rendered tolerogenic with biological/pharmacological agents (23), drives the differentiation of Tr1 cells. We and others have reported that repetitive stimulations of human naïve CD4+ T cells with immature DC drive the in vitro generation of Tr1 cells via an IL-10-dependent mechanism (24, 25). Recently, we have identified and characterized a novel population of human tolerogenic DC, termed DC-10, which are particularly suitable for Tr1 cell induction. In addition to secreting high levels of IL-10,but low amounts of IL-12, DC-10 express high levels of the tolerogenic molecules ILT2, ILT3, ILT4, and membrane-bound HLA-G1. Their phenotype is stable in vitro since it is not modified by activation, indicating that once exposed to viral or bacterial antigens, DC-10 maintained their tolerogenic potential. DC-10 are inducible in vitro from peripheral blood monocytes in the presence of IL-10, but importantly, DC-10 are present in vivo in peripheral blood and in secondary lymphoid tissues underlying their physiological role in immunohomeostasis (26). During the last decade, Tr cell-based therapy has become an attractive nonpharmacological therapeutic option for immunomodulation in different pathological settings. Therefore, much effort has been devoted to the development of ad hoc methods to isolate and to expand or generate Tr cells in vitro for clinical application (9). Several studies demonstrated the efficacy of nTr or Tr1 cell therapy in treating immunomediated pathologies in preclinical models. Specifically, adoptive transfer of Tr1 cells has been proven to be effective in controlling inflammatory bowel disease (6, 27), colitis (28), experimental encephalomyelitis (17), graft vs. host disease (29), and islet allograft rejection (30) and (31). We focused our work on the definition of suitable protocols to generate in vitro Ag-specific Tr1 cells for cell therapy. A reproducible manufacturing protocol for the GMP production of allo-specific anergic T cells, using either exogenous IL-10 or tolerogenic DC-derived IL-10, has been developed. A clinical protocol of immunotherapy with ex vivo IL-10-anergized cells of donor origin in patients receiving haploidentical HSCT is nearly completed at the San Raffaele Hospital (http://www.risetfp6.org), confirming the safety and the feasibility of the proposed approach (Bacchetta et al., manuscript submitted). In addition, a different manufacturing protocol for the GMP production of human ovalbumin-specific Tr1 cells using artificial APC has been established (32). This protocol allows the expansion of autologous ovalbumin-specific Tr1

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cell clones for the treatment of severe forms of Crohn’s disease within a clinical trial started in March 2008. In this chapter, we outline protocols suitable for in vitro generation of human polyclonal and Ag-specific Tr1 cells and experimental assays used to characterize the resulting Tr1 cells.

2. Materials 2.1. Cell Culture and Activation Reagents

1. RPMI 1640 (BioWhittaker, Verniers, Belgium) supplemented with 10% FBS (BioWhittaker), 100 U/ml penicillin/streptomycin, and 2 mM glutamine with or without 2-mercaptoethanol 50 nM. 2. X-VIVO 15 (BioWhittaker) supplemented with 5% HS (BioWhittaker) and 100 U/ml penicillin/streptomycin. 3. TPA is diluted in X-VIVO 15 (BioWhittaker) supplemented with 5% HS (BioWhittaker) and 100 U/ml penicillin/streptomycin at 1 mg/ml. 4. Ionomycin is diluted in X-VIVO 15 (BioWhittaker) supplemented with 5% HS (BioWhittaker) and 100 U/ml penicillin/ streptomycin at 1 mg/ml. 5. Proleukin (IL-2) (Chiron Corporation, Emeryville, CA) is added to X-VIVO 15 (BioWhittaker) supplemented with 5% HS (BioWhittaker) and 100 U/ml penicillin/streptomycin at the required concentrations. 6. Anti-CD3 mAb (clone OKT3) (Orthoclone Janssen-Cilag, Cologno Monzese, Italy) is diluted in PBS at the 10 mg/ml. 7. Anti-CD28 mAb (clone V5T CD28.05) (BD Bioscience) is diluted in X-VIVO 15 (BioWhittaker) supplemented with 5% HS (BioWhittaker) and 100 U/ml penicillin/streptomycin at the required concentrations. 8. (3H)-thymidine ((3H)-thy) (Amersham Biosciences, Uppsala, Sweden) diluted in X-VIVO 15 (BioWhittaker) supplemented with 5% HS (BioWhittaker) and 100 U/ml penicillin/streptomycin at 1 mCi/ml. 9. CFSE, 5-(and-6)-carboxyfluorescein diacetate succinimidyl ester (CFSE; Molecular Probes, Eugene, OR) is diluted in PBS at 20 mM. Working solution is 5 mM. 10. Leukocyte Activation Cocktail (BD Pharmingeu) containing PMA, ionomycin and Brefeldim A.

2.2. Cytokine Detection and Cytofluorimetric Analyses

1. PBS (Euroclone Life Sciences Division, Pero, Italy) with or without 2 or 5% FBS. 2. Anti-IL-10 capture mAbs (clone M010, Endogen, Pierce, Rockford, USA) is diluted in PBS supplemented with 5% FBS at 5 mg/ml.

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3. Anti-IL-10 detection mAbs (clone M011B, Endogen, Pierce, Rockford, USA) is diluted in PBS supplemented with 5% FBS at 1 mg/ml. 4. PE-labeled anti-hIL-4, anti-hIL-2, or anti-hIL-10, and FITCcoupled anti-hIFN-g (BD Bioscience) are diluted in FBS supplemented with 2% FBS. 5. FACS antibodies: Anti-CD14 FITC/PE conjugated, antiCD83 PE conjugated, anti-CD80 PE conjugated, anti-CD86 PE conjugated, anti-HLA-DR PE conjugated, and anti-CD11c PE conjugated (BD Bioscience, San Diego, CA) are diluted in PBS supplemented with 2% FBS.

3. Methods The methods described below outline: (1) induction of polyclonal Tr1 cell lines, (2) induction of allo-specific Tr1 cell lines using monocyte and IL-10, (3) induction of tolerogenic DC-10, (3) biological characterization of tolerogenic DC-10, (4) induction of Tr1 cell lines using DC-10, and (5) biological characterization of Tr1 cell lines. 3.1. Induction of Polyclonal Tr1 Cells 3.1.1. CD4+ T Cell Preparation

3.1.2. Differentiation of Polyclonal Tr1 Cell Lines

Naïve CD4+ T cells can be selected from either umbilical cord blood or peripheral blood. Isolation of these cells requires two consecutive steps: (1) isolation of total CD4+ T cells by negative selection and (2) negative selection of CD45RA+ T cells. Peripheral blood mononuclear cells (PBMC) are obtained by centrifugation over Lymphoprep gradients, and total CD4 + T cells are separated using the CD4+ T cell isolation kit II, strictly following the manufacturer’s instructions (see Note 1). CD4+CD45RO− T cells are negative-selected using with magnetic beads coupled to anti-CD45RO mAb (see Note 2). The phenotype of the resulting cells is routinely greater than 90% CD4+CD45ROCD45RA+. Polyclonal Tr1 cell lines can be generated using murine L cells expressing hCD32 (FCRII), hCD58 (LFA-3), and hCD80 as surrogate APC. Adherent L cells cultured in RPMI 1640 supplemented with 10% FBS, 100 U/ml penicillin/streptomycin, and 2 mM glutamine are detached from the plastic by incubation with trypsin-EDTA and irradiated (7,000 rad). After washing, plate L cells in 24-well plates at an initial density of 2 × 105 cells/ml in 500 ml of complete medium (X-VIVO 15 supplemented with 5% HS and 100 U/ml penicillin/streptomycin). Let L cells remain for 2 h at 37°C and then add 500 ml of naïve CD4+ T cells at an initial density of 2 × 105 cells/ml in complete medium containing rhIL-2 (100 U/ml), and rhIL-15 (1 ng/ml), and 0.1 mg/ml of

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anti-CD3 mAb as previously described (16). Tr1 and Th0 cultures are generated in parallel by using the following polarizing conditions: 1. Tr1 polarizing condition: rhIL-10 (10 ng/ml) and rhIFN-a (5 ng/ml). 2. Th0 polarizing condition: complete medium alone. T cells are split as necessary, and rhIL-2 and rhIL-15 together with polarizing cytokines are replenished in all cultures. At day 7 collect, wash, and count T cells, and restimulate them under identical conditions for an additional 7 days. After a total of 14 days of in vitro culture collect, wash, count, and analyze T cells for their profile of cytokine production and proliferative capacity. Note that cells under Tr1 polarizing conditions will expand poorly during the second stimulation, and thus, 5-10 more wells should be used for this condition. Timing for Tr1 cell lines differentiation: Day 0: Activate naïve CD4+ T cells with L cells in the presence of polarizing cytokines Day 7: Restimulate T cells under identical conditions Day 14: Tr1 cell lines are differentiated 3.2. Induction of Allo-Specific T Cells Using Monocyte and IL-10 3.2.1. Monocytes Preparation

Monocytes are isolated from PBMC as following: 1. Wash PBMC, isolated as previously described (see Subheading 3.1.1), but taking care to remove platelets by washing twice at 800 rpm for 10 min. 2. Resuspend PBMC in PBS supplemented with 2% FBS at the final concentration of 10 × 106 cells/ml (see Note 3). 3. Add the number of CD3-dynal-beads on the bases of the percentage of CD3+ T cells in the sample and according to the manufacturer’s instructions and incubate for 40 min at 4°C gently tilting. 4. Remove beads by two consecutive washing and count.

3.2.2. Induction of Allo-Specific IL-10-Anergized T Cells

IL-10-anergized T cells can be generated from pairs with different degree of HLA disparities. Total PBMC, total CD4+ T cells, or naïve CD4+ T cells can be used as responders, and allogeneic CD3-depleted cells as stimulators. 1. CD3-depleted cells, prepared as described before (see Subheading 3.2.1), are irradiated (6,000 rad). 2. Following washing, resuspend CD3-depleted cells in complete medium and count (see Note 4). 3. Cocultured irradiated CD3-depleted cells (1 × 106 cells/ml) with allogeneic responder cells (1 × 106 cells/ml) in 24-well plates in a final volume of 1 ml of complete medium.

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4. Add rhIL-10 (10 ng/ml) and culture for an additional 7 days (see Note 5). 5. At 7 day half of the medium, with or without rhIL-10, is replaced with fresh medium without cell suspension (see Note 6). 6. At day 10 collect and wash T cells two times with HBSS supplemented with 2% HS and resuspend in complete medium. Cells are ready to be tested. Note that under IL-10 culture cells will decrease in number (yield as low as 30% of the input), and thus, more wells should be used for this condition. Timing for allo-specific IL-10-anergized differentiation: Day 0: Plate responder cells and allogeneic CD3-depleted cells with or without rhIL-10 Day 7: Replace half of the medium with fresh medium containing or not rhIL-10 Day 10: IL-10-anergized cells are differentiated 3.3. Differentiation of Allo-Antigen-Specific Tr1 Cell Lines Using Tolerogenic DC-10 3.3.1. Tolerogenic DC-10 Preparation

1. Wash PBMC, isolated as previously described (see Subheading 3.1.1), but taking care to remove platelets (which interfere with monocyte adherence) by washing two times at 800 rpm for 10 min. A third wash may be necessary if a significant number of platelets remain. 2. To isolate CD14+ monocytes as the adherent fraction, plate 10 × 106 PBMC/well in a 6-well plate, in 2 ml of RPMI 1640 at 37°C. 3. Let remain at 37°C for 1–2 h (see Note 7). 4. After incubation, wash away the nonadherent cells by gently adding 2 ml of warm RPMI/well, and swirling. Do this at least four times, or until you can see under the microscope that all the nonadherent cells are eliminated. 5. Differentiate adherent monocytes into DC-10 by culturing them in 2 ml of DC medium containing 10 ng/ml rhIL-4, 100 ng/ml rhGM-CSF, and 10 ng/ml rhIL-10. 6. In parallel differentiate adherent monocytes into DC by culturing them in 2 ml of DC medium containing 10 ng/ml rhIL-4, and 100 ng/ml rhGM-CSF (see Note 8). 7. At day 3, add 2 ml of fresh DC medium containing IL-4 (20 ng/ml) and GM-CSF (200 ng/ml) with or without IL-10 (20 ng/ml). 8. After additional 2 days, half of the medium, with or without rhIL-10, is replaced with fresh one. 9. DC cultured in the presence of rhIL-4 and rhGM-CSF are matured by addition of LPS (1 mg/ml) (see Note 9). 10. After an additional 2 days, collect and irradiate (6,000 rad) DC-10 and mature DC, and use them to stimulate responder cells. In parallel, freeze mature DC for readouts (10).

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The purity and maturation state of DC are routinely checked by flow cytometric analysis to determine expression of CD1a, CD14, CD83, CD86, and HLA-DR. Timing of DC preparation: Day 0: Start DC differentiation Day 3: Add 2 ml of DC medium containing rhIL-4, rhGM-CSF with or without rhIL-10 Day 5: And rhIL-10 (20 ng/ml) for DC-10. In parallel mature DC are activated with 1 mg/ml LPS Day 7: DC-10 and mature DC are ready to use DC-10 are characterized by analyzing their: –

Phenotype

–

Cytokine-production profile

–

Stimulatory capacity

Phenotype: DC-10 are characterized by the expression of CD14 and CD11c. They do not express CD1a, and express CD83, CD86, CD80 at levels comparable to that observed in mature DC (Fig. 1). Mature DC

CD1a

DC-10

53%

76%

CD83

CD14 45%

48%

2.4%

18% HLA-DR

CD86

3.3.2. Biological Characterization of DC-10

95%

93%

CD11c

Fig. 1. DC-10 phenotype. Monocyte-derived DC were differentiated in IL-4 and GM-CSF in the presence of IL-10 (DC-10) for 7 days, or in IL-4 and GM-CSF for 5 days and cultured for additional 2 days with LPS (mature DC). Expression of CD1a, CD14, CD83, HLA-DR, CD11c, CD80, and CD86 was evaluated by FACS analysis. A representative donor out of twenty tested is presented.

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Cytokine production profile: DC-10 are characterized for their cytokine production. To this end: 1. Plate 2 × 105 DC-10 alone in the presence of IFN-g (50 ng/ ml) and LPS (200 ng/ml) in a 96-well plate in complete medium in a final volume of 200 ml and incubate at 37°C. 2. After 24 and 48 h collect culture supernatants to evaluate IL-12p70 (24 h) IL-10, IL-6, and TNF-a (40 h) by ELISA performed according to manufacturer’s instructions or Bioplex (see Note 10). Stimulatory capacity: DC-10 are tested for their ability to promote proliferation of allogeneic CD4+ T cells or total PBMC. 1. Co-cultured irradiated DC-10 (1 × 104) with CD4+ T or PBMC (1 × 105) in 96-well plate in complete medium at the final volume of 200 ml. 2. Pulse the cultures after 4 days, for 14–16 h with 1 mCi/well of (3H)thy (see Note 11). 3.3.3. Differentiation of Allo-Antigen-Specific Tr1 Cell Lines Using DC-10

Allo-Ag-specific Tr1 cell lines can be generated using DC-10 obtained from healthy, haploidentical, and HLA-matched unrelated donors. Total PBMC, total CD4+ T cells, or naïve CD4+ T cells are isolated as previously described (see Subheading 3.1.1) and are used as responder cells and allogeneic DC-10 as stimulators. 1. DC (1 × 105) are cocultured with allogeneic naïve CD4+ T cells (1 × 106) in 1 ml of complete medium. 2. Add rhIL-2 (20 ng/ml) after 6/7 days, in cocultures, and let expand T cells for an additional 7/8 days. 3. 14 days after initiation of the culture, collect and wash CD4+ T cells two times with HBSS supplemented with 2% HS. Resuspend cells in complete medium and test then for cytokine production profile, proliferative, and suppressive capacity (see below). T cells cultured with DC-10 typically expand threefold less in comparison to cultures stimulated with mature DC. Timing of T-cell differentiation: Day 0: Start cocultures (1 × 105 DC + 1 × 106 allogeneic T cells) Day 6/7: Add rhIL-2 Day 14: T-cell lines are ready to be tested

3.4. Enrichment of IL-10-Producing T Cells from Bulk Tr1 Cell Lines

Tr1 cell lines generated with L cells (see Subheading 3.1.2) or DC-10 (see Subheading 3.3.3) contained up to 15% of IL-10-producing T cells. In case an enriched population of IL-10-producing T cells is required for additional investigations, IL-10-producing T cells can be isolated using IL-10-secretion assay (Miltenyi), according to the manufacturer’s instructions.

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Briefly, Tr1 cell lines are activated with TPA (10 ng/ml) and immobilized anti-CD3 mAb (10 mg/ml) for 4 h and IL-10producing T cells are isolated using anti-CD45/anti-IL10 mAb followed by PE conjugated anti-IL-10 mAb. After incubation, IL-10-producing T cells are positively selected by PE conjugated beads using MS columns according to the manufacturer’s instructions (see Note 12). 3.5. Biological Characterization of Tr1 Cell Lines 3.5.1. Cytokine Production Profile of Tr1 Cell Lines

Tr1 cell lines polyclonally generated using with L cells (see Subheading 3.1.2) or induced in the presence of DC-10 (see Subheading 3.3.3) are characterized by their cytokine production pattern. Cytokines are measured in culture supernatants by ELISA. Alternatively, the frequency of IL-10-producing T cells can be determined by intracytoplasmic staining or ELISPOT. ELISA: To test cytokine production following TCR-mediated activation, Tr1 cells (1 × 106/ml) are activated with immobilized anti-CD3 mAb (10 mg/ml) and soluble anti-CD28 mAb (1 mg/ml) in a total volume of 200 ml/well. To determine cytokine production following allo-specific activation plate allo-specific Tr1 cell lines generated with DC-10 (5 × 105/ml) (see Subheading 3.3.3) with allogeneic monocytes (5 × 105/ml) or allogeneic mature DC (5 × 104/ml) from the same donor used in the priming, in a total volume of 200 ml/well. Collect supernatants after 24 h to determine the amounts of IL-2, and after 48 h to detect IL-4, IL-10, IFN-g, and TGF-b, and after 72 h to evaluate IFN-g and TGF-b by ELISA performed according to the manufacturer’s instructions. Intracytoplasmic staining: Tr1 cell lines (1.5 × 106/ml) generated with L cells (see Subheading 3.1.2) or induced in the presence of DC-10 (see Subheading 3.3.3) are stimulated with Leukocyte Activation Cocktail in complete medium for 5 h, according to the manufacturer’s instructions. After activation, CD4+ T cells are collected, washed in PBS 2% FBS, and stained with anti-CD4 mAbs (2 × 105/sample). After surface staining, cells are fixed and permeabilized with Fixation/Permeabilization Solution Kit, according to the manufacturer’s instructions. Permeabilized T cells are incubated with PE-coupled anti-hIL-2, or anti-hIL-10, and FITC-coupled anti-hIFN-g or anti-hIL-4 mAbs. The samples are analyzed using flow cytometer on CD4+ gated cells (Fig. 2). ELISPOT: IL-10-secreting T cells in cultures obtained with L cells (see Subheading 3.1.2), monocytes + IL-10 (see Subheading 3.2.2), or DC-10 (see Subheading 3.3.3) can be enumerated by enzyme-linked immunospot (ELISPOT) assay. 1. Coat ELISPOT plates with anti-IL-10 mAb, incubate at 4°C. 2. After incubation, wash plates and add 200 ml/well of PBS supplemented with 5% FBS, incubate for 1 h at RT, and after incubation wash them.

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Fig. 2. Cytokine production profile of Tr1 cell lines. Naive CD4+ T cells from peripheral blood of normal donor are activated by anti-CD3 mAbs cross-linked on L cells in the presence of IL-10 and IFN-a (Tr1), or in the absence of IL-10 (Th0). After two rounds of identical stimulation, IL-2, IFN-g, IL-4, and IL-10 expression is determined by intracytoplasmic staining upon stimulation with Leukocyte Activation Kit for 5 h. One representative donor is shown. Numbers represent percentage of positive cells.

3. Plate 1 × 105 IL-10-anergized cells in anti-IL-10-coated ELISPOT plates, add TPA (10 ng/ml) and Ionomycin (150 ng/ml) in a final volume of 200 ml, and incubate for 48 h at 37°C. 4. After culture, wash plates and add anti-IL-10 detection mAb and incubate for 2 h at RT. 5. After incubation, wash plates and reveal spots with avidinPOD, and count spots. 3.5.2. Allo-Specific T Cell Anergy

Once IL-10-anergized T cells are generated with either allogeneic monocytes + IL-10 (see Subheading 3.2.2) or allogeneic DC-10 (see Subheading 3.3.3) are tested for their proliferative response specifically to the allo-antigen used in the priming (allogeneic monocytes or mature DC). 1. Plate 1 × 105 IL-10-anergized T cells with 1 × 105 irradiated (6,000 rad) CD3-depleted cells in round-bottom 96-well plates or 1 × 104 irradiated (6,000 rad) mature DC. 2. After 2 days of culture, collect 50 ml of supernatants to measure IFN-g (see below) and add 50 ml of fresh medium. 3. Pulse the cultures after 2 days, for 14–16 h with 1 mCi/well of (3H)thy (see Note 13).

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4. IFN-g in culture supernatants is detected using an ELISA test performed according to the manufacturer’s instructions. 5. To calculate the percentage of anergy, use the following: (MLR cpm-MLR/10 cpm)/MLR × 100 or (MLR/mDC − MLR/DC-10)/MLR/mDC × 100. 3.5.3. Proliferative Response to Nominal Antigen

Once IL-10-anergized T cells are generated with either allogeneic monocytes + IL-10 (see Subheading 3.2.2) or allogeneic DC-10 (see Subheading 3.3.3), they are tested for their ability to proliferate in response to nominal antigens (Tetanus Toxoid) and to polyclonal stimulation (TPA/ionomycin). 1. For polyclonal proliferation, plate 1 × 105 IL-10-anergized T cells with TPA (10 ng/ml) and ionomycin (150 ng/ml; Sigma) (see Note 13). 2. After 2 days of culture, collect 50 ml of supernatants to measure IFN-g (see Note 10) and add 50 ml of fresh medium. 3. Pulse the cultures after 2 days, for 14–16 h with 1 mCi/well of (3H)thy (see Note 11). 4. IFN-g in culture supernatants is detected using an ELISA test performed according to the manufacturer’s instructions. 5. For proliferation in response to nominal antigens, plate 1 × 105 IL-10-anergized T cells with 1 × 105 irradiated (6,000 rad) autologous CD3-depleted cells in 96-well plates in the presence of Tetanus Toxoid at 5 mg/ml (see Note 13). 6. Pulse the cultures after 4 days, for 14–16 h with 1 mCi/well of (3H)thy (see Note 11).

3.5.4. Suppressive Function

To test the suppressive capacity of Tr1 cell lines generated with L cells (see Subheading 3.1.2) by flow cytometry, allogeneic or autologous PBMC are labeled with 5-(and-6)-CFSE before the stimulation with immobilized anti-CD3 (10 mg/ml) and soluble anti-CD28 (1 mg/ml) mAbs and cocultured with Tr1 cell lines at 5:2 ratio. After 4 days of culture, proliferation of CFSE-labeled PBMC is determined by flow cytometry, gating the responder cells for CD4+ cells (Fig. 3). Alternatively, autologous CD4+ T cells are labeled CFSE and stimulated with allogeneic irradiated CD3-depleted cells (see Subheading 3.2.1) at 1:1 ratio and immobilized anti-CD3 (1 mg/ml) in the presence of Tr1 cell lines. After 4 days of culture, proliferation of CFSE-labeled CD4+ cells is determined by flow cytometry. To test for the capacity of Tr1 cells generated with DC-10 to suppress proliferation and/or cytokine production, culture autologous CD4+ T cells (5 × 104 cells/well), which are cryopreserved at the start of the experiment, with allogeneic irradiated (6,000 rad) mature DC (10:1, T:DC). Stimulate naïve CD4+ T cells alone,

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Responder CD4+ T cells + Responder CD4+ T cells

Th0

Tr1

Fig. 3. Tr1 cell lines suppress proliferative response of autologous CD4+ T cells. Purified autologous CD4+ T cells stained with CSFE were stimulated with allogeneic APC with anti-CD3 (1 mg/ml) alone or in the presence of T-cell lines at 1:1 ratio. Cytofluorometric analysis of CSFE dilution was performed after 4 days of co-culture. Dot plots and histograms of CSFE-positive population are shown. The percentages of undivided cells are reported in the dot plots. In the histograms, the overlay between the responder cells cocultured with the T-cell lines (black line) and the responder cell alone (gray solid line) is shown. The percentages reported in the histograms represent suppression of responder proliferation.

T(DC-10) + MLR T(mDC) + MLR MLR

cpm x 10−3

300 250 200 150 100 50 0 2

3 days of culture

4

Fig. 4. DC-10 induce Tr1 cell with suppressive activity. Naïve CD4+ T cells were stimulated with allogeneic DC-10 (T(DC-10)), or mDC (T(mDC)) for 14 days. After stimulation, T-cell lines were tested for their ability to suppress responses of autologous CD4+ T cells activated with mDC (MLR). Naïve CD4+ T cells were stimulated with mDC alone (MLR) or in the presence of T(DC-10), and T(mDC) cell lines at a 1:1 ratio. (3H)-thymidine was added after 3 days of culture for an additional 16 h. Results of one experiment representative of eight independent experiments are shown.

or in the presence of IL-10-anergized Tr1 cells (1:1 ratio), in a final volume of 200 ml of complete medium in 96-well roundbottom plates (see Note 11). After 2, 3, or 4 days of culture, pulse the cultures for 14–16 h with 1 mCi/well of (3H)thy (see Note 13) (Fig. 4). Alternatively, collect supernatants for

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analysis of inhibition of IFN-g production. To calculate percentage of suppression, use the following: MLR cpm − (MLR + T(DC-10) cpm)/MLR × 100.

4. Notes 1. To improve the purity of CD4+ T cells, do not load more than 2 × 108 cells per column. 2. To purify naïve CD4+ T cell from peripheral blood, use LD columns (Miltenyi) for negative selection. 3. Do not put more than 50 × 106 PBMC in one vial; in case more cells need to be depleted, use additional vials. 4. Keep CD3-depleted cells on ice. 5. PBMC primed with allo-monocytes in the absence of hrIL-10 are routinely generated and used in parallel to IL-10anergized cells, as control MLR. 6. If necessary, split cultures generated in the absence of rhIL-10 (control MLR). 7. Do not let the PBMC adhere for more than 2 h as the monocytes will start to detach from the plastic. 8. Mature DC are routinely used to generate, in parallel to IL-10-anergized cells, control effector T cells. 9. Only DC differentiated in the presence of rhIL-4 and rhGMCSF are activated with LPS; DC-10 are left inactivated. 10. Bioplex analysis allows the detection of more cytokines in 50 ml of supernatant; much larger volumes are required for ELISA. Therefore, when cell numbers are limiting, Bioplex is usually preferable. 11. Cells are harvested and counted in a scintillation counter. 12. Cell recovery is very low. 13. Prepare triplicate wells for each condition.

Acknowledgments The authors thank Claudia Sartirana and Chiara Francesca Magnani for technical help. This work was supported by a grant from the Italian Telethon Foundation, Cariplo Foundation, the Italian Association for Cancer Research (Associazione Italiana per la Ricerca sul Cancro, AIRC), and the EU graut to MGR, Riset Consortion (www.risetFp6.org)

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References 1. Gavin, M.A., J.P. Rasmussen, J.D. Fontenot, V. Vasta, V.C. Manganiello, J.A. Beavo, and A.Y. Rudensky. 2007. Foxp3-dependent programme of regulatory T-cell differentiation. Nature 445:771–775. 2. Bacchetta, R., E. Gambineri, and M.G. Roncarolo. 2007. Role of regulatory T cells and FOXP3 in human diseases. J Allergy Clin Immunol 120:227–235; quiz 236–227. 3. Vignali, D.A., L.W. Collison, and C.J. Workman. 2008. How regulatory T cells work. Nat Rev Immunol 8:523–532. 4. Roncarolo, M.G., S. Gregori, M. Battaglia, R. Bacchetta, K. Fleischhauer, and M.K. Levings. 2006. Interleukin-10-secreting type 1 regulatory T cells in rodents and humans. Immunol Rev 212:28–50. 5. Bacchetta, R., M. Bigler, J.L. Touraine, R. Parkman, P.A. Tovo, J. Abrams, R. de Waal Malefyt, J.E. de Vries, and M.G. Roncarolo. 1994. High levels of interleukin 10 production in vivo are associated with tolerance in SCID patients transplanted with HLA mismatched hematopoietic stem cells. J Exp Med 179: 493–502. 6. Groux, H., A. O’Garra, M. Bigler, M. Rouleau, S. Antonenko, J.E. de Vries, and M.G. Roncarolo. 1997. A CD4+ T-cell subset inhibits antigen-specific T-cell responses and prevents colitis. Nature 389:737–742. 7. Bacchetta, R., C. Sartirana, M.K. Levings, C. Bordignon, S. Narula, and M.G. Roncarolo. 2002. Growth and expansion of human T regulatory type 1 cells are independent from TCR activation but require exogenous cytokines. Eur J Immunol 32:2237–2245. 8. Vieira, P.L., J.R. Christensen, S. Minaee, E.J. O’Neill, F.J. Barrat, A. Boonstra, T. Barthlott, B. Stockinger, D.C. Wraith, and A. O’Garra. 2004. IL-10-secreting regulatory T cells do not express Foxp3 but have comparable regulatory function to naturally occurring CD4+CD25+ regulatory T cells. J Immunol 172:5986–5993. 9. Allan, S.E., L. Passerini, R. Bacchetta, N. Crellin, M. Dai, P.C. Orban, S.F. Ziegler, M.G. Roncarolo, and M.K. Levings. 2005. The role of 2 FOXP3 isoforms in the generation of human CD4 Tregs. J Clin Invest 115:3276–3284. 10. Levings, M.K., S. Gregori, E. Tresoldi, S. Cazzaniga, C. Bonini, and M.G. Roncarolo. 2005. Differentiation of Tr1 cells by immature dendritic cells requires IL-10 but not CD25+ CD4+ Tr cells. Blood 105:1162–1169.

11. Cobbold, S.P., E. Adams, L. Graca, and H. Waldmann. 2003. Serial analysis of gene expression provides new insights into regulatory T cells. Semin Immunol 15:209–214. 12. Pot, C., H. Jin, A. Awasthi, S.M. Liu, C.Y. Lai, R. Madan, A.H. Sharpe, C.L. Karp, S.C. Miaw, I.C. Ho, and V.K. Kuchroo. 2009. Cutting edge: IL-27 induces the transcription factor c-Maf, cytokine IL-21, and the costimulatory receptor ICOS that coordinately act together to promote differentiation of IL-10-producing Tr1 cells. J Immunol 183: 797–801. 13. Haringer, B., L. Lozza, B. Steckel, and J. Geginat. 2009. Identification and characterization of IL-10/IFN-gamma-producing effectorlike T cells with regulatory function in human blood. J Exp Med 206:1009–1017. 14. Carpentier, A., F. Conti, F. Stenard, L. Aoudjehane, C. Miroux, P. Podevin, O. Morales, S. Chouzenoux, O. Scatton, H. Groux, C. Auriault, Y. Calmus, V. Pancre, and N. Delhem. 2009. Increased expression of regulatory Tr1 cells in recurrent hepatitis C after liver transplantation. Am J Transplant 9:2102–2112. 15. Grossman, W.J., J.W. Verbsky, B.L. Tollefsen, C. Kemper, J.P. Atkinson, and T.J. Ley. 2004. Differential expression of granzymes A and B in human cytotoxic lymphocyte subsets and T regulatory cells. Blood 104:2840–2848. 16. Levings, M.K., R. Sangregorio, F. Galbiati, S. Squadrone, R. de Waal Malefyt, and M.G. Roncarolo. 2001. IFN-alpha and IL-10 induce the differentiation of human type 1 T regulatory cells. J Immunol 166:5530–5539. 17. Barrat, F.J., D.J. Cua, A. Boonstra, D.F. Richards, C. Crain, H.F. Savelkoul, R. de WaalMalefyt, R.L. Coffman, C.M. Hawrylowicz, and A. O’Garra. 2002. In vitro generation of interleukin 10-producing regulatory CD4(+) T cells is induced by immunosuppressive drugs and inhibited by T helper type 1 (Th1)- and Th2-inducing cytokines. J Exp Med 195: 603–616. 18. Kemper, C., A.C. Chan, J.M. Green, K.A. Brett, K.M. Murphy, and J.P. Atkinson. 2003. Activation of human CD4+ cells with CD3 and CD46 induces a T-regulatory cell 1 phenotype. Nature 421:388–392. 19. Murugaiyan, G., A. Mittal, R. Lopez-Diego, L.M. Maier, D.E. Anderson, and H.L. Weiner. 2009. IL-27 is a key regulator of IL-10 and IL-17 production by human CD4+ T cells. J Immunol 183:2435–2443.

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20. Paccani V., Gregori S, Chini L, Corrente S, Chianca M, Moschese V, Rossi P, Roncarolo MG, Angelini F. 2010. Induction of anergic allergen-specific suppressor T cells using tolerogenic dendritic cells derived from children with allergies to house dust mites. J Allergy Clin Immunol 125:727–36. 21. Gregori, S., R. Bacchetta, L. Passerini, M.K. Levings, and M.G. Roncarolo. 2007. Isolation, expansion, and characterization of human natural and adaptive regulatory T cells. Methods Mol Biol 380:83–105. 22. Bacchetta R., Gregori S, Serafini G, Sartirana C, Schultz U, Zino E, Tomiuk S, Janben U, Ponzoni M, Paties CT, Fleischhauer K, Roncarolo MG. 2010. Molecular and functional characterization of alloantigen-specific anergic T cells suitable for cell therapy. Haematologica, in press. Epub ahead of print August 2010. 23. Morelli, A.E. and A.W. Thomson. 2007. Tolerogenic dendritic cells and the quest for transplant tolerance. Nat Rev Immunol 7: 610–621. 24. Levings, M.K. and M.G. Roncarolo. 2005. Phenotypic and functional differences between human CD4+CD25+ and type 1 regulatory T cells. Curr Top Microbiol Immunol 293: 303–326. 25. Jonuleit, H., E. Schmitt, M. Stassen, A. Tuettenberg, J. Knop, and A.H. Enk. 2001. Identification and functional characterization of human CD4(+)CD25(+) T cells with regulatory properties isolated from peripheral blood. J Exp Med 193:1285–1294. 26. Gregori S., Tomasoni D, Pacciani V, Scirpoli M, Battaglia M, Magnani Cf, Hauben E, Roncarolo MG. 2010 May 6. [Epub ahead of

27.

28.

29.

30.

31.

32.

print]. Differentation of type 1 T regulatory (Tr1) cells by tolerogenic DC-10 requires the IL-10-dependent ILT4/HLA-G pathway. Blood. Foussat, A., F. Cottrez, V. Brun, N. Fournier, J.P. Breittmayer, and H. Groux. 2003. A comparative study between T regulatory type 1 and CD4+CD25+ T cells in the control of inflammation. J Immunol 171:5018–5026. Cong, Y., C.T. Weaver, A. Lazenby, and C.O. Elson. 2002. Bacterial-reactive T regulatory cells inhibit pathogenic immune responses to the enteric flora. J Immunol 169:6112–6119. Zeller, J.C., A. Panoskaltsis-Mortari, W.J. Murphy, F.W. Ruscetti, S. Narula, M.G. Roncarolo, and B.R. Blazar. 1999. Induction of CD4+ T cell alloantigen-specific hyporesponsiveness by IL-10 and TGF-beta. J Immunol 163:3684–3691. Battaglia, M., A. Stabilini, E. Draghici, B. Migliavacca, S. Gregori, E. Bonifacio, and M.G. Roncarolo. 2006. Induction of tolerance in type 1 diabetes via both CD4+CD25+ T regulatory cells and T regulatory type 1 cells. Diabetes 55:1571–1580. Gagliani N., Jofra T, Stabilini A, Valle A, Atkinson M, Roncarolo MG, Battaglia M. 2010. Antigen-specific dependence of Tr1-cell therapy in preclinical models of islet transplant. Diabetes. 59(2):433–9. Brun, V., H. Bastian, V. Neveu, and A. Foussat. 2009. Clinical grade production of IL-10 producing regulatory Tr1 lymphocytes for cell therapy of chronic inflammatory diseases. Int Immunopharmacol 9: 609–613.

Chapter 4 Ex Vivo Generation of Regulatory T Cells: Characterization and Therapeutic Evaluation in a Model of Chronic Colitis Fridrik Karlsson, Sherry A. Robinson-Jackson, Laura Gray, Songlin Zhang, and Matthew B. Grisham Abstract Naturally occurring regulatory T cells (nTregs; CD4+CD25+Foxp3+) are capable of suppressing the chronic inflammation observed in a variety of different animal models of autoimmune and chronic inflammatory diseases such as inflammatory bowel diseases, diabetes, and arthritis. A major limitation in exploring how and where nTregs exert their suppression in vivo is the relative paucity of these regulatory cells. Although several laboratories have described different methods to expand flow-purified nTregs or convert conventional/naïve T cells (CD4+Foxp3−) to Foxp3-expressing “induced” Tregs (iTregs; CD4+Foxp3+) ex vivo, we have found that many of these approaches are encumbered with their own limitations. Therefore, we sought to develop a relatively simple ex vivo method to generate large numbers of Foxp3expressing iTregs that can be used to evaluate their trafficking properties, suppressive activity, and therapeutic efficacy in a mouse model of chronic gut inflammation in vivo. We present a detailed protocol demonstrating that polyclonal activation of conventional CD4+ T cells in the presence of IL-2, TGFb, and all trans retinoic acid induces >90% conversion of these T cells to Foxp3-expressing iTregs as well as promotes a three- to fourfold increase in proliferation following a 4-day incubation period in vitro. This protocol enhances modestly the surface expression of the gut-homing adhesion molecule CCR9 but not a4b7. Furthermore, we provide preliminary data demonstrating that these iTregs are significantly more potent at suppressing T-cell activation in vitro and are equally effective as freshly isolated nTregs at attenuating chronic colitis in vivo. Finally, we report that this protocol has the potential to generate 30–40 million iTregs from one healthy mouse spleen. Key words: iTregs, TGFb, Retinoic acid, Colitis, Inflammatory bowel disease

1. Introduction Naturally occurring regulatory T cells (nTregs; CD4+CD25+Foxp3+) are known to suppress a wide range of immune responses via several different mechanisms including production of regulatory cytokines, Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_4, © Springer Science+Business Media, LLC 2011

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competition for essential cytokines, and contact-dependent mechanisms (1–4). Because nTregs have been shown to suppress Th1 and Th17 autoimmune responses in vivo, a great deal of interest has been generated regarding the possible use of these cells to treat patients with chronic inflammatory disorders such as the inflammatory bowel diseases (IBD; Crohn’s disease, ulcerative colitis), diabetes, arthritis, and graft vs. host disease. A major limitation in defining how and where nTregs exert their suppression in vivo is the relative paucity of these regulatory cells as they constitute 90% conversion of these T cells to Foxp3-expressing iTregs as well as promotes a 3-4-fold increase in proliferation following a 4-day incubation period in vitro. In addition, this protocol enhances modestly the surface expression of the gut-homing adhesion molecule CCR9 but not a4b7. Furthermore, we provide preliminary data demonstrating that these iTregs are significantly more potent at suppressing T-cell activation in vitro and are equally effective as nTregs at attenuating chronic colitis in vivo. Finally, we report that this protocol has the potential to generate 30–40 million iTregs from one healthy mouse spleen.

2. Materials 2.1. Animals

C57Bl/6 wild-type (WT) and recombinase activating gene-1deficient (RAG-1−/−) mice were obtained from the Jackson Laboratory (Bar Harbor, Maine), whereas Foxp3GFP “knockin” mice were obtained from the LSUHSC breeding facility (originally obtained from Dr. Alexander Rudensky, University of Washington).

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All mice were housed under specific pathogen-free conditions in the LSUHSC-Shreveport animal care facility. 2.2. Tissue Culture Plastic Ware and Reagents

1. Costar® 24-Well Clear TC-Treated Microplates, Sterile (Corning). 2. Mouse CD3e-monoclonal antibody (mAb) (eBioscience). 3. Phosphate-buffered saline (PBS) pH 7.4.

2.3. Splenocyte and CD4 + T-Cell Preparation

1. PBS with 4% fetal bovine serum (PBS/FBS). 2. Red blood cells lysis buffer (RBC-LB): 0.14 M NH4Cl and 0.0165 M Tris base in water with pH adjusted to 7.1–7.2. 3. Trypan Blue solution: 0.4% solution. 4. Dynal® Mouse CD4 Cell Negative Isolation Kit (Invitrogen). 5. Dynal buffer: 1× PBS with 0.1% bovine serum albumin (BSA) and 2 mM EDTA. 6. Fetal bovine serum.

2.4. T-Cell Conversion

1. RPMI-10 Complete medium: RPMI-1640 (Sigma) supplemented with L-glutamine, antibiotic/antimycotic solution, 50 mM b-mercaptoethanol, and 10% FBS. 2. Recombinant Human TGF-b1 (2 mg; R&D Systems) is first dissolved in 40 ml of a 4 mM HCl solution to which 360 ml of 0.1% BSA in PBS is then added to yield a 5 mg/ml stock solution. Aliquots of this solution are stored at −80°C (see Note 1). 3. Recombinant Human IL-2 (Chiron) is dissolved in distilled water to given a final concentration of 18 × 106 U/ml. This stock solution is kept at 4°C. 4. A 10 mM stock solution of all trans Retinoic acid (Acros) is made by dissolving in dimethyl sulfoxide (DMSO) and stored in small aliquots at −80°C. Subsequent dilutions can be made using RPMI-10 as described later.

2.5. Flow-Cytometric Analysis

Allophycocyanin (APC)-conjugated CD4 antibody (clone GK1.5), Phycoerythrin (PE)-conjugated Foxp3 antibody (clone FJK-16s), and Foxp3 staining buffer set (all from eBioscience).

3. Methods The overall objective of the protocol described below is to generate large numbers of iTregs from a mouse splenocyte preparation in a relatively short period of time using common immunological methods and laboratory instrumentation. If performed carefully,

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this method does not require the use of fluorescence-activated cell sorting (FACS) and can be accomplished using a variety of different genetically engineered mutant mouse T cells. Indeed, we describe the use of genetically engineered Foxp3GFP knockin mice (14) in which expression of the green fluorescent protein (GFP) is driven by the Foxp3 promoter, thereby providing a “nonlethal” method to quantify (by flow cytometry) Foxp3 expression within T cells without permeabilizing/killing the lymphocytes. We compare conversion of these Foxp3GFP T cells with that obtained using T cells obtained from WT mice or mice deficient in specific selectins and/or integrins. Finally, we present a detailed protocol for assessing the suppressive activity of the iTregs in vitro and in a model of chronic gut inflammation in vivo. 3.1. Preparation of CD3 mAb-Coated, 24-Well Tissue Culture Plates

1. A 10 mg/ml solution of CD3 mAb is prepared by adding 120 ml of the stock CD3-antibody (1 mg/ml) into 12 ml of cold sterile PBS, which is then mixed by vortex (see Note 2). 2. 500 ml aliquots of the 10 mg/ml CD3 mAb solution is pipetted into each well of the 24 wells. 3. The plate is sealed with Parafilm to prevent evaporation and stored overnight in refrigerator at 4°C (The CD3 mAb solution will be removed in step 4, Subheading 3.4).

3.2. Preparation of a Single Cell Suspension from Mouse Spleens

This section describes the preparation of splenocytes from either Foxp3GFP or WT spleens and is applicable for most mouse strains and phenotypes. All steps outlined in this section should be performed under sterile conditions using sterile reagents and plasticware. 1. Each spleen is aseptically dissected from the mouse (using sterile instruments) and placed in a small Petri dish containing 10 ml of ice-cold PBS/FBS (see Note 3). 2. Using the frosted sides of two microscope slides, press against the spleen and grind it into a homogenous pulp in the Petri dish. 3. Using a 10-ml syringe without the needle, aspirate the cell suspension, attach a 26-G needle, and expel the contents of the syringe through a 70-mm cell strainer and into a 15-ml sterile conical centrifuge tube. 4. Rinse the Petri dish with an additional 5 ml of PBS/FBS, aspirate the cell suspension into the 10-ml syringe, and repeat step 3 combining the contents into the same sterile 15-ml conical centrifuge tube from above. 5. Centrifuge the cells for 10 min at 400 × g in a cooled centrifuge (4°–8°C). Aspirate and discard most of the supernatant leaving approximately 500 ml of residual fluid to resuspend

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the cell pellet. Using a 1-ml micropipettor, gently resuspend the pellet in the residual fluid and measure this volume. 6. Remove 20 ml of the cell suspension and add it to a 500-ml microcentrifuge tube containing 60 ml of RBC-LB. Allow the cells to incubate at room temperature in the RBC-LB for 3 min to insure complete lysis of erythrocytes. 7. After the 3-min incubation in RBC-LB, remove 10 ml of the cell solution and add it to new microcentrifuge tube containing 90 ml of 0.4% trypan blue. Gently finger tap the tube to mix the cells (do not vortex) and remove 10 ml to count on a hemocytometer. 3.3. CD4 + T-Cell Enrichment Using Negative Selection

This section describes the steps required to enrich the splenocyte preparation for CD4+ T cells. Although the protocol described here was performed using the Dynal® Mouse CD4 Cell Negative Isolation Kit (Invitrogen), any negative selection protocol for enriching CD4+ T cells from a splenocyte preparation may be used. 1. Splenocyte concentration is adjusted to 1 × 107 cells per 100 ml using Dynal buffer (see Note 4). 2. For every 100 ml of cell suspension, 20 ml of FBS and 20 ml of antibody mix from the Invitrogen kit are added to the tube. The cell suspension is incubated at 4°C for 20 min on a gently rocking platform. 3. Following the incubation period, the tubes are filled with Dynal buffer, gently mixed by inverting the tubes and centrifuged at 400 × g for 10 min at 4°C. The supernatant is aspirated and discarded. 4. For every 1 × 107 cells present in the cell pellet, 800 ml of Dynal buffer and 200 ml of prewashed Dynabeads are added, and the cells are resuspended (instructions for washing beads can be found in the kit instructions) (see Note 5). The cell and bead suspension is then incubated at room temperature with gentle rocking for 15 min. 5. Following this 15-min incubation period, the cell and bead mixture is gently resuspended five times using a 1-ml micropipettor and an additional 1 ml of Dynal buffer for every 1 × 107 cells is added to the tube. The tube is then placed against the magnet and separation is allowed to proceed for 3 min at room temperature according the companies protocol. This step removes B-cells, CD8+ T cells, macrophages, PMNs, erythrocytes, monocytes, and dendritic cells by promoting the magnetic binding of these cells to wall of the tube leaving an enriched population of CD4+ T cells in the fluid phase.

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6. The fluid from the tube is removed being careful not to disturb the beads bound to the sides of the tube by the magnet and then transferred into a new sterile 15-ml plastic conical tube. The tube is filled with Dynal buffer and centrifuged at 400 × g for 10 min at 4°C. The supernatant is aspirated and discarded. The pellet is resuspended in PBS/FCS. This fraction contains a highly enriched population of CD4+ T cells that will be used for Treg conversion. A small aliquot of these cells should be removed and analyzed for CD4-purity using flow cytometry. We routinely achieve CD4+ T-cell enrichment of 85–92% using the Dynal negative selection kit. 3.4. Conversion of CD4+ Foxp3 − T Cells to iTregs

The protocol outlined in this section describes the methods required to convert 12 million negatively selected CD4+ T cells to iTregs in one 24-well tissue culture plate. We have found that negatively selected CD4+ T cells are routinely >85–92% conventional T cells (CD4+Foxp3− T cells) and will generate >90% CD4+Foxp3+ iTregs using the conversion protocol described below. If greater purity is required for, FACS may be used to generate iTreg purity of >98%. 1. In order to minimize well-to-well variability during the plating of large numbers of cells into multiple wells in a 24-well plate, we first prepare a 24-ml cell suspension in RPMI-10 containing 12 million CD4+ T cells, 135 U/ml of human recombinant IL-2, 20 ng/ml TGFb, and 1 nM (1 pmol/ml) all trans retinoic acid (RA). 2. To do this, we add small aliquots (90% of the viable CD4+ T cells express Foxp3 by day 4 (Fig. 3). Indeed, we observe an initial decrease in cell number lasting from 24–48 h. This decrease is then followed by a time-dependent increase in T-cell proliferation such that maximal proliferation is observed at day 4 reaching a three- to fourfold expansion from what was originally plated (Fig. 3).This conversion protocol produces very similar results using negatively selected CD4+ T cells obtained from Fopx3GFP or WT mice (Fig. 3) (see Note 6).

3.6.3. In Vitro Suppression Assay

To compare the suppressive activity of iTregs with freshly isolated nTregs, we performed a standard in vitro suppression assay with minor modifications (19). Briefly, freshly isolated, negatively selected CD4+ T cells from Foxp3GFP were sorted for nTregs (CD4+GFP+ T cells) and CD4+GFP− (responder) T cells. Varying numbers of CD4+GFP+ T cells (nTregs) or iTregs (from Foxp3GFP mice) were cultured with 5 × 104 responder CD4+GFP− T cells, 105 irradiated antigen-presenting cells, and 1 mg/ml CD3 Ab.

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Fig. 3. Time-dependent induction and expansion of Foxp3-expressing iTregs ex vivo. Foxp3 expression and proliferation were quantified at different times in vitro for CD4+T cells obtained from Foxp3GFP “knockin” mice (a) or WT mice (b) activated with platebound CD3 mAb in the presence of IL-2, TGFb, and RA. Proliferation was expressed as fold change from the original cell number.

Antigen-presenting cells were prepared by irradiating (2,000 rad) splenocytes. The cells were cultured for 96 h, with the addition of 1 mCi/well of tritiated (3H)-thymidine for the final 24 h. These same suppression assays were also performed with flow-purified WT nTregs (CD4+CD25+ T cells) or iTregs in the presence of WT responder (CD4+CD25-) T cells, irradiated WT splenocytes, and CD3 Ab. Figure 4 demonstrates that iTregs generated from Foxp3GFP or WT T cells were significantly more suppressive than freshly nTregs. 3.6.4. Foxp3 Expression in CD4+ T Cells Deficient in L-Selectin, b7 Integrin, or Both

Because our laboratory is interested in T-cell trafficking, we wished to determine whether our conversion protocol could be used to convert CD4+ T cells obtained from mice deficient in different selectins and/or integrins. Therefore, we enriched for CD4+ T cells by negative selection of splenocytes obtained from

Conversion of Conventional T-Cell to Tregs

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Fig. 4. Suppressive activity of freshly isolated nTregs or ex vivo-generated iTregs. (a) Freshly isolated, negatively selected CD4+ T cells from Foxp3GFP mice were sorted for nTregs (CD4+GFP+ T cells) and naïve (CD4+GFP−) T cells. Varying numbers of CD4+GFP+ T cells (nTregs) or ex vivo-generated iTregs (from Foxp3GFP mice) were cultured with 5 × 104 naïve (responder) CD4+GFP− T cells, 105 irradiated splenocytes (from Foxp3GFP mice), and 1 mg/ml CD3 mAb. The cells were incubated for 96 h at 37°C with 1 mCi/well of 3H-thymidine added during the final 24 h. (b) This same suppression assay was performed with flow-purified WT nTregs (CD4+CD25+ T cells) or ex vivo-generated iTregs in the presence of WT responder cells (CD4+CD25− T cells), irradiated WT splenocytes and CD3 mAb. Data represent the mean ± SEM for triplicate samples performed at least two different times. Black bars represent T-cell proliferation in the absence of Tregs; shaded bars represent T-cell proliferation in the presence of nTregs; and white bars represent T-cell proliferation in the presence of iTregs. *P < 0.05 compared to proliferation in the absence of Tregs.

C576Bl/6 mice that were deficient in L-selectin (CD62L−/−), b7 integrin (b7−/−), or both (CD62L−/− × b7−/−) and subjected these cells to our conversion protocol. We found, using our standard 4-day incubation period, that all three mutant T-cell populations converted to the iTreg phenotype were quantitatively similar to WT CD4+ T cells (Fig. 5). These data demonstrate the utility of this protocol for producing iTregs from several different mutant CD4+ T-cell populations.

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b7−/−

CD62L−/−฀x b7−/−

Fig. 5. Foxp3 expression in CD4+ T cells obtained from mice deficient in L-Selectin, b7 integrin, or both. Negatively selected CD4+ T cells obtained from mice deficient in L-Selectin (CD62L−/−), b7 integrin (b7−/−), or both (CD62L−/− × b7−/−) were plate into CD3 mAb-coated wells containing IL-2, TGFb, and RA for 4 days at 37°C. Cells were permeabilized and stained for CD4 and Foxp3. Cells were gated on CD4+ T cells and Foxp3 expression was determined by flow cytometry. The shaded curves represent isotype control Ab staining.

3.6.5. Suppressive Activity of iTregs in a Mouse Model of Chronic Gut Inflammation

Having established the feasibility of producing large numbers of iTregs with potent suppressive activity in vitro, we next wished to assess the therapeutic efficacy of these ex vivo-generated iTregs in suppressing the development of chronic colitis in mice in vivo. To do this, we utilized the well-characterized T-cell transfer model of chronic colonic inflammation in mice (20). Although a detailed description of this model is beyond the scope of the current chapter, we refer the reader to the recently published protocol for inducing chronic colitis using this model (20). Briefly, chronic colitis is induced in lymphopenic recombinase activating gene-1 deficient mice (RAG−/−) by adoptive transfer of 0.5 × 106 WT CD4+CD45RBhigh(naïve) T cells. Chronic colitis develops within 6–8 weeks post T-cell transfer. To compare the suppressive activity of cultured iTregs with freshly isolated nTregs (CD4+CD25+ T cells) in this model of inflammatory bowel disease, we injected (i.p.) 1 × 105 iTregs or nTregs into RAG−/− recipients 10 days following the adoptive transfer of the disease-producing CD4+CD45RBhigh T cells. At 8-week posttransfer of naïve T cells, mice were euthanized, their colons removed and cleansed of fecal material, and fixed in PBS formalin. Colons were then embedded in paraffin, sectioned, and stained with H&E and scored by a pathologist unfamiliar with the treatment groups. Figure 6 demonstrates that iTregs are as effective as freshly isolated nTregs in suppressing the development of chronic colitis in this model of IBD.

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Fig. 6. Suppressive activity of freshly isolated nTregs or ex vivo-generated iTregs in a mouse model of chronic colitis. Chronic colitis was induced in recombinase activating gene-1 deficient (RAG−/−) mice by adoptive transfer of 5 × 105 WT CD4+CD45RBhigh(naïve) T cells. Freshly isolated nTregs (1 × 105 cells; CD4+CD25+ T cells) or ex vivo-generated iTregs (1 × 105 cells) from WT mice were injected (i.p.) into RAG−/− recipients 10 days following the adoptive transfer of the disease-producing CD4+CD45RBhigh T cells. At 8-week posttransfer of naïve T cells, mice were euthanized and their colons removed for blinded histopathological evaluation.

4. Conclusions 1. We describe a detailed protocol for generating large numbers of iTregs from a mouse splenocyte preparation in a relatively short period of time using common immunological methods and laboratory instrumentation. This method does not require cell sorting for most studies and is therefore less expensive and time-consuming than some protocols described elsewhere. Indeed, we have found no obvious advantage of using flow-purified CD4+GFP− or CD4+CD25− T cells for conversion to iTregs using our protocol (data not shown). 2. Conversion of negatively selected CD4+ T cells to iTregs can be performed with cells obtained from different populations of C57Bl/6 mice including WT and Foxp3GFP mice as well as mice with genetic ablation of different T-cell-associated selectins and/or integrins. 3. This conversion protocol induces modestly the expression of the gut-homing chemokine receptor CCR9 but not a4b7.

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4. Ex vivo-generated iTregs possess potent suppressive activity that is equal to or exceeds that of freshly isolated nTregs in in vitro models of T-cell activation as well as in a mouse model chronic gut inflammation. 5. Data obtained using this protocol predict that one healthy mouse spleen has the potential to generate between 21 and 36 million iTregs following a 4-day incubation period.

5. Notes 1. When working with TGF-b, it should be kept in mind that repeated freeze–thaw cycles should be avoided for this protein. Therefore, we recommend making single-use aliquots of TGF-b. 2. This can be scaled down if the number of iTregs required is much less than will be generated with a conversion in a full 24-well plate. 3. Prior to dissection, the mouse abdomen should be wiped 2–3 times with 70% ethanol. 4. Up to 50 million cells can be processed in a single 15-ml sterile conical centrifuge tube, and up to 200 million cells can be processed in a single 50-ml conical tube. 5. When resuspending the cell pellet in the Dynal buffer, it is important to use only part of the final volume required to resuspend the pellet to achieve optimal resuspension. Once the pellet has been uniformly resuspended, the remainder of the Dynal buffer, followed by the beads, is added to the tube. 6. We have observed that culturing cells for longer periods of time and/or adding CD28 mAb does not increase iTreg yield and may actually induce cell death such that proliferative activity decreases. Therefore, all experiments are performed in the absence of CD28 mAb with a 4-day incubation period unless otherwise noted. References 1. Sakaguchi, S. et al. (2006) Foxp3+ CD25+ CD4+ natural regulatory T cells in dominant self-tolerance and autoimmune disease. Immunol. Rev. 212, 8–27. 2. Scheffold, A., Murphy, K.M. & Hofer, T. (2007) Competition for cytokines: T(reg) cells take all. Nat. Immunol. 8, 1285–1287. 3. Uhlig, H.H. et al. (2006) Characterization of Foxp3+CD4+CD25+ and IL-10-secreting

CD4+CD25+ T cells during cure of colitis. J. Immunol. 177, 5852–5860. 4. Vignali, D.A., Collison, L.W. & Workman, C.J. (2008) How regulatory T cells work. Nat. Rev. Immunol. 8, 523–532. 5. Earle, K.E. et al. (2005) In vitro expanded human CD4+CD25+ regulatory T cells suppress effector T cell proliferation. Clin. Immunol. 115, 3–9.

Conversion of Conventional T-Cell to Tregs 6. Tang, Q. et al. (2004) In vitro-expanded antigen-specific regulatory T cells suppress autoimmune diabetes. J. Exp. Med. 199, 1455–1465. 7. Battaglia, M., Stabilini, A. & Roncarolo, M.G. (2005) Rapamycin selectively expands CD4+CD25+FoxP3+ regulatory T cells. Blood 105, 4743–4748. 8. Taylor, P.A., Lees, C.J. & Blazar, B.R. (2002) The infusion of ex vivo activated and expanded CD4(+)CD25(+) immune regulatory cells inhibits graft-versus-host disease lethality. Blood 99, 3493–3499. 9. Chen, W. et al. (2003) Conversion of peripheral CD4+. J. Exp. Med. 198, 1875–1886. 10. Benson, M.J., Pino-Lagos, K., Rosemblatt, M. & Noelle, R.J. (2007) All-trans retinoic acid mediates enhanced T reg cell growth, differentiation, and gut homing in the face of high levels of co-stimulation. J. Exp. Med. 204, 1765–1774. 11. Fantini, M.C., Dominitzki, S., Rizzo, A., Neurath, M.F. & Becker, C. (2007) In vitro generation of CD4+ CD25+ regulatory cells from murine naive T cells. Nat. Protoc. 2, 1789–1794. 12. Kang, S.G., Lim, H.W., Andrisani, O.M., Broxmeyer, H.E. & Kim, C.H. (2007) Vitamin A metabolites induce gut-homing FoxP3+ regulatory T cells. J. Immunol. 179, 3724–3733. 13. Mucida, D. et al. (2007) Reciprocal TH17 and regulatory T cell differentiation mediated by retinoic acid. Science 317, 256–260.

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14. Fontenot, J.D. et al. (2005) Regulatory T cell lineage specification by the forkhead transcription factor foxp3. Immunity 22, 329–341. 15. Zheng, S.G., Wang, J.H., Wang, P., Gray, J.D. & Horwitz, D.A. (2007) IL-2 is essential for TGF-beta to convert naive CD4(+)CD25(−) cells to CD25(+)Foxp3(+) regulatory T cells and for expansion of these cells. J. Immunol. 178, 2018–2027. 16. Sun, C.M. et al. (2007) Small intestine lamina propria dendritic cells promote de novo generation of Foxp3 T reg cells via retinoic acid. J. Exp. Med. 204, 1775–1785. 17. Hill, J.A. et al. (2008) Retinoic acid enhances Foxp3 induction indirectly by relieving inhibition from CD4+CD44hi Cells. Immunity 29, 758–770. 18. Mucida, D. et al. (2009) Retinoic acid can directly promote TGF-beta-mediated Foxp3(+) Treg cell conversion of naive T cells. Immunity 30, 471–472. 19. Thornton, A.M. & Shevach, E.M. (1998) CD4+CD25+ immunoregulatory T cells suppress polyclonal T cell activation in vitro by inhibiting interleukin 2 production. J. Exp. Med. 188, 287–296. 20. Ostanin, D.V. et al. (2009) T cell transfer model of chronic colitis: concepts, considerations, and tricks of the trade. Am. J. Physiol. Gastrointest. Liver Physiol. 296, G135–G146.

www

Chapter 5 Phenotypic and Functional Characterization of CD8+ T Regulatory Cells Séverine Ménoret, Carole Guillonneau, Séverine Bezié, Lise Caron, Ignacio Anegon, and Xian-Liang Li Abstract Increasing evidence shows the presence and significance of CD8+ T regulatory cells (CD8+ Tregs) in both human and rodent transplant recipients, as well as in autoimmune disease models. We, hereafter, review all available data on the phenotypic and functional characterization of CD8+ Tregs, and we also provide detailed protocols to purify them and analyze their suppressive function. Different subsets of dendritic cells (DCs) and CD4+ effector T cells may modulate the suppression mediated by CD8+ Tregs. By analyzing the proliferation of CFSE-labeled naïve CD4+CD25− T cells in coculture MLR and transwell experiments, we explored the mutual modulation of CD8+ Tregs, DC subsets, and CD4+ T effector cells. The suppressive function of CD8+ Tregs was mediated by both cell-contact-dependent and -independent mechanisms. Key words: CD8+ regulatory T cells, CFSE, Suppressive function, Transwell analysis, Dendritic cells, CD4+ effector T cells

1. Introduction The CD8+ suppressor T cells mediating the adoptive transfer of tolerance in mice were first identified in the 1970s by Gershon and colleagues (1, 2). In 1984, human CD8+ T cells with immune regulatory potential were demonstrated (3). These observations prompted the first series of studies for CD8+ regulatory T cells (Tregs) in the late 1980s and early 1990s. Most of the CD8+ Tregs that have been characterized for their phenotype and function are summarized in Table 1. The functional study of human CD8+ Tregs with suppressive function induced with an autologous renal allograft-derived T cell Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_5, © Springer Science+Business Media, LLC 2011

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Induced by MTB in vitro

Induced by pDCs in vivo, ovarian cancer

Induced by activated allogeneic pDCs

LAG3+

CD45RO+

CD8+

Mouse

Induced by virus

Foxp3+

Natural

Induced by epithelial cells through TGF-b

CD25+

CD25+

Induced by alloantigen

CD103+

Induced by IL-4 IL-12 and dexamethasone

Induced by CD40 activated B cells

CD8high

CD8+

Induced by anti-CD3 and TGFb

Foxp3+

Induced by anti-CD137 and polyI:C

Induced after allotransplantation

CD28+

CD8+

Induced by immature DCs pulsed with MP

CD8+

CD45RC

Natural from MHC II-deficient mice

Induced by allotransplantation

CD28−

low

Induced by anti-CD3 mAb

CD25+

Human

Generation

Identification

Species

Table 1 The different subsets of CD8+ Tregs

(36)

(35)

CD44high, GITR+, CTLA-4+ CD25+, CD28− CD25+, Foxp3+, TGF-b, IL-10

(34)

(33)

(32)

(31)

(30)

(29)

(28)

(27)

(26)

(25)

(24)

(23)

(6)

(22)

References

CD45RB+,CD28+, contact dependent, IL-10/TGFb independent

CD103+, IFNg induced TGF-b-mediated suppression

IL-10 production, cell contact dependent

Foxp3+,CD25+, CD28+, CD62L+,CD45RO+, cell contact dependent, IL-10, TGFb, CTLA-4 dependent suppression

CTLA4+, CD45RO+, CD28+, CD25+, IFN-g production

IL-10,TGFb, and CTLA4+ dependent suppression

Peptide specific and contact dependent suppression

IL-10 and TGFb

IL-10-mediated suppression

CCR7+ IL-10+, IL-10-dependent

CD25+Foxp3+CCL4+, CD28+, CD62L+

CTLA4+, GITR+, IFN-g

CD27+, Foxp3+

Foxp3+, CTLA4+, CD45RA+

Phenotypic and function and characterization

64 Ménoret et al.

Induced by uveitogenic peptide

Induced by CD40Ig

Induced by UV-B-irradiated DST

Induced by DST

Induced by oral administration of MBP

CD8+

CD8+

CD8+

CD45RC

CD45RC

low

Natural

low

CD45RClow

Natural

CD103high

Induced by oral alloantigen

Natural

Qa-1 restricted

CD8+

Induced by anti-CD137

CD11c+

Induced by DST

Induced by anti-ICOSmAb

PD1+

CD28−

Natural

CD122+

Induced by T-cell vaccination

Natural

CD122+

CD28−

Induced by peptides

Foxp3+

Antigen-driven bystander suppression

Adoptive transfer of tolerance by CD8+ and CD4+ T cells

Donor-specific unresponsiveness

Foxp3−, CD40L-, IFN-g, IDO, contact dependent and independent

Foxp3+ after stimulation

CTLA4+, IL-10, IL-4, IL13, contact dependent

IL-4, prolong graft survival

Foxp3+, contact dependent

Foxp3−, inhibit corneal xenograft rejection

CD44−, TGF-b production

CD8aa+TCRab+, killing of CD4+Qa-1+ cells

IFN-g and IDO dependent

(54)

(53)

(52)

(11), unpublished

(51)

(50)

(49)

(48)

(47)

(46)

(44, 45)

(43, 44)

(42)

(41)

CD44high, IL-10 CD45RB+, CD27+, CD45RA+, PD-1+, CD25−

(39, 40)

(38)

(37)

Foxp3−, IL-10

TGF-b, suppress anti-DNA IgG production

CD62L+, CD44low, CD45RBhigh, action through IL-10 and TGFb

GVHD graft vs. host disease, VIP vasoactive intestinal peptide, MTB Mycobacterium tuberculosis, IDO indoleamine 2,3-dioxygenase, MP influenza matrix peptide, MBP myelin basic protein

Rat

Natural

CD28−

Phenotypic and Functional Characterization of CD8+ T Regulatory Cells 65

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line was first reported in 1988 (4). The presence and expansion of CD8+ Tregs have also been associated with absence or reduced immunosuppression in transplant recipients of liver-intestine (5), heart (6, 7), and kidney allografts (8, 9). Recently, CD8+ Tregs have equally been described in human cardiac allografts with a potential to specifically inhibit antidonor immune reactivity (10). Furthermore, by blockade of CD40−CD40L interactions with CD40Ig in a rodent heart allograft model, the adoptive transfer of tolerance was mediated by tolerogenic CD8+CD45RClow regulatory T cells (11). All these observations highlight the potentially significant role of CD8+ Tregs in allograft tolerance. The crucial role of CD8+ suppressor T cells in autoimmune diseases was firstly reported in a murine model of experimental autoimmune encephalomyelitis (EAE) (12, 13). Mice that were deficient of CD8+ T cells developed more severe autoimmune myocarditis and were prone to a relapse of autoimmune arthritis, suggesting that the regulatory activity of CD8+ T cells might reside in the CD8+ memory pool (14, 15). CD8+ Tregs have also been implicated in the regulation of human autoimmune diseases such as inflammatory bowel disease and multiple sclerosis (16–18). Mounting evidence has also pointed to the role of antigen-specific CD8+ Tregs in allergy, asthma, and cancers (19, 20). As a result, the interest in the role of CD8+ Tregs has been rekindled and several types of CD8+ Tregs with distinct phenotypes and suppressive mechanisms have been described both in humans and in rodents (Table 1). The suppressive functions can be categorized into cellcontact-dependent or -independent mechanisms, and cytotoxicity to the target cells and anti-inflammatory cytokine secretion have been described in both of these categories (18) (Table 1). Subsets of DCs may influence the suppressive function of CD8+ Tregs. In a rat cardiac allograft transplantation model, plasmacytoid DCs (pDCs) promote the expression of Foxp3 in CD8+ Tregs and are needed for donor alloantigen-specific suppression. In addition, the interaction of immature CD8+ Tregs with pDCs, but not with conventional DCs, is crucial so as to induce tolerance and prevent rejection (Li et al. submitted). The in vivo outcome of DC and Treg encounters additionally depends on the effects of CD4+ effector cells, which can potentiate and modify the suppressive mechanisms of CD4+ Tregs (21). This observation introduces an important new concept concerning the suppressive mechanism of Tregs, indicating that it is not only a bidirectional regulation with DC subsets but also a triangular modulation in which T effector cells also play a key role in potentiating Treg function. To test the suppressive function of CD8+ Tregs, the coculture mixed leukocyte reaction (MLR) systems are indispensable. The conventional 3H-thymidine incorporation assay is widely used to

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assess the suppressive function of regulatory T cells (see Note 1). However, this system cannot specifically identify the different influences of APC subsets on the suppressive function because the proliferation of CD4+ T effector cells can be masked by the proliferation of other cells in the MLR system, such as CD8+ Tregs. We have, thus, developed a more specific MLR and transwell system using CFSE-labeled CD4+ T cells by which we have analyzed the suppressive mechanisms of CD8+ Tregs and the role of different APC subsets as well as CD4+ T effector cells on the CD8+ Treg function.

2. Materials 2.1. Equipment

1. A basic anesthetic delivery system with an isoflurane anesthesia facility for harvesting the spleen or lymph nodes. 2. Stainless-steel mesh, dissection instruments, and syringes, as well as needles for isolation of lymphocytes. All instruments should be cleaned and sterilized with 70% ethanol. 3. Magnetic bead separator for separating the lymphocyte subpopulations. 4. Flow cytometer; FACS LSR II for analysis, FACSAria for lymphocyte subpopulation sorting. 5. Microscope and hemocytometer for cell counting. 6. 37°C, Water-jacketed CO2 incubator with 95% humidity. 7. Calibrated pipettes and multichannel pipettes. 8. Cell strainer with a pore size of 100 or 60 mm. 9. Conical tubes: 15-ml and 50-ml BD Falcon tubes and 5-ml V-bottom sterilized tubes. 10. Culture plates and dishes: 96-well U-bottom and 24-well plates, 10-cm dishes, 3.5-cm dishes, cell culture insert with a 0.4-mm pore size. 11. Centrifuges capable of spinning for Ficoll Hypaque, Nycodenz, and washing cells.

2.2. Reagents

1. Phosphate Buffer Solution without calcium and magnesium. 2. Red blood cell lysis solution: 8.29-g NH4Cl, 1-g KHCO3, 37.2-mg Na2 EDTA dissolved in 1-l H2O, with a pH adjusted to 7.2. 3. Ficoll-Paque Plus. 4. Nycodenz: 14.5-g Nycodenz/100-ml PBS-FCS-EDTA solution.

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Table 2 Antibodies used for FACS analysis or sorting Labeled antibody

Clone

Supplier

Purified mouse anti-rat TCRgd

V65

Hybridoma from ECCC

Purified mouse anti-rat CD45R

His24

BD Biosciences

Purified mouse anti-rat NK cells

3.2.3

Hybridoma from ECCC

Purified mouse anti-rat CD11b/c

OX42

Hybridoma from ECCC

Purified mouse anti-rat T cells

R73

Hybridoma from ECCC

Biotin mouse anti-rat CD45RC

OX22

Hybridoma from ECCC

Alexa 488 mouse anti-rat TCR

R73

Hybridoma from ECCC

FITC mouse anti-rat CD6

OX56

Hybridoma from ECCC

FITC mouse anti-rat CD45R

His24

BD Biosciences

FITC mouse anti-rat TCR

R73

Hybridoma from ECCC

PE mouse anti-rat CD8a

OX8

Hybridoma from ECCC

PE mouse anti-rat CD4

OX35

Hybridoma from ECCC

PE mouse anti-rat CD45R

His24

BD Biosciences

APC mouse anti-rat CD4

OX35

Hybridoma from ECCC

APC mouse anti-rat CD103

OX62

Hybridoma from ECCC

Alexa 647 mouse anti-rat CD25

OX39

Hybridoma from ECCC

5. Collagenase D: 1 g collagenase in 500-ml RPMI-1640 + 5-ml Hepes + 2% fetal bovine serum (FCS). 6. Antibodies for FACS analysis or cell sorting: see Table 2. 7. PBS-FCS-EDTA solution: 500-ml PBS + 10-ml FCS + 2.5-ml 0.1 M EDTA. 8. Microbeads: Goat anti-mouse IgG (see Note 2). 9. 500 ml complete RPMI-1640 medium with 5-ml penicillin (80 U/ml)-streptomycin (80 mg/ml), 5-ml L-glutamine, 5-ml nonessential amino acids (100×), 5-ml pyruvate sodium (100 mM), 5-ml HEPES buffer (1 M), 2.5-ml b-mercaptomethanol (7 ml of 2-b mercaptoethanol stock diluted in 10-ml RPMI) (see Notes 3 and 4). 10. Fetal bovine serum (see Note 3). 2.3. Animals and Other Antibodies

1. Male Lewis 1W and Lewis 1A Rats (8 weeks) (see Note 5). 2. A description of antibodies used is provided in Table 2.

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3. Methods 3.1. Protocols for the Analysis of Regulatory CD8+ T Cell Suppressive Function

3.2. Protocol for Harvesting the Spleen or Other Organs

3.2.1. Protocol for Harvesting the Spleen

Analyzing antigen-specific CD4+ T-cell proliferation is a major technique for assessing the suppressive capacity of CD8+ Tregs. CD4+ lymphocytes proliferate in response to antigenic peptides associated with class II major histocompatibility complex (MHC) molecules on APCs. A mixed lymphocyte proliferation (MLR) assay measures the ability of lymphocytes placed in short-term tissue culture to proliferate when stimulated in vitro by donor alloantigens. Our method involves isolating and labeling naive CD4+ CD25− T lymphocytes with CFSE, placing the cells in a 96-well plate with or without CD8+CD45RClow regulatory T cells, and stimulating with various stimuli, such as allogeneic plasmacytoid or conventional DCs, as well as recipient DC subsets loaded with donor alloantigens. The cells are incubated for six days at 37°C in a CO2 incubator. A transwell system can be used to test the contact dependency of Treg–DC cells, or Treg–Teff cells. Proliferation is quantified on the 6th day by analyzing the dilution of CFSE (excitation, 488 nm; detection, 530 nm) on a LSR II cytometer. After gating of TCR+CD4+ T cells, the proliferation is shown by a histogram of CFSE intensity. 1. Anesthetize 8-week-old male Lewis 1A and Lewis 1W rats by isofluorane (rat body weight around 250 g). 2. Sterilize the abdomen of the animals with 70% ethanol. All surgical procedures should be performed with sterilized instruments. 1. Make a surgical incision in the abdomen with the dissection scissors. 2. Expose the spleen, separate from other organs, and ligate the blood vessels of the spleen with 4–0 surgical thread, then remove it aseptically without the fat tissues by forceps and scissors. 3. Store the spleen in cold PBS solution in a 50-ml BD Falcon tube on ice.

3.2.2. Protocol for Harvesting the Donor Heart for Indirect Antigen Presentation Analysis

1. Make a middle thoracic incision exposing the rib cage and cut bilaterally. 2. Expose the heart by removing the rib cage and harvest the heart without the thymus using sterile instruments. 3. Store the donor heart in cold PBS solution in a 50-ml BD Falcon tube on ice.

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3.3. Protocol for the Purification of T Lymphocyte Subpopulations

1. Collect the spleen from the storage tube and transfer to a 10-ml culture dish with 3–4 ml of PBS.

3.3.1. Preparation of T Lymphocytes: CD8+ Tregs, CD4+CD25− T Cells, and CD4+CD25+ Tregs

3. Prepare another 10-ml dish with a sterilized steel mesh, transfer the small pieces of spleen with a 10-ml pipette onto the mesh, and add 15 ml of PBS.

2. Mince the spleen into small pieces with sterilized surgical blades.

4. Further disrupt the spleen tissue by pressuring using the syringe column. 5. Collect the suspension of splenocytes in a 50-ml tube and wash the dish with cold PBS. 6. Centrifuge the suspension of splenocytes for 10 min at 524 × g, 4°C. 7. Remove the supernatant and resuspend the pellet of splenocytes with 10 ml of red cell lysis buffer for at least 5 min at room temperature. 8. Wash the splenocytes twice with 50 ml of PBS to remove the lysis buffer. 9. After washing, resuspend the splenocytes in 10-ml PBS solution, filter the splenocyte suspension with a cell strainer with a pore size of 100 mm, and collect the splenocytes in another 50-ml BD Falcon tube. 10. Count the splenocyte suspension with a hemocytometer and resuspend it at a concentration of 20–30 × 106 cells/ml in PBS-FCS-EDTA solution. 11. To enrich for T cells, incubate the splenocyte suspension for at least 10 min or more at 4°C with a cocktail of antibodies against: gdT cells (anti-TCRgd, clone V65), B cells (anti-CD45R, clone His24), NK cells (anti-CD161, clone 3.2.3), and monocytes (anti-CD11b/c, clone OX42). 12. After the incubation of the cocktail of antibodies, wash the splenocytes twice with PBS-FCS-EDTA solution at 524 × g for 10 min, 4°C. 13. During the washing of the antibodies, prepare the Dynabeads by calculating the number of cells that require purifying, for example, to enrich T lymphocytes, 50% of the total number of splenocytes need to be removed using four beads per cell. Wash the Dynabeads at least three times with PBS-FCS-EDTA solution using a magnetic bead separator. After washing, resuspend the Dynabeads at a 1:10 dilution in PBS-FCSEDTA solution. (see Note 2). 14. Resuspend the splenocyte pellet in magnetic Dynabeads and incubate under slight agitation in a solar agitator for 10 min at 4°C (cold room).

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15. After incubation, add 30 ml of PBS-FCS-EDTA solution to the tube and separate the splenocytes using a magnetic bead separator by collecting the suspension of the cells in another tube after at least 1-min incubation in the separator. Repeat at least three times to remove all the conjugated beads. 16. Wash the enriched T lymphocytes at least twice by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution. 17. Count the T lymphocytes and resuspend at 20–30 × 106 cells/ml. 18. Label the enriched T cells with a cocktail of fluorescence antibodies for at least 20 min or more at 4°C in the dark (anti-CD45RC-biotin (OX22), anti-CD8a-PE (OX8) or anti-CD4–PE (OX35), anti-CD6-FITC (OX56), and antiCD25-Alexa 647 (OX39)). 19. Wash the enriched T lymphocytes at least twice by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution. 20. Resuspend the T lymphocytes at 30 × 106 cells/ml in cold PBS-FCS-EDTA solution and label the secondary antibodies with Strepavidin-PE-Cy7 in the dark for at least 15 min at 4°C. 21. Wash the enriched T lymphocytes at least twice by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution. 22. Count the T lymphocyte suspension with a hemocytometer and resuspend at a concentration of 30 × 106 cells/ml in cold PBS-FCS-EDTA solution. Filter the cell suspension using a cell strainer with a pore size of 60 mm. 23. Add DAPI to the cell suspension (10 mg/ml) to exclude dead cells in the FACS analysis. 24. Sort the subpopulations of T lymphocytes using a FACSAria in the following gates, CD6+CD8+CD45RClow T cells, CD6+CD4+CD25+ T cells, and CD6+CD4+CD25− T cells. The purity of the sorted subpopulations should be systematically more than 98%. 25. Refer to the pseudocolor plot display of the FACSAria to sort the targeted population (Fig. 1). (The figures are provided with the permission of the authors.) 3.4. Preparation of DC Subsets

1. Collect the spleen from the storage tube and transfer to a 5-ml culture dish and add 3–4 ml Collagenase D, prewarmed in a 37°C water bath. 2. Inject collagenase D with syringes into the spleen and mince the spleen into small pieces with sterilized surgical blades.

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CD8 PE

CD4 PE

Fig. 1. The pseudocolor plot display of the FACSAria for sorting of the target population. Sort the subpopulations of T lymphocytes using a FACSAria in the following gates: CD6+CD8+CD45RClow T cells, CD6+CD4+CD25+ T cells, and CD6+CD4+CD25− T cells. Left panel, CD8+CD45RClow T cells and right panel, CD4+CD25+ T cells and CD4+CD25− T cells.

3. Incubate the spleen pieces with collagenase D at 37°C in a water-jacketed CO2 incubator for at least 30 min. 4. After incubation, add 0.4 ml of 0.1 M EDTA to stop the reaction. 5. Prepare another 10-ml dish with a sterilized steel mesh, transfer the small pieces of spleen with a 10-ml pipette onto the mesh, and add 15 ml of PBS-FCS-EDTA solution (see Note 4). 6. Further lyse the spleen tissue by syringe column pressure or pipette. 7. Collect the suspension of splenocytes in a 50-ml tube and wash the dish with cold PBS-FCS-EDTA solution. 8. Wash the splenocytes with 524 × g for10 min at 4°C in PBS-FCS-EDTA solution. 9. After washing, resuspend the splenocytes in 8-ml PBS-FCSEDTA solution, filter the splenocyte suspension using a cell strainer with a pore size of 100 mm, and collect the splenocytes in another 50-ml BD tube. 10. Prepare 4 ml of 14.5% Nycodenz in two 15-ml BD tubes. Very slowly, load 4 ml of splenocytes onto each of the Nycodenz layers. 11. Centrifuge the splenocytes with Nycodenz for 13 min at 1,825 × g, 4°C without braking. 12. After centrifugation, collect the cells from the interface; which are enriched in conventional DCs (cDCs). Wash the enriched cDCs with PBS-FCS-EDTA solution by centrifuging for 10 min at 630 × g, 4°C. 13. Rewash the enriched cDCs by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution and count the cells. Resuspend at a concentration of 30 × 106 cells/ml in cold PBS-FCS-EDTA solution and ready for staining.

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14. Incubate with the following cocktail of antibodies used to label cDCs: anti-TCR (R73-alexa 488), anti-CD45RA (clone OX33-FITC), CD103 (OX62-APC), and anti-CD4 (OX35-PE) on ice for at least 10 min. 15. Collect the cell pellet after running the Nycodenz centrifugation and resuspend in 15 ml of PBS-FCS-EDTA solution. 16. Prepare 15 ml of Ficoll Hypaque Plus in 50-ml BD tubes, and very slowly add 7.5 ml of resuspended splenocytes onto the Ficoll Hypaque Plus solution. 17. Centrifuge for 20 min at 931 × g and at room temperature without braking. 18. Collect the cells from the interface and wash with cold PBSFCS-EDTA for 10 min at 630 g, 4°C. Resuspend at a concentration of 30 × 106 cells/ml in cold PBS-FCS-EDTA solution, ready for staining. 19. To further enrich the pDC population, incubate the cells collected from Ficoll centrifugation with a cocktail of antibodies staining T cells (clone R73), gdT cells (anti-TCRgd, clone V65), and B cells (anti-CD45RA, clone OX33) for at least 10 min at 4°C. 20. After the incubation of the antibody cocktail, wash the cells twice with PBS-FCS-EDTA solution for10 min at 524 × g, 4°C. 21. During the washing of the antibodies, prepare the Dynabeads by calculating the number of cells requiring purification: for example, to enrich for pDCs, at least 50% of total splenocytes need to be removed using four beads per cell. Wash the Dynabeads at least three times in PBS-FCS-EDTA solution using a magnetic beads separator. After washing, resuspend the Dynabeads at a 1:10 dilution in PBS-FCS-EDTA solution (see Note 2). 22. Resuspend the cell pellet with washed magnetic Dynabeads and incubate under slight agitation in a solar agitator for 10 min at 4°C (cold room). 23. After incubation, add 30 ml of PBS-FCS-EDTA solution to the tube and separate the splenocytes using a magnetic bead separator by collecting the cell suspension in another tube after at least 1-min incubation in the separator. Repeat at least three times to remove all conjugated beads. 24. Wash the enriched pDC population by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution, at least twice. Count and resuspend at 20–30 × 106 cells/ml. 25. The following cocktail of antibodies should be used to label pDCs: anti-TCR (clone R73-FITC), anti-CD45R (clone His24-PE), and anti-CD4 (clone OX35-APC) for at least 10 min at 4°C.

OX62 A lexa 647

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CD45R PE

pD Cs

CD4 PE

CD 4-cD Cs/CD 4+ cD Cs

Fig. 2. The pseudocolor plot display of the FACSAria for sorting of the target population. Sort the subpopulations of DCs using a FACSAria in the following gates: cDCs are sorted as TCR−CD45RA−CD103+ cells and further classified by the expression of CD4 into CD4+ cDCs and CD4− cDCs subpopulations. pDCs are defined as TCR−CD45RA−CD45R+CD4+ cells. Left panel, pDCs, right panel, CD4+/CD4−cDCs.

26. Wash the fluorescence antibody-labeled cell populations at least twice by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution. 27. Count the cell suspension with a hemocytometer and resuspend at a concentration of 30 × 106 cells/ml in cold PBS-FCSEDTA solution. Filter the cell suspension using a cell strainer with a pore size of 60 mm. 28. Add DAPI to the cell suspension (10 mg/ml) to exclude dead cells in the FACS analysis. 29. Sort the subpopulations of DCs using a FACSaria in the following gates: cDCs are sorted as TCR−CD45RA−CD103+ cells and further classified by the expression of CD4 into CD4+ cDCs and CD4− cDCs subpopulations. pDCs are defined as TCR−CD45RA−CD45R+CD4+ cells. The purity of the sorted subpopulations should be systematically greater than 98%. 30. Refer to the pseudocolor plot display of the FACSaria to sort the target population (Fig. 2). (The figures are provided with the permission of the authors.) 3.5. CFSE Labeling of Responder T Cells and MLR by the Direct Alloantigen Presentation Pathway

1. Wash the sorted CD4+CD25− T cells with PBS solution twice and dilute with PBS to a concentration of 25 × 106/6 ml (see Note 6). 2. Label naive CD4+CD25− T cells with 3 ml of 5-carboxy fluorescein succinimidyl ester (CFSE) (5 mM) for 3–5 min at room temperature. 3. Stop the labeling reaction by adding 1.25 ml of FCS for 1 min at room temperature, and wash the cells twice with complete RPMI-1640. Then, count the cell numbers.

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In normal circumstances, 30–50% of cells will die during the labeling process. 4. If the cells numbers are less than 5 × 10 6, the cells should be resuspended in 2-ml PBS and stained with 2-ml CFSE. The reaction is stopped with 500-ml FCS. 5. To assess the suppressive function of regulatory T cells, coculture MLR experiments are used to analyze the proliferation of naive CD4+CD25− T cells. Purified naive CD4+CD25− T cells labeled with CFSE (2 × 104 cells) are cocultured with cDCs or pDCs (5 × 103 cells, CD4+ T cell/DC ratio of 4/1) in a round-bottom 96-well plate in a final volume of 200 ml of complete RPMI-1640 medium, with or without suppressive cells, for 6 days at 37°C in 5% CO2. The ratio of suppressor cells to effector cells should be varied from 1:1 to 0.1/1. 6. To investigate the cell–cell contact dependency, a transwell experiment is used to analyze the proliferation of naive CD4+CD25− T cells. Load the purified naive CD4+CD25− T cells (4 × 105 cells) labeled with CFSE and sorted DCs (105 cells) in a V-bottom tube (5 ml) as the lower chamber. Then, add a cell culture insert with a 0.4-mm HD pore size membrane into the V tube as the upper chamber. Load the CD8+ Tregs (0.4 × 106 cells) with or without sorted DCs onto the cell culture insert membrane. The cells are cultured at 37°C in 5% CO2 in a final volume of 1 ml of complete RPMI-1640 medium for 6 days (see Note 7). 7. After 6 days of culture, the cells from the coculture MLR experiment or from the lower chamber of the transwell system are harvested and stained with TCR (R73-biotin, further conjugated with Strepavidin-PE-Cy7) and CD4 (OX35-PE) in V-bottom 96-well plates. 8. The proliferation of CFSE-labeled naive CD4+CD25− T cells induced by allogeneic DCs is analyzed by the dilution of CFSE (excitation, 488 nm; detection, 530 nm) on a LSR II cytometer. After gating of TCR+CD4+ T cells, the proliferation is shown by a histogram of CFSE intensity. The data are analyzed using FLOWJO software. 9. The suppressive function assay is performed by analyzing the dilution of CFSE-labeled naive CD4+CD25− T cells in the presence of immature pDCs after 6 days of MLR coculture (Fig. 3). (The figures are provided with the permission of the authors.) 10. The suppressive function assay is performed by analyzing the dilution of CFSE-labeled naive CD4+CD25− T cells in the presence of immature pDCs after 6 days of the transwell experiment (Fig. 4). (The figures are provided with the permission of the authors.)

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Medium + CD4+ CD25−

pDCs−1W + CD4+ CD25-

CD8+ Tregs + pDCs−1W + CD4+ CD25-

CFSE

Fig. 3. The suppressive function assay by analyzing the dilution of CFSE-labeled naive CD4+CD25− T cells in the presence of immature pDCs after 6 days of coculture MLR. Left panel: there is no proliferation of the naive CD4+CD25− T cells with medium alone. Middle panel: strong proliferation of naive CD4+CD25− T cells with the stimulation of allogeneic pDCs. Right panel: the proliferation of naive CD4+CD25− T cells is strongly inhibited by naive CD8+CD45RClow regulatory T cells.

Upper chamber Lower chamber

Medium CD4+ CD25−

Medium CD4+ CD25− pDCs-1W

CD8+ Tregs pDCs

CD8+ Tregs

CD4+ CD25− pDCs-1W

CD4+ CD25− pDCs-1W

CFSE

Fig. 4. The suppressive function assay by analyzing the dilution of CFSE-labeled naive CD4+CD25− T cells in the presence of immature pDCs after 6 days of the transwell experiment. Left panel: there is no proliferation of the naive CD4+CD25− T cells with medium alone. Left middle panel: strong proliferation of naive CD4+CD25− T cells with the stimulation of allogeneic immature pDCs. Right middle panel: the proliferation of naive CD4+CD25− T cells is strongly inhibited when the naive CD8+CD45RClow regulatory T cells and allogeneic immature pDCs are loaded in the upper chamber. Right panel: there is no suppression by naive CD8+CD45RClow regulatory T cells if there are no allogeneic pDCs in the upper chamber.

3.6. MLR by the Indirect Alloantigen Presentation Pathway

1. Collect the donor heart from the storage tube and transfer to a 5-ml culture dish, add 3–4 ml collagenase D, prewarmed in a 37°C water bath (see Note 8). 2. Inject collagenase D with syringes into the heart and mince it into small pieces with sterilized surgical blades.

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3. Incubate naive heart pieces with collagenase D at 37°C in a Water-jacketed CO2 incubator for at least 45 min. 4. After incubation, add 0.4 ml of 0.1 M EDTA to stop the reaction. 5. Prepare another 10-ml dish with a sterilized steel mesh, transfer the small pieces with a 10-ml pipette onto the mesh, and add 15 ml of PBS-FCS-EDTA solution. 6. Further disrupt the heart tissue by applying pressure using the syringe column. 7. Collect the suspension of cells in a 50-ml tube and wash the dish with cold PBS-FCS-EDTA solution. 8. Wash the cells by centrifuging at 524 × g for 10 min at 4°C in PBS-FCS-EDTA solution. 9. After washing, resuspend the cells in 10-ml PBS-FCS-EDTA solution, filter the cell suspension using a cell strainer with a 100-mm pore size, and collect the cells in another 50-ml BD tube. 10. Prepare 15 ml of Ficoll Hypaque Plus in 50-ml BD Falcon tubes and very slowly add 5 ml of resuspended cells onto the Ficoll Hypaque Plus solution layer. 11. Centrifuge for 20 min at 930 × g and at room temperature without braking. 12. Collect the cells from the interface: these are naive heart infiltrating lymphocytes. Wash with cold PBS-FCS-EDTA for 10 min by centrifuging at 630 × g, 4°C and count the numbers. 13. Apoptosis of naive heart infiltrating lymphocytes is induced by UV irradiation (TUV 30 W/G 30 TB; Philips, The Netherlands; wavelength 280–340 nm), UV irradiation at room temperature for 3 h in complete RPMI-1640 medium. 14. Apoptotic cells are cocultured overnight with recipient (LEW.1A) DC subsets in a 96-well U-bottom culture plate, with complete RPMI-1640 culture medium at 37°C in a 5% CO2 incubator. The ratio of the apoptotic cells and DC subset is 1:1 or less. The concentration of the coculture cells is 1 × 106/ml. 15. After one night of coculture, Ficoll gradient centrifugation is used to harvest DCs loaded with donor alloantigens and to eliminate apoptotic cells. Wash the cells collected from the inter-phase with cold PBS-FCS-EDTA by centrifuging for 10 min at 630 × g, 4°C and count the numbers. Resuspend the cells in complete RPMI-1640 medium and use for the MLR experiment.

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Medium CD4+ CD25−

pDC-1A CD4+ CD25-

Apoptotic cells CD4+ CD25-

pDC-1A Apoptotic cells CD4+ CD25−

CD8 + Tregs pDC-1A Apoptotic cells CD4+ CD25-

CFSE

Fig. 5. The suppressive function assay by analyzing the dilution of CFSE-labeled naive CD4+CD25− T cells in the presence of indirect allogeneic stimulation after 6 days of MLR coculture. There is no proliferation of the naive CD4+CD25− T cells with medium alone, recipient immature pDCs, or the donor apoptotic naive heart resident leukocytes. A strong proliferation of naive CD4+CD25− T cells with the stimulation of recipient immature pDCs after loading with donor apoptotic naive heart resident leukocytes, and strong inhibition when the naive CD8+CD45RClow regulatory T cells are present.

16. The suppressive function assay is performed by analyzing the dilution of CFSE-labeled naive CD4+CD25− T cells in the presence of indirect allogeneic stimulation after 6 days of coculture MLR (Fig. 5). (The figures are provided with the permission of the authors.)

4. Notes 1. The 3H incorporation assay is not recommended since the proliferation of DCs and suppressor cells cannot be excluded. 2. In some cases, the toxicity of Dynabeads or its stock solution will strongly influence the viability of T cells and DC subsets. The beads must be washed thoroughly. In addition, using less quantity of beads may partly prevent toxicity. 3. Check each individual serum before using in MLRs to ensure that each batch qualifies for proliferation assays (Fig. 6). Additionally, sera from different companies will exert a significant influence on the suppressive function of CD8+ Tregs. Finally, the same culture medium from different companies also influences the MLRs, the function of the CD8 Tregs, and the function and viability of DC subsets. Thus, the identification of the best combination of serum and coculture medium is very important to ensure consistent and reproducible results. For the functional analysis of rat CD8+ Tregs,

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Fig. 6. The influence of sera on the suppressive function of CD8+CD45RClow regulatory T cells. Purified CD4+CD25− T cells labeled with CFSE were cocultured with allogeneic pDCs for 6 days with or without the suppressive CD8+CD45RClow T regulatory cells. After 6 days of coculture, proliferation of CFSE-labeled naive CD4+CD25− T cells is analyzed by the dilution of CFSE after gating of TCR+CD4+ T cells. The proliferation is shown by a histogram of CFSE intensity.

FCS from Lonza Verniers and RPMI-1640 culture medium from Sigma are reliable. (The figures are provided with the permission of the authors.) 4. Always use PBS-FCS-EDTA solution for purification of the DC subsets and perform all the procedures in ice. RPMI-1640 medium with L-glutamine should be used for cell culture, and L-glutamine should be added when the medium is used. Fresh L-glutamine should be added if this is not used within a month. 5. Rats should be kept under specific pathogen-free conditions and used at 8–16 weeks of age. 6. Prepare enough number of naive CD4+CD25− T cells before labeling the CFSE since this process could cause the cells to die. 7. V-bottom tubes, but not flat-bottom transwell culture plates, are required to ensure optimal cell–cell contact. It is vital that a perfect proliferation of naive CD4+CD25− T cells is obtained in the transwell experiment. 8. For alloantigen indirect presentation pathway analysis, the use of naive donor heart leukocytes, but not donor spleen, is paramount, since apoptotic donor naive heart infiltrating cells do not induce any proliferation of naive recipient T cells, while donor splenocytes do. For testing the phagocytosis function of recipient DCs, donor naive heart leukocytes could be labeled with PKH prior to the irradiation. The uptake of PKH fluorescence by recipient DC subsets can be traced by confocal microscopy (Fig. 7). The figures are provided with the permission of the authors.

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Fig. 7. For testing the phagocytosis function of recipient DCs. Donor naive heart infiltrating cells can be labeled with PKH prior to irradiation. The uptake of PKH fluorescence by recipient DCs subsets can be traced by confocal microscopy. Left panel: unloaded recipient pDCs immediately after FACS sorting and labeling nuclei with Topro3 (confocal analysis, ×63). Right panel: recipient pDCs loaded for 24 h with donor PKH-labeled apoptotic cells (nuclei labeled with Topro3, confocal analysis, ×63).

Acknowledgments This work was supported by a research fellowship from the Transplantation Society and the Senior Basic Science Grant from the European Society for Organ Transplantation (to X.L.L.) as well the grants from the Roche Organ Transplantation Research Foundation and the Fondation Transvie (to I.A.).

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References 1. Gershon, R.K. and Kondo, K. 1971. Infectious immunological tolerance. Immunology 21:903–914. 2. Cantor, H., Shen, F.W., and Boyse, E.A. 1976. Separation of helper T cells from suppressor T cells expressing different Ly components. II. Activation by antigen: after immunization, antigen-specific suppressor and helper activities are mediated by distinct T-cell subclasses. J Exp Med 143: 1391–1340. 3. Thomas, Y., Rogozinski, L., Rabbani, L., Chess, A., Goldstein, G., and Chess, L. 1984. Functional analysis of human T cell subsets defined by monoclonal antibodies. VI. Distinct and opposing immunoregulatory functions within the OKT8+ population. J Mol Cell Immunol 1:103–113. 4. Emara, M., Miceli, M.C., Finn, O.J., and Sanfilippo, F. 1988. Human suppressor T cells induced in vitro with an autologous renal allograft-derived T cell line. I. Suppressor cell induction, function, and specificity. Hum Immunol 23:223–240. 5. Sindhi, R., Manavalan, J.S., Magill, A., Suciu-Foca, N., and Zeevi, A. 2005. Reduced immunosuppression in pediatric liver-intestine transplant recipients with CD8+CD28− Tsuppressor cells. Hum Immunol 66:252–257. 6. Colovai, A.I., Mirza, M., Vlad, G., Wang, S., Ho, E., Cortesini, R., and Suciu-Foca, N. 2003. Regulatory CD8+CD28− T cells in heart transplant recipients. Hum Immunol 64:31–37. 7. Manavalan, J.S., Kim-Schulze, S., Scotto, L., Naiyer, A.J., Vlad, G., Colombo, P.C., Marboe, C., Mancini, D., Cortesini, R., and Suciu-Foca, N. 2004. Alloantigen specific CD8+CD28− FOXP3+ T suppressor cells induce ILT3+ ILT4+ tolerogenic endothelial cells, inhibiting alloreactivity. Int Immunol 16:1055–1068. 8. Ciubotariu, R., Vasilescu, R., Ho, E., Cinti, P., Cancedda, C., Poli, L., Late, M., Liu, Z., Berloco, P., Cortesini, R., et al. 2001. Detection of T suppressor cells in patients with organ allografts. Hum Immunol 62:15–20. 9. Trzonkowski, P., Zilvetti, M., Chapman, S., Wieckiewicz, J., Sutherland, A., Friend, P., and Wood, K.J. 2008. Homeostatic repopulation by CD28−CD8+ T cells in alemtuzumabdepleted kidney transplant recipients treated with reduced immunosuppression. Am J Transplant 8:338–347. 10. Dijke, I.E., Caliskan, K., Klepper, M., de Kuiper, R., Balk, A.H., Maat, A.P., Weimar, W.,

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

and Baan, C.C. 2009. Donor-specific immune regulation by CD8 lymphocytes expanded from rejecting human cardiac allografts. Am J Transplant 9:397–403. Guillonneau, C., Hill, M., Hubert, F.X., Chiffoleau, E., Herve, C., Li, X.L., Heslan, M., Usal, C., Tesson, L., Menoret, S., et al. 2007. CD40Ig treatment results in allograft acceptance mediated by CD8CD45RC T cells, IFN-gamma, and indoleamine 2,3-dioxygenase. J Clin Invest 117:1096–1106. Jiang, H., Zhang, S.I., and Pernis, B. 1992. Role of CD8+ T cells in murine experimental allergic encephalomyelitis. Science 256: 1213–1215. Koh, D.R., Fung-Leung, W.P., Ho, A., Gray, D., Acha-Orbea, H., and Mak, T.W. 1992. Less mortality but more relapses in experimental allergic encephalomyelitis in CD8−/− mice. Science 256:1210–1213. Penninger, J.M., Neu, N., Timms, E., Wallace, V.A., Koh, D.R., Kishihara, K., Pummerer, C., and Mak, T.W. 1993. The induction of experimental autoimmune myocarditis in mice lacking CD4 or CD8 molecules (corrected). J Exp Med 178:1837–1842. Tada, Y., Ho, A., Koh, D.R., and Mak, T.W. 1996. Collagen-induced arthritis in CD4- or CD8-deficient mice: CD8+ T cells play a role in initiation and regulate recovery phase of collagen-induced arthritis. J Immunol 156: 4520–4526. Brimnes, J., Allez, M., Dotan, I., Shao, L., Nakazawa, A., and Mayer, L. 2005. Defects in CD8+ regulatory T cells in the lamina propria of patients with inflammatory bowel disease. J Immunol 174:5814–5822. Tennakoon, D.K., Mehta, R.S., Ortega, S.B., Bhoj, V., Racke, M.K., and Karandikar, N.J. 2006. Therapeutic induction of regulatory, cytotoxic CD8+ T cells in multiple sclerosis. J Immunol 176:7119–7129. Suzuki, M., Konya, C., Goronzy, J.J., and Weyand, C.M. 2008. Inhibitory CD8+ T cells in autoimmune disease. Hum Immunol 69: 781–789. Vlad, G. and Suciu-Foca, N. 2008. Resurgence or emergence of CD8+ Ts. Hum Immunol 69:679–680. Niederkorn, J.Y. 2008. Emerging concepts in CD8(+) T regulatory cells. Curr Opin Immunol 20:327–331. Collison, L.W., Pillai, M.R., Chaturvedi, V., and Vignali, D.A. 2009. Regulatory T cell suppression is potentiated by target T cells in

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23.

24.

25.

26.

27.

28.

29.

30.

31.

Ménoret et al. a cell contact, IL-35- and IL-10-dependent manner. J Immunol 182:6121–6128. Bisikirska, B., Colgan, J., Luban, J., Bluestone, J.A., and Herold, K.C. 2005. TCR stimulation with modified anti-CD3 mAb expands CD8+ T cell population and induces CD8+CD25+ Tregs. J Clin Invest 115:2904–2913. Billerbeck, E., Blum, H.E., and Thimme, R. 2007. Parallel expansion of human virusspecific FoxP3− effector memory and de novogenerated FoxP3+ regulatory CD8+ T cells upon antigen recognition in vitro. J Immunol 179:1039–1048. Joosten, S.A., van Meijgaarden, K.E., Savage, N.D., de Boer, T., Triebel, F., van der Wal, A., de Heer, E., Klein, M.R., Geluk, A., and Ottenhoff, T.H. 2007. Identification of a human CD8+ regulatory T cell subset that mediates suppression through the chemokine CC chemokine ligand 4. Proc Natl Acad Sci U S A 104:8029–8034. Wei, S., Kryczek, I., Zou, L., Daniel, B., Cheng, P., Mottram, P., Curiel, T., Lange, A., and Zou, W. 2005. Plasmacytoid dendritic cells induce CD8+ regulatory T cells in human ovarian carcinoma. Cancer Res 65: 5020–5026. Gilliet, M. and Liu, Y.J. 2002. Generation of human CD8 T regulatory cells by CD40 ligand-activated plasmacytoid dendritic cells. J Exp Med 195:695–704. Ordonez, L., Bernard, I., L’Faqihi-Olive, F.E., Tervaert, J.W., Damoiseaux, J., and Saoudi, A. 2009. CD45RC isoform expression identifies functionally distinct T cell subsets differentially distributed between healthy individuals and AAV patients. PLoS One 4:e5287. Dhodapkar, M.V. and Steinman, R.M. 2002. Antigen-bearing immature dendritic cells induce peptide-specific CD8(+) regulatory T cells in vivo in humans. Blood 100:174–177. Cai, J., Lee, J., Jankowska-Gan, E., Derks, R., Pool, J., Mutis, T., Goulmy, E., and Burlingham, W.J. 2004. Minor H antigen HA-1-specific regulator and effector CD8+ T cells, and HA-1 microchimerism, in allograft tolerance. J Exp Med 199:1017–1023. Siegmund, K., Ruckert, B., Ouaked, N., Burgler, S., Speiser, A., Akdis, C.A., and Schmidt-Weber, C.B. 2009. Unique phenotype of human tonsillar and in vitro-induced FOXP3+CD8+ T cells. J Immunol 182: 2124–2130. Zheng, J., Liu, Y., Qin, G., Chan, P.L., Mao, H., Lam, K.T., Lewis, D.B., Lau, Y.L., and

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

Tu, W. 2009. Efficient induction and expansion of human alloantigen-specific CD8 regulatory T cells from naive precursors by CD40-activated B cells. J Immunol 183:3742–3750. Uss, E., Rowshani, A.T., Hooibrink, B., Lardy, N.M., van Lier, R.A., and ten Berge, I.J. 2006. CD103 is a marker for alloantigen-induced regulatory CD8+ T cells. J Immunol 177: 2775–2783. Myers, L., Croft, M., Kwon, B.S., Mittler, R.S., and Vella, A.T. 2005. Peptide-specific CD8 T regulatory cells use IFN-gamma to elaborate TGF-beta-based suppression. J Immunol 174:7625–7632. Noble, A., Giorgini, A., and Leggat, J.A. 2006. Cytokine-induced IL-10-secreting CD8 T cells represent a phenotypically distinct suppressor T-cell lineage. Blood 107:4475–4483. Bienvenu, B., Martin, B., Auffray, C., Cordier, C., Becourt, C., and Lucas, B. 2005. Peripheral CD8+CD25+ T lymphocytes from MHC class II-deficient mice exhibit regulatory activity. J Immunol 175:246–253. Sugita, S., Futagami, Y., Horie, S., and Mochizuki, M. 2007. Transforming growth factor beta-producing Foxp3(+)CD8(+) CD25(+) T cells induced by iris pigment epithelial cells display regulatory phenotype and acquire regulatory functions. Exp Eye Res 85:626–636. Menager-Marcq, I., Pomie, C., Romagnoli, P., and van Meerwijk, J.P. 2006. CD8(+)CD28(−) Regulatory T lymphocytes prevent experimental inflammatory bowel disease in mice. Gastroenterology 131:1775–1785. Hahn, B.H., Singh, R.P., La Cava, A., and Ebling, F.M. 2005. Tolerogenic treatment of lupus mice with consensus peptide induces Foxp3-expressing, apoptosis-resistant, TGFbetasecreting CD8+ T cell suppressors. J Immunol 175:7728–7737. Rifa’i, M., Kawamoto, Y., Nakashima, I., and Suzuki, H. 2004. Essential roles of CD8+CD122+ regulatory T cells in the maintenance of T cell homeostasis. J Exp Med 200:1123–1134. Endharti, A.T., Rifa’I, M., Shi, Z., Fukuoka, Y., Nakahara, Y., Kawamoto, Y., Takeda, K., Isobe, K., and Suzuki, H. 2005. Cutting edge: CD8+CD122+ regulatory T cells produce IL-10 to suppress IFN-gamma production and proliferation of CD8+ T cells. J Immunol 175:7093–7097. Chen, X., Priatel, J.J., Chow, M.T., and Teh, H.S. 2008. Preferential development of CD4

Phenotypic and Functional Characterization of CD8+ T Regulatory Cells

42.

43.

44.

45.

46.

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and CD8 T regulatory cells in RasGRP1deficient mice. J Immunol 180:5973–5982. Izawa, A., Yamaura, K., Albin, M.J., Jurewicz, M., Tanaka, K., Clarkson, M.R., Ueno, T., Habicht, A., Freeman, G.J., Yagita, H., et al. 2007. A novel alloantigen-specific CD8+PD1+ regulatory T cell induced by ICOS-B7h blockade in vivo. J Immunol 179: 786–796. Ju, S.A., Park, S.M., Lee, S.C., Kwon, B.S., and Kim, B.S. 2007. Marked expansion of CD11c+CD8+ T-cells in melanoma-bearing mice induced by anti-4-1BB monoclonal antibody. Mol Cells 24:132–138. Seo, S.K., Choi, J.H., Kim, Y.H., Kang, W.J., Park, H.Y., Suh, J.H., Choi, B.K., Vinay, D.S., and Kwon, B.S. 2004. 4-1BB-mediated immunotherapy of rheumatoid arthritis. Nat Med 10:1088–1094. Hu, D., Ikizawa, K., Lu, L., Sanchirico, M.E., Shinohara, M.L., and Cantor, H. 2004. Analysis of regulatory CD8 T cells in Qa-1deficient mice. Nat Immunol 5:516–523. Ho, J., Kurtz, C.C., Naganuma, M., Ernst, P.B., Cominelli, F., and Rivera-Nieves, J. 2008. A CD8+/CD103high T cell subset regulates TNF-mediated chronic murine ileitis. J Immunol 180:2573–2580. Wang, J., Jiang, S., Shi, H., Lin, Y., Wang, J., and Wang, X. 2008. Prolongation of corneal xenotransplant survival by T-cell vaccinationinduced T-regulatory cells. Xenotransplantation 15:164–173.

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48. Liu, Y., Chen, N., Chen, G., and You, P. 2007. The protective effect of CD8+CD28− T suppressor cells on the acute rejection responses in rat liver transplantation. Transplant Proc 39:3396–3403. 49. Zhou, J., Carr, R.I., Liwski, R.S., Stadnyk, A.W., and Lee, T.D. 2001. Oral exposure to alloantigen generates intragraft CD8+ regulatory cells. J Immunol 167:107–113. 50. Xystrakis, E., Dejean, A.S., Bernard, I., Druet, P., Liblau, R., Gonzalez-Dunia, D., and Saoudi, A. 2004. Identification of a novel natural regulatory CD8 T-cell subset and analysis of its mechanism of regulation. Blood 104:3294–3301. 51. Han, G., Shao, H., Peng, Y., Zhang, P., Ke, Y., Kaplan, H.J., and Sun, D. 2007. Suppressor role of rat CD8+CD45RClow T cells in experimental autoimmune uveitis (EAU). J Neuroimmunol 183:81–88. 52. Oluwole, S.F., Ng, A.K., Reemtsma, K., and Hardy, M.A. 1989. The mechanism of the induction of immunologic unresponsiveness to rat cardiac allografts by recipient pretreatment with donor lymphocyte subsets. Transplantation 48:281–288. 53. Kataoka, M., Margenthaler, J.A., Ku, G., and Flye, M.W. 2003. Development of infectious tolerance after donor-specific transfusion and rat heart transplantation. J Immunol 171:204–211. 54. Miller, A., Lider, O., and Weiner, H.L. 1991. Antigen-driven bystander suppression after oral administration of antigens. J Exp Med 174:791–798.

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Chapter 6 Regulatory CD4– CD8– Double Negative T Cells* Edward Y. Kim, Stephen C. Juvet, and Li Zhang Abstract Peripheral abTCR+CD3+CD4–CD8– NK1.1/CD56– double-negative (DN) Treg cells are a relatively rare subset of regulatory cells found in both humans and mice, typically comprising less than 5% of the total peripheral T-cell pool. Numerous studies have shown that DN Tregs can inhibit CD4+ and CD8+ T-cell responses in vitro and in vivo using a variety of model systems [Zhang et al., Nature Medicine 6:782, 2000; Young et al., Blood 100:3408, 2002; Ford et al., Experimental Medicine 196:261, 2002; Young et al., Journal of Immunology 171:134, 2003; Ford et al., European Journal of Immunology 37:2234, 2007; Zhang et al., Blood 109:4071, 2007; Fischer et al., Blood 105:2828, 2005]. This chapter describes published methods for the phenotypic identification of DN Tregs, their isolation from secondary lymphoid organs of mice or human peripheral blood, activation and expansion, and assays for their ability to suppress T-cell proliferation, induce apoptosis, and promote tolerance to allografts in vivo. Key words: Double-negative T cells, Regulatory cells, Transplant tolerance, Autoimmunity, Isolation, Activation/expansion, Suppression, Apoptosis

1. Introduction Numerous Treg subsets have been identified that possess distinct phenotypes in various disease models. Among these subsets, peripheral ab-TCR+CD3+CD4–CD8– NK1.1– double-negative (DN) Treg cells have been shown to inhibit a range of immune responses mediated by CD4+ and CD8+ T cells in murine models, using a number of mechanisms that may relate, in part, to the mode of stimulation employed for their activation/expansion. We identified a novel regulatory mechanism in an allo-specific TCR transgenic murine model (2C TCR), wherein DN Treg cells activated in vivo via donor-lymphocyte induced tolerance to *Kim and Juvet contributed equally to this chapter. Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_6, © Springer Science+Business Media, LLC 2011

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allogeneic skin grafts via the acquisition of allo-MHC to mediate antigen-specific, Fas/FasL-dependent cytolysis of target allospecific T cells (1). We also showed in a non-TCR transgenic murine model that DN Treg cells from C57BL/6 (B6).Faslpr/lpr mice, which possess an abnormal accumulation of DN T cells, can dose dependently kill syngeneic CD8+ and CD4+ T cells from syngeneic B6.Faswild-type mice through Fas/Fas ligand interactions in vitro and in vivo (2). 2C DN Tregs were also able to inhibit graft-versus-host disease (GVHD) mediated by 2C CD8+ T cells in a single MHC class I mismatch model of bone marrow transplantation (13). In another model system, islet peptide-specific transgenic DN Tregs were able to inhibit autoimmune diabetes development (14). More recently, it has been demonstrated that peripheral DN Treg cells capable of inducing apoptosis and suppressing the proliferation of antigen-specific CD8+ T cells are also found in humans (3). Hence, a variety of model systems have been used to study DN Treg cells. Due to the lack of a unique cell surface marker on murine and human DN Treg cells, isolation methods have relied on the depletion of non-DN Treg cells (i.e., CD4+, CD8+, NK+/ DX5+/CD56+, CD19+, gd TCR+, etc.) using magnetic bead-cell separation technology or fluorescence-activated cell sorting. Due to their relative low abundance, more stringent methods of isolation are necessary. DN Treg cells can be activated and/or expanded using stimulation relevant to the model system of study; for instance, allogeneic antigen-presenting cells, donor-lymphocyte infusion, or peptide-pulsed antigen-presenting cells have all been employed. These methods can thereby activate alloreactive or antigen-specific DN Treg cells, which can be functionally assayed for their suppression of CD4+ or CD8+ T cell proliferation, induction of apoptosis, or promotion of allograft tolerance in vivo using well-established assays.

2. Materials 2.1. Equipment

1. Laminar flow hood for tissue culture. 2. Atmosphere-controlled incubator set to 37°C with 5% CO2. 3. Microscope and hemocytometer. 4. Centrifuge. 5. Analytical flow cytometer. 6. Fluorescence-activated cell sorter (required for Subheading 3.3). 7. Supply of 51Cr and g emission counter (e.g., TopCount, Packard Instruments) for cytotoxicity assay (required for Subheading 3.7). 8. Institutionally approved anesthetic drugs, surgical instruments, and facilities (for Subheading 3.8).

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1. Mouse spleens and/or lymph nodes. 2. Cold erythroycte lysis buffer: 0.15 M NH4Cl, 10 mM KHCO3, 0.1 mM EDTA. 3. Wire mesh or other suitable filter. 4. 10 mL syringe and plunger. 5. Petri dish.

2.3. Isolation of Lymphocytes from Human Peripheral Blood

1. Blood donors. 2. Venipuncture equipment and blood collection tubes. 3. (Optional) leukapheresis system for enrichment of PBMC. 4. Ficoll or other suitable density gradient for lymphocyte isolation.

2.4. Monoclonal Antibodies for Purification and Flow Cytometry of DN T Cells

1. Fluorescence-activated cell sorting (FACS) buffer: phosphatebuffered saline (PBS), pH 7.2 with 2% fetal bovine serum (FBS). 2. For murine DN T cells, the following fluorochrome-conjugated antibodies are used: anti-TCR mAb (a-chain, clone: H57597, eBioscience), anti-CD8 mAb (a-chain, clone: 53.67, eBioscience), anti-CD4 mAb (clone: GK1.5, eBioscience), and anti-NK1.1 mAB (clone: PK146, eBioscience). 3. For human DN T cells, the following fluorochrome-conjugated antibodies are used: anti-TCRab mAb, anti-CD8 mAb, antiCD4 mAb, and anti-CD56 mAb. 4. For depletion of non-DN T-cell subsets using Dynabeads (Invitrogen), purified (i.e., nonfluorochrome-conjugated) monoclonal antibodies (mAb) are required. 5. For isolation of murine dendritic cell precursors, fluorochrome-conjugated anti-Gr1, anti-Ter119, and anti-B220 antibodies are required (eBioscience). 6. For activation of T cells, plate-bound anti-CD3 antibodies (clone 145-2C11) and anti-CD28 antibodies (clone 37.51) can be employed.

2.5. Magnetic Cell Sorting

1. IgG-coated Dynabeads. 2. Dynal magnet.

2.5.1. Using Dynal Dynabeads (Invitrogen) 2.5.2. Using MACS Columns and Microbeads (Miltenyi)

1. MACS microbeads (e.g., anti-PE, anti-biotin, anti-FITC and/ or beads specific for a particular antigen, e.g., anti-CD4). 2. LD columns (for depletion); LS or MS columns (for positive selection). 3. MiniMACS (for MS columns) or MidiMACS (for LD or LS columns) magnets.

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4. MACS support stand for magnets. 5. MACS Buffer: 0.5% bovine serum albumin (BSA) in PBS with 2 mM EDTA. 2.6. Expansion/ Activation of Regulatory DNT Cells

1. Culture medium: for DN T-cell cultures, a-MEM supplemented with 10% FBS and b-mercaptoethanol (50 mM, Gibco); for dendritic cell cultures, RPMI1640 supplemented with 10% FBS and b-mercaptoethanol (50 mM, Gibco). 2. Tissue culture plates: 96-well round-bottom tissue culture plate (Corning), 24-well flat-bottom tissue culture plate (Corning). 3. Exogenous cytokines: recombinant human IL-2 (R&D Systems), recombinant human IL-15 (R&D Systems), and recombinant IL-4 (R&D Systems) stored in aliquots at –80°C, to be added to cultures as required. 4. Generation of allogeneic mature bone marrow-derived dendritic cells (mDCs) in vitro: recombinant GM-CSF (R&D Systems) and LPS (Escherichia coli 0127:B8; Sigma-Aldrich). Teflon cell scrapers (Corning). PE-conjugated antibodies: anti-mouse Gr1, anti-mouse Ter119, and anti-mouse B220 antibodies (eBioscience). FITC or PE-conjugated antibodies to CD40 and CD86 (eBioscience).

2.7. CFSE Labeling of Responder T Cells

1. Carboxyfluorescein succinimdyl ester (CFSE, Invitrogen) reconstituted as a concentrated 10–100 mM stock in the highly purified dimethyl sulfoxide (DMSO) provided. 2. PBS, pH 7.2. 3. Fetal Bovine Serum (FBS). 4. Complete a-MEM: a-MEM supplemented with 10% FBS and 50 mM b-mercaptoethanol.

2.8. Mouse Skin Allografting

1. Sterile scalpel blades (#11). 2. Clear spray bandage (e.g., NewSkin). 3. Thin glass tubing.

3. Methods 3.1. Phenotype/ Identification of DNT Cells 3.1.1. Murine DNT Cells

1. Disrupt mouse spleens and lymph nodes in a-MEM in a sterile filter or wire mesh placed in a petri dish, and crush the remnants using the plunger of a syringe. Rinse the filter or mesh well with the a-MEM to generate a single cell suspension. 2. Pellet the cells by centrifugation and lyse erythrocytes in cold ammonium chloride lysis buffer. Incubate on ice for 5 min, pellet cells, and wash twice in complete a-MEM.

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bTCR Fig. 1. Phenotypic identification of DN Treg cells. Single cell suspensions were prepared from the spleen of C57B/6 (B6) and B6.Fas lpr/lpr mice and stained with PE-conjugated anti-CD4, anti-CD8 and anti-NK1.1 monoclonal antibodies, and FITC-conjugated antibTCR monoclonal antibody. The rectangle gate within each dot plot corresponds to DN Treg cells, and their percentage within the total spleen is shown.

3. Lymph node and spleen cells (1 × 10 6) are stained with fluorochrome-conjugated antibodies against b-TCR, CD4, CD8, and NK1.1 (or DX5) at 4°C in FACS buffer at recommended dilutions per manufacturer’s protocol. After 20 min, the stained cells are washed three times in FACS buffer and then resuspended in 400 mL of FACS buffer for analysis by flow cytometry. 4. An example of the flow cytometry results from the spleens of B6 and B6.Faslpr/lpr mice is shown in Fig. 1. 3.1.2. Human DNT Cells

1. PBMCs are isolated from leukapheresis products by Ficoll density gradient centrifugation (obtained from healthy donors with informed consent). 2. PBMCs are stained with fluorochrome-conjugated antibodies against ab TCR, CD4, CD8, and CD56 at 4°C in FACS buffer at recommended dilutions per manufacturer’s protocol. After 20 min, the stained cells are washed three times in FACS buffer and then resuspended in 400 mL of FACS buffer for analysis by flow cytometry.

3.2. Isolation of Murine DN T Cells Using Magnetic Cell Separation

1. Obtain a single cell suspension of mouse spleen and lymph node cells. 2. Pellet the cells by centrifugation and resuspend the pellet in cold ammonium chloride erythrocyte lysis buffer. Once the cells are resuspended, incubate the tube on ice for 5 min. 3. Pellet the cells and wash twice in a-MEM. The cell suspension can be passed through a 40 mM filter at this stage to remove debris.

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4. Count cells in a hemocytometer. Pellet the cells and resuspend in PBS supplemented with 0.2% BSA. Add the following biotinylated or fluorochrome-conjugated antibodies: anti-CD4, anti-CD8, anti-NK1.1, or anti-DX5, and anti-CD19 at appropriate dilutions. Purity of the final DN T-cell population can be enhanced by adding anti-CD11b, anti-CD11c, antiTer119, anti-Gr1, and anti-gamma delta TCR antibodies. Incubate at 4°C for 15 min. 5. Wash cells twice in MACS buffer. 6. Pellet cells and remove supernatant completely. Now, add MACS buffer and anti-PE, anti-FITC, or anti-biotin-conjugated microbeads (Miltenyi) in the appropriate ratio to total volume according to the manufacturer’s instructions. Incubate at 4°C for at least 20 min. 7. Wash cells in MACS buffer, pellet, and resuspend in at least 500 uL of MACS buffer. Remove any debris at this stage; this can be accomplished using a sterile 40 mM filter. Take a small aliquot of the sample for flow cytometric analysis. 8. Degas MACS buffer for >5 min. 9. Prepare an LD column (Miltenyi) by inserting it into a MidiMACS magnet and equilibrate with 2 mL of degassed MACS buffer. 10. Place a fresh tube under the column labeled “negative fraction” and add the cell suspension to the column. Allow the suspension to run into the column. 11. Wash the column at least twice with 1 mL of MACS buffer while collecting effluent in the same tube. More than two washes may be required to optimize DN T-cell yield depending on the initial cell number. 12. Now remove the column from the magnet and place it in a fresh tube marked “positive fraction.” Add 5 mL MACS buffer to the column and force it through using the supplied plunger. 13. Take aliquots from the positive and negative fractions for flow cytometry. Stain both of these as well as the precolumn sample with an anti-TCRb antibody conjugated to a different fluorochrome than the one used for depletion (e.g., if PE antibodies were used, stain with FITC-conjugated anti-TCRb antibodies). Incubate at 4°C in the dark for 15 min, and wash twice in PBS with 0.2% BSA. 14. While staining, pellet the cells in the negative fraction and resuspend in complete a-MEM. Keep cells on ice until ready for use for in vitro or in vivo experiments. 15. Determine DN T-cell purity by flow cytometry which is the percentage of TCRb+CD4–CD8–DX5– cells in the negative fraction. See Note 1.

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16. Determine the total number of DN T cells present in the negative fraction by counting viable cells in a hemocytometer and multiplying the result by the fraction of TCRb+CD4– CD8–DX5– cells as determined in step 15. In our hands, using B6.Faslpr/lpr mice, this protocol results in 70–95% TCRb+ cells, of which >97% are CD4–CD8–DX5–. Owing to the rarity of DN Tregs in normal B6 mice, yield and purity are generally much lower using this protocol. See Note 2. 3.3. Isolation of Human DNT Cells Using FluorescenceActivated Cell Sorting

1. This method was described by Fischer et al. (3). 2. Obtain peripheral blood and centrifuge over a Ficoll gradient as recommended by the manufacturer. 3. Deplete CD4+, CD8+, CD14+, CD19+, and CD56+ cells as follows. Incubate Ficoll-enriched PBMC with purified specific antibodies to these antigens. Wash cells in PBS containing 0.2% BSA and incubate in the same buffer with IgG-coated Dynabeads (Invitrogen) according to the manufacturer’s protocol. Incubate on a rotary mixer at 4°C for 30 min. 4. Apply tube to a magnet and remove nonbound cells in the buffer and place in a fresh tube. Pass over the magnet a second time if any beads remain in the buffer. 5. Stain cells with anti-TCRb, anti-CD4, and anti-CD8 antibodies conjugated to different fluorochromes, wash in 0.2% BSA-PBS, and sort TCRb+CD4–CD8– cells using a fluorescence-activated cell sorter. 6. Count cells and culture as needed; one may now proceed to in vitro activation (see Subheading 3.5).

3.4. Activation and Expansion of Regulatory Murine Allo-Reactive DNT Cells 3.4.1. Generation of Allo-Reactive Murine DNT Cells In Vitro

Allo-reactive murine DN T cells can be generated using different preparations of alloantigen-presenting cells as stimulators. 1. Alloreactive H-2d-specific DN T cells are generated by incubating the purified cells with irradiated (20 Gy) BALB/c or [B6xBALB/c]F1 splenocytes (2, 4); this method can also be used for other alloantigens, e.g., B6bm1 (5). 2. After isolation, seed 1 × 105 DN T cells and 1 × 105 irradiated allogeneic splenocytes per well in a 96-well plate. 3. Add rhIL-2 (50 U/mL) and rIL-4 (30 U/mL) to the cultures.

3.4.1.1. Stimulators: Irradiated Allogeneic Splenocytes

4. Incubate the cultures for 5–6 days at 37°C in a 5% CO2 environment.

3.4.1.2. Stimulators: Allogeneic Mature Bone Marrow-Derived Dendritic Cells

Alloreactive DN T cells can be generated via conversion from CD4+ T cells that were expanded using allogeneic mature bone marrow-derived dendritic cells (mDCs) (6). The generation of mDCs utilizes a modified version of the Lutz method (7).

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1. Harvest bone marrow cells from femurs and tibiae of allogeneic (e.g., BALB/c) mice by dissecting out these bones, cutting off the ends with sterile scissors, and flushing the marrow cavities into a sterile tube. The latter can be accomplished using a 5 mL syringe filled with RPMI1640 connected to a 23G needle that is inserted into the opening at the end of the bone. 2. Remove erythrocytes using erythrocyte lysis buffer. 3. Incubate bone marrow cells with PE-conjugated antibodies: anti-mouse Gr1, anti-mouse Ter119, and anti-mouse B220 antibodies for 20 min at 4°C, wash in MACS buffer, and then incubate with anti-PE microbeads as per the manufacturer’s protocol (Miltenyi Biotec, Auburn CA). 4. Deplete Gr1+ Ter119+ B220+ cells using a MACS LD column as per the manufacturer’s protocol (Miltenyi Biotec, CA). Typically this results in >90% Gr1– Ter119– B220– cells. 5. Seed 2 × 106 purified Gr1– Ter119– B220– bone marrow cells in 100 mm tissue culture dishes in RPMI with 20 ng/mL rmGM-CSF. Change media and add fresh rmGM-CSF (20 ng/mL) every 2–3 days. On day 5, add 0.1 mg/ml LPS (E. coli 0127:B8; Sigma-Aldrich) to induce maturation of DCs. 6. Mature DCs (mDCs) are harvested on day 6 for use as stimulators. Collect mDCs from the plates, wash with FACS buffer, and incubate with fluorchrome-conjugated anti-CD40 and anti-CD86 antibodies (eBioscience) at the appropriate dilutions in FACS buffer for 20 min at 4°C. 7. Wash twice in MACS buffer and incubate with anti-fluorochromeconjugated microbeads (Miltenyi) for 20 min at 4°C as per the manufacturer’s protocol. Wash in MACS buffer. 8. Enrich CD40hi CD86hi mDCs by positive selection through an MS column (Miltenyi) as per the manufacturer’s protocol. Additional washes may increase the cell yield. 9. Count the purified CD40hi CD86hi mDCs and determine their purity by flow cytometry. 10. Seed 1 × 105 purified CD4+ T cells per well and 0.25 × 105 of the purified mDCs per well in 96-well, round-bottom plates. 11. Add rhIL-2 and rmIL-15 to the culture at concentrations of 5 ng/mL and 500 ng/mL, respectively. Incubate cultures at 37°C in a 5% CO2 environment. 12. Harvest cells after 5 days of culture and stain with flurochromeconjugated antibodies to bTCR, CD4, CD8, and either NK1.1 or DX5. DN T cells can now be sorted for subsequent functional assays.

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Donor-lymphocyte infusion (DLI) has been shown to activate regulatory murine DN T cells in vivo, which are capable of promoting allograft tolerance (8). 1. (2C × dm2)F1 or (B6 × dm2)F1 mice (both Ld–) can be used as recipients. Intravenously inject mice with 4 × 107 splenocytes in PBS from sex-matched Ld+ (B6 × BALB/c)F1 mice, as described previously (1, 8). Depending on the experiment, it may be useful to leave additional (B6xdm2)F1 or (2Cxdm2)F1 mice uninjected as controls. 2. B6.Faslpr/lpr mice can also be used as recipients and given intravenous infusions of a mixture of 4 × 107 spleen and lymph node cells from allogeneic mice, e.g., B6.bm1 or B6.bm12 mice (2). 3. Harvest spleen and lymph node cells from the DLI-treated B6.Faslpr/lpr mice on d7 post-DLI for subsequent isolation and functional assays of activated DN Treg cells.

3.5. Activation and Expansion of Regulatory Human DN T Cells

Human DN T cells can be activated and expanded in vitro using allogeneic PBMCs or DCs (3). See Note 3. 1. Isolate PBMCs from leukapheresis products (obtained from healthy donors with informed consent) by Ficoll density gradient centrifugation and irradiate them (30 Gy) for use as stimulators. 2. DCs are generated from leukapharesis products, as previously described (9). Monocytes are enriched by countercurrent elultriation. 3. After enrichment, culture monocytes in M’ medium supplemented with 2% autologous serum, rGM-CSF, and rIL-4. 4. On day 6, add fresh medium containing rGM-CSF, rIL-4, TNF-a, IL-6, IL-1b, and PGE2 to the culture. 5. After an additional 48 h, harvest nonadherent cells for use as stimulators. 6. Seed 1 × 10 5 freshly purified DN T cells per well with 1 × 105 irradiated allogeneic PBMCs or DCs per well in 96-well round-bottom plates in the presence of rhIL-2 (100 U/mL). 7. Incubate the culture at 37°C in a 5% CO2 environment for 4 days.

3.6. Assay for Murine DN Treg-Mediated Suppression of T Cell Proliferation Using CFSE

1. Prepare either congenic CD90.1 or CD45.1 CD4+ or CD8+ T cells using microbeads conjugated to anti-CD4 or antiCD8 antibodies, followed by positive selection on MS or LS columns in a manner similar to the protocol for DN T-cell isolation described above. Alternatively, “untouched” CD4+ or CD8+ T cells can be purified by depleting non-CD4+ or

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non-CD8+ cells from the cell suspension using appropriate biotinylated or fluorochrome-conjugated antibodies and antifluorochrome- or anti-biotin-conjugated beads, followed by depletion of these populations using an LD column. 2. Establish the purity of the responder CD4+ or CD8+ T cells using flow cytometry. 3. Wash the responder cells in PBS twice and adjust the cell concentration to 1 × 107 cells/mL. 4. Prepare a stock solution of CFSE (Invitrogen) at a concentration of 1 mM. Add 1 mL of the stock solution to each 1 mL of cell suspension (to give a working concentration of 1 mM) and vortex immediately. Incubate at room temperature in the dark for 8 min. Add an equal volume of FBS to halt the labeling process. 5. Wash at least three times in complete a-MEM, take an aliquot, and confirm successful CFSE labeling by flow cytometry (high FL1 signal). Obtain the viable cell count. 6. Prepare either anti-CD3 mAb (clone 145-2C11)-coated plates or irradiated allogeneic spleen cells to stimulate responder cells. 7. In a 96 well plate, add 5 × 104–1 × 105 CFSE-labeled responder T cells and rIL-2 (50 U/mL) per well. If plates are coated with anti-CD3 mAb, add soluble anti-CD28 mAb at the start of culture. If plates are not coated with anti-CD3 mAb, add an equal number of irradiated allogeneic splenocytes as well. Add DN Tregs at varying ratios (e.g., 5:1, 2.5:1, 1.25:1, 0.6:1). 8. Include control wells with no DN Tregs (0:1) and wells with no stimulator cells or anti-CD3 antibodies (responder T cells alone). 9. Incubate for 3–5 days at 37°C in a 5% CO2 atmosphere. 10. Using different fluorochrome-conjugated (non-FITC) antibodies, stain cells for CD90.1 (or CD45.1), CD4 (or CD8) and, if desired, propidium iodide. Analyze proliferation, as measured by CFSE dilution, in the responder T-cell population by flow cytometry (see Fig. 2). See Note 4. 3.7. Assay of Cytotoxic Function of DNT Cells by 51Cr Release

1. Prepare CD4+ or CD8+ T cells and activate using plate-bound anti-CD3 and soluble anti-CD28 antibodies (3 days) or irradiated splenocytes (5 days). 2. Simultaneously, activate DN T cells in a similar fashion. 3. Pellet 1 × 106 activated responder T cells and add 1 mCi 51Cr. Incubate for 45 min at 37°C in a 5% CO2 atmosphere. 4. While target cells are being labeled, count viable DN T cells and resuspend in complete a-MEM at 1 × 106 cells/mL.

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51

88

CFSE DN:CD4 Ratio

5:1

0.6:1

Alloantigen

+

+

83

48

CFSE DN:CD4 Ratio

0:1

0:1

Alloantigen

−

+

Fig. 2. Suppression of alloantigen-induced proliferation of B6.Thy1.1 CD4+ T cells by syngeneic B6.Faslpr/lpr DN Treg cells. Responder B6.Thy1.1 CD4+ T cells were labeled with 1 mM CFSE and co-cultured in a 96-well plate (105 per well) with irradiated CB6F1 (Thy1.2+) splenocytes (105 per well) and 50 U/mL rhIL-2. B6.Faslpr/lpr DN T cells (Thy1.2+) were added to cultures at the indicated ratios, and CFSE dilution in the live, Thy1.1+ CD4+ T cell population was determined by flow cytometry after 5 days. Numbers in the top left corner of each plot indicate the percentage of undivided (CFSEhi) Thy1.1+ CD4+ T cells.

Plate 200 mL of this suspension in each of three wells of a 96 well V-bottom plate. Designate these wells “10:1.” 5. Fill three additional wells with 100 mL complete a-MEM for each of the following ratios: 5:1, 2.5:1, 1.25:1, 0.6:1, 0.3:1, and “0:1/spontaneous.” Serially dilute DN T cells by taking 100 mL from each of the three 10:1 wells and pipetting up and down in the 5:1, 2.5:1, 1.25:1, 0.6:1, and 0.3:1 wells, taking care not to add any DN T cells to the 0:1/spontaneous wells. 6. Wash activated responder T cells three times in complete a-MEM. Count viable cells.

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7. Adjust the concentration of responder T cells to 1 × 105 cells/ mL, and add 100 U/mL rIL-2, and 5 ´ 105/mL fresh irradiated allogeneic splenocytes. 8. Add 100 mL of the 51Cr-labeled T-cell suspension to each of the wells containing DN T cells, and to the 0:1 (spontaneousrelease) wells. 9. Add 100 mL of the suspension to each of three additional wells, labeled “maximum release,” and immediately add 100 uL of a dilute bleach solution to each of these wells. 10. Incubate the plate at 37°C in a 5% CO2 atmosphere for up to 24 h. 11. Transfer 80 mL of the supernatant from each well onto a lumiscence plate (e.g., LumaPlate), ensuring that only the cell-free supernatant is transferred. 12. Read the plate in an appropriate device designed to quantify g-ray emissions (e.g., TopCount). 13. Determine the mean triplicate results.

51

Cr release for each ratio from the

14. Calculate the percent specific lysis for each ratio as follows: % Specific lysis = [(cpmratio−cpmspontaneous)/(cpmmax release− cpmspontaneous)] × 100% 15. Plot the percent specific lysis vs. DN:target ratio. See Note 4. 3.8. Inhibition of Skin Graft Rejection by DNT Cells

1. Prepare erythrocyte-depleted splenocytes from mice syngeneic to the graft donors. 2. Inject each allogeneic or xenogeneic graft recipient with 4 × 107 of the splenocytes, and leave additional recipient mice uninjected to serve as controls. House the mice in an animal facility overnight. 3. Anesthetize the mice. Using sterile technique, remove a 1 × 0.5cm2 piece of skin from the tail of each donor mouse and replace it with a comparably sized piece of skin (including dermis and epidermis) from a Ld+ (B6xBALB/c)F1 mouse. Cover each graft with clear spray bandage (e.g., New Skin®) followed by a loosely-fitting piece of glass tubing. House the mice in an animal facility. 4. On a daily basis, monitor grafts visually for signs of necrosis. Score a graft as rejected when >90% necrotic. 5. Plot percentage of surviving grafts vs. time and perform survival analysis using the log rank test. 6. Isolate DN T cells from tolerant and rejecting mice and adoptively transfer to fresh recipient mice one day prior to skin grafting. Assess graft survival as described in step 4. See Note 5.

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4. Notes 1. The methods of purification described here are in common use in our laboratory. Other methods of purification can be used; for example, a more limited depletion of CD4+, CD8+, and NK1.1+ populations followed by positive selection of Thy1.2+ cells using microbeads has been described (10). T cells can be enriched from mouse spleen and lymph node cell suspensions by passage over nylon wool; then, elimination of CD4+ and CD8+ cells using specific antibodies and complement can be performed (11). Regardless of which method is selected, the purity of the DN T-cell population must always be confirmed by flow cytometry prior to use in experiments. 2. Since DN T cells comprise a small population, it is sometimes necessary to perform additional rounds of purification to remove other contaminating populations and/or increase yield. In the case of MACS separation, insufficient purity can be improved by passing the negative fraction over a second column. In contrast, if increased yield is required, the positive fraction can be passed over a second column, assuming that it still contains DN T cells. 3. Anti-CD3/CD28 mAb-mediated stimulation can also be used to activate cytokine production by human DN T cells (3). 4. There is likely to be functional and phenotypic heterogeneity within the DN T-cell population that has not yet been fully characterized. As such, some variability in DN T-cell regulatory function can be observed depending on the preparation. 5. Various allo- and xenograft models in which DN T cells promote tolerance have been described. In general, the more profound the donor–recipient mismatch, the greater the difficulty in establishing tolerance. For instance, in the (2Cxdm2) F1 model, in which only a single MHC class I mismatch is present, 100% long-term skin allograft acceptance can be obtained with DLI-mediated activation of recipient DN T cells alone (1). In contrast, rejection of rat-to-mouse cardiac xenografts was prevented only when CD4+ T cells were also depleted (12).

Acknowledgments The authors would like to thank Ramesh Vanama and Betty Joe for technical assistance. E.Y.K and S.C.J. are recipients of Terry Fox Foundation Research Fellowships from the National Cancer

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Institute of Canada. S.C.J. was also supported by the ClinicianScientist Training Program, Department of Medicine, University of Toronto. This work was supported by Canadian Institutes of Health Research grant # 14431 and CCS grant # 17157 to LZ. References 1. Zhang ZX, Yang L, Young KJ, DuTemple B, Zhang L. (2000) Identification of a previously unknown antigen-specific regulatory T cell and its mechanism of suppression. Nature Medicine 6 :782. 2. Ford MS, Young KJ, Zhang Z, Ohashi PS, Zhang L. (2002) The immune regulatory function of lymphoproliferative double negative T cells in vitro and in vivo. Journal of Experimental Medicine 196 :261. 3. Fischer K, Voelkl S, Heymann J, Przybylski GK, Mondal K, Laumer M, Kunz-Schughart L, Schmidt CA, Andreesen R, Mackensen A. (2005) Isolation and characterization of human antigen-specific TCR alpha beta(+) CD4(–) CD8(–) double-negative regulatory T cells. Blood 105:2828. 4. Ford McIntyre MS, Young KJ, Gao J, Joe B, Zhang L. (2008) Cutting edge: in vivo trogocytosis as a mechanism of double negative regulatory T cell-mediated antigen-specific suppression. Journal of Immunology 181:2271. 5. Ford MS, Zhang ZX, Chen W, Zhang L. (2006) Double-negative T regulatory cells can develop outside the thymus and do not mature from CD8+ T cell precursors. Journal of Immunology 177:2803. 6. Zhang D, Yang W, Degauque N, Tian Y, Mikita A, Zheng XX. (2007) New differentiation pathway for double-negative regulatory T cells that regulates the magnitude of immune responses. Blood 109 :4071. 7. Lutz MB, Kukutsch N, Ogilvie AL, Rössner S, Koch F, Romani N, Schuler G. (1999) An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. Journal of Immunological Methods 223:77.

8. Young KJ, Yang L, Phillips MJ, Zhang L. (2002) Donor-lymphocyte infusion induces transplantation tolerance by activating systemic and graft-infiltrating double-negative regulatory T cells. Blood 100 :3408. 9. Meidenbauer N, Marienhagen J, Laumer M, Vogl S, Heymann J, Andreesen R, Mackensen A. (2003) Survival and tumor localization of adoptively transferred Melan-A-specific T cells in melanoma patients. Journal of Immunology 170 :2161. 10. Zhang ZX, Ma Y, Wang H, Arp J, Jiang J, Huang X, He KM, Garcia B, Madrenas J, Zhong R. (2006) Double-negative T cells, activated by xenoantigen, lyse autologous B and T cells using a perforin/granzyme-dependent, Fas-Fas ligand-independent pathway. Journal of Immunology 177:6920. 11. Zhang ZX, Stanford W, Zhang L. (2002) Ly-6A is critical for the function of double negative regulatory T cells. European Journal of Immunology 32:1584. 12. Chen W, Ford M.S., Young K.J., Zhang L. (2003) Infusion of in vitro-generated DN T regulatory cells induces permanent cardiac allograft survival in mice. Transplantation Proceedings 35:2479. 13. Young KJ, DuTemple B, Phillips MJ, Zhang L. (2003) Inhibition of graftversus-host disease by double-negative regulatory T cells. Journal of Immunology 171:134. 14. Ford MS, Chen W, Wong S, Li C, Vanama R, Elford AR, Asa SL, Ohashi PS, Zhang L. (2007) Peptide-activated double-negative T cells can prevent autoimmune type-1 diabetes development. European Journal of Immunology 37:2234.

Chapter 7 Identifying Regulatory B Cells (B10 Cells) that Produce IL-10 in Mice Takashi Matsushita and Thomas F. Tedder Abstract Regulatory B cells that produce IL-10 are now recognized as an important component of the immune system. We have identified a rare antigen-specific regulatory B-cell subset with a unique CD1dhiCD5+CD19hi phenotype in the spleens of wild-type mice. We call these cells B10 cells because they are responsible for most B cell IL-10 production, they appear to only produce IL-10 after 5 h of in vitro stimulation, and to distinguish them from other potential regulatory B cell subsets. B10 progenitor (B10pro) cells have also been identified within the spleen CD1dhiCD5+CD19hi B-cell subset, and within other lymphoid tissues. Herein, four methods for identifying and isolating regulatory IL-10-producing B10 cells in mice are provided. The first two methods are used to identify and enumerate B10 and B10pro cells based on their cell surface phenotypes and cytoplasmic IL-10 staining. The last two methods are used to isolate viable B10 cells for adoptive transfer and functional studies. These methods should facilitate the study of B10 cells in inflammation, autoimmune disease, immune responses, and cancer therapy. Key words: B cells, Autoimmunity, Inflammation, Regulatory cells, CD5, Interleukin-10

1. Introduction Historically, B cells have been characterized as positive regulators of humoral immune responses and are uniquely distinguished by their ability to terminally differentiate into antibody (Ab)-secreting plasma cells (1). B cells can also serve as antigen (Ag)-presenting cells (APCs), with the capacity to present Ag more efficiently than dendritic cells and other APCs (2). B cell expression of CD80, CD86, OX40L, and other costimulatory molecules is also important for optimal T-cell activation (3, 4). As a result, B-cell Ag presentation and/or costimulation is required for optimal Ag-specific CD4+ T-cell activation, expansion, memory formation, and cytokine production (5–7). B cells can also positively regulate Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_7, © Springer Science+Business Media, LLC 2011

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CD8+ T-cell responses in some models (8, 9). Thus, B cells can not only produce Abs that have effector functions but also positively regulate cellular immune responses. Evidence for B-cell negative regulatory function has accumulated over the past 30 years. B-cell suppression of an immune response was first reported in 1974, where column depletion of B cells from guinea pig splenocyte preparations eliminated the ability of adoptively transferred cells to inhibit delayed-onset skin reactions (10, 11). This suggested that suppressor B cells can regulate hypersensitivity responses and T-cell function. Over the past decade, B-cell negative regulation has been demonstrated in multiple mouse models of autoimmunity (12–18). Mizoguchi et al. were the first to use the term “regulatory B cells” to designate B cells with negative regulatory properties (12). However, B-cell regulatory activities and negative regulation through the secretion of IL-10 have been attributed to multiple phenotypically diverse B-cell subsets in different mouse models. In addition, Lund et al. have identified Be1 and Be2 cell populations, both of which have the capacity to produce IL-10 (19). Although identifying regulatory B-cell subsets and their mechanisms of action has taken over 30 years, regulatory B cells are now recognized as an important component of the immune system. We have identified a relatively rare B-cell subset with negative regulatory functions that is predominantly contained within a phenotypically unique CD1dhiCD5+CD19hi subset in the spleens of naïve wild-type mice (17). This regulatory B-cell subset is Ag-specific and significantly influences T-cell activation and inflammatory responses through the secretion of IL-10 (17, 18). Given that multiple regulatory B cell subsets are likely to exist, as is now well recognized for T cells, we have specifically labeled this IL-10 competent CD1dhiCD5+CD19hi regulatory subset as B10 cells because they are responsible for most B-cell IL-10 production and they appear to only produce IL-10 (20). B10 cells are further defined as B cells that are competent to express detectable IL-10 after 5 h of in vitro stimulation with a cocktail of lipopolysaccharide (LPS), phorbol 12-myristate 13-acetate (PMA), ionomycin, and monensin (L+PIM). B10 cells are predominantly found at low frequencies within the spleen and peritoneal cavity of adult mice (Table 1). B10 progenitor (B10pro) cells have also been identified within the spleen CD1dhiCD5+CD19hi B-cell subset (20). B10pro cells are defined as those B cells that are not induced to express cytoplasmic IL-10 after L+PIM stimulation for 5 h, but that can be induced to mature into IL-10 competent B10 cells by in vitro culture with agonistic CD40 monoclonal antibody (mAb) stimulation for 48 h. B10pro cells that do not express either CD5 or CD1d are also found in other lymphoid tissues such as the peritoneal cavity, lymph nodes, and blood (Table 1). However, since IL-10 expression is the defining marker for B10 cells, it is not

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Table 1 B10 and B10pro cell distributions within tissues of C57BL/6 mice B10 cells

B10pro + B10 cells

Tissue

Percentage

Number (×10−6)

Percentage

Number (×10−6)

Blood

1.5 ± 0.1

0.029 ± 0.01

3.5 ± 0.5

0.07 ± 0.02

Bone marrow

2.8 ± 0.5

0.20 ± 0.03

3.5 ± 0.2

0.25 ± 0.03

Lymph nodes

0.7 ± 0.1

0.01 ± 0.01

1.5 ± 0.2

0.03 ± 0.01

Spleen

2.0 ± 0.3

1.18 ± 0.1

6.2 ± 0.3

3.7 ± 0.25

Peritoneal cavity

25.8 ± 3.0

0.51 ± 0.03

30.2 ± 2.0

0.62 ± 0.12

B10 cell frequencies and numbers (±SEM) in three mice between 8 and 10 weeks of age were quantified by flow cytometry as described (17, 18, 20) after culturing viable B cells with L+PIM for 5 h before CD19 and live/dead cell staining, permeabilization and intracellular staining for IL-10. B10pro cells were induced by culturing B cells with agonistic CD40 mAb for 48 h, with L+PIM added during the final 5 h of culture (20). The number of cells within the B10pro cell subset includes B10 cells. Values represent the average percentages of cytoplasmic IL-10+ cells within the CD19+ B cell compartment isolated from the indicated tissues. Numbers indicate IL-10+CD19+ B cells within each tissue, except for blood where values represent 10−6 cells/ml. Lymph nodes include pooled bilateral axial, inguinal, and brachial lymph nodes. Bone marrow leukocytes from both femurs were pooled to calculate IL-10+CD19+ cell numbers.

possible to differentiate between B10 cells and B10pro cells after 48 h in vitro cultures. Given the unique functional capabilities of B10 cells and their characteristic patterns of cell surface molecules that are expressed by multiple other B-cell subsets, it is likely that most if not all regulatory B-cell functions can be attributed to B10 cells or their progenitors, B10pro cells (20, 21). Herein, four methods for identifying and isolating regulatory B10 cells are provided.

2. Materials 2.1. Equipment

1. Fluorescence-activated cell sorter (FACS) tubes: 5 ml (12 × 75 mm) polystyrene round-bottom tubes with snap caps. 2. Pipet-Man pipettes and sterile pipette tips. 3. Sterile, disposable transfer pipettes. 4. Refrigerated centrifuge. 5. Biosafety cabinet/hood for tissue culture, and tissue culture supplies. 6. Vortexer. 7. CO2 incubator, 37°C, 5% CO2. 8. Flow cytometer and/or sorter with multiparameter cytometry capability. 9. Wet ice, to maintain cells and reagents at ~4°C.

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2.2. Immunofluorescence Staining of B10 Cells

1. Freshly prepared single-cell suspensions of isolated mouse tissue mononuclear cells. Single-cell suspensions of bone marrow (bilateral femurs), spleen, and peripheral lymph node (paired axillary and inguinal) lymphocytes are generated by gentle dissection. To isolate peritoneal cavity leukocytes, 10 ml of cold 0.2% bovine serum albumin (BSA) in Dulbecco’s phosphate-buffered saline (D-PBS) is injected by syringe into the peritoneal cavity followed by a gentle massage of the abdomen before aspiration of the fluid back into the syringe. 2. Cell culture medium: sterile RPMI 1640 containing 10% fetal bovine serum (FBS), 200 mg/ml penicillin, 200 U/ml streptomycin, 4 mM L-Glutamine, and 5 × 10−5 M 2-mercaptoethanol. The medium is filter-sterilized (0.2 mM pore size) after production and stored at 4°C. 3. D-PBS without Ca2+ and Mg2+ kept sterile, and stored at 4°C. 4. Staining Buffer: D-PBS supplemented with 2% FBS, stored at 4°C. 5. MACS buffer: D-PBS containing 0.5% BSA and 2 mM ethylenediaminetetraacetic acid (EDTA), filter-sterilized (0.2 mM pore size) after production and stored at 4°C. 6. Tissue culture plates: sterile 48-well flat-bottom culture plates. 7. PMA (Sigma) dissolved at 1 mg/ml in dimethyl sulfoxide (DMSO) and stored in single-use (50 mg/50 ml) aliquots at −20°C. 8. Ionomycin (Sigma) dissolved at 1 mg/ml in DMSO and stored in single-use aliquots (50 mg/50 ml) at −20°C. 9. LPS (Escherichia coli serotype 0111:B4, Sigma) dissolved at 10 mg/ml in sterile D-PBS, filter-sterilized (0.2 mM pore size), and stored at 4°C. 10. Monensin (eBioscience). Concentration is 2 mM. 11. LIVE⁄DEAD® Fixable Green Dead Cell Stain Kit (Invitrogen): Add 12.5 ml of DMSO to the vial of reactive dye (2 mg/ml). Mix well and visually confirm that all of the dye has dissolved (see Note 1). Store the vial at −20°C for a maximum of three thaws or 1 month post reconstitution.

2.3. Immunofluorescence Staining of B10 and B10pro Cells

1. Purified agonistic antimouse CD40 mAb (Clone HM40/3, BD Biosciences, Cat No. 553721). 2. Purified antimouse CD16/CD32 mAb (Mouse BD Fc Block™, Clone 2.4G2, BD Biosciences, Cat No. 553142). 3. Phycoerythrin (PE)-Cy5 labeled antimouse CD19 mAb (Clone eBio1D3, eBioscience, Cat No. 15-0193-82) that has been previously titrated in the lab for optimum staining.

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4. PE-labeled antimouse IL-10 mAb (Clone JES5-16E3, BioLegend, Cat No. 505008) that has been previously titrated in the lab for optimum staining. 5. PE-labeled Rat IgG2b Isotype control (Clone KLH/G2b-1-2, eBioscience, Cat No. 12-4331-82). Many commercial isotype control mAbs stain permeabilized cells, so make sure that the isotype control mAb is not reactive with cytoplasmic components of the cells being assayed. 6. BD Cytofix/Cytoperm™ cell fixation buffer (BD Biosciences). 7. BD Perm/Wash™ buffer for cell permeabilization (BD Biosciences). 8. 1.5% Formaldehyde fixing solution: 15% formalin diluted tenfold using Ca2+- and Mg2+- free PBS, and stored at room temperature for up to 1 month. 2.4. CD1d hiCD5+ B Cell Isolation

1. Antimouse CD19 mAb-coated microbeads (Miltenyi Biotec). 2. Fluorescein isothiocyanate (FITC)-labeled antimouse CD19 mAb (Clone eBio1D3, eBioscience, Cat No. 11-0193-85). 3. PE-labeled antimouse CD1d mAb (Clone 1B1, BioLegend, Cat No. 123510). 4. PE-Cy5-labeled antimouse CD5 mAb (Clone 53-7.3, BioLegend, Cat No. 100610). 5. Nylon mesh filter.

2.5. IL-10-Secreting B Cell Isolation

1. MACS Mouse IL-10 Secretion Assay – Detection Kit (Miltenyi Biotec). 2. DNase I (Roche) dissolved at 10 mg/ml in D-PBS and stored at −20°C.

3. Methods The first two methods provided are for identifying B10 and B10pro cells based on their cell surface phenotype and cytoplasmic IL-10 staining. Purified B cells or single-cell mixed mononuclear cell populations are isolated from spleen or other tissues and stimulated in vitro for 5 h with a cocktail of LPS, PMA, and ionomycin (L+PI) to induce IL-10 expression. Monensin is also added to the stimulation cocktail (L+PIM) to block intracellular protein transport and thereby retain newly generated IL-10 within the cell cytoplasm for optimal detection. The B cells are then labeled by immunofluorescence staining, with subsequent flow cytometry (FACS) analysis to determine B10 cell numbers. The second method provided is for identifying B10pro cells by inducing their maturation into IL-10 competent B10 cells after 48 h of in vitro

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stimulation through cell surface CD40 ligation using agonistic CD40 mAb. During the final 5 h of culture, matured IL-10 competent B10pro and B10 cells are induced to express cytoplasmic IL-10 by adding L+PIM to the culture medium. The B cells are then labeled by immunofluorescence staining, with subsequent flow cytometry analysis. Using this culture system to identify B10pro cells, it is not possible to determine whether individual cells are in vitro matured B10pro cells or are already mature IL-10 competent B10 cells. Therefore, it is best to determine both B10 cell and B10pro+B10 cell numbers individually using the appropriate stimulation and culture conditions in cases where it is important to determine B10 cell numbers relative to B10pro+B10 cell numbers. The last two methods are for isolating viable B10 cells. Staining for cytoplasmic IL-10 expression as described above results in cell death due to membrane permeabilization required for cytoplasmic IL-10 staining. The third method is for isolating the CD1dhiCD5+ subset of spleen B cells, which contains most spleen IL-10 competent B10 cells and B10pro cells. This technique is generally useful for isolating sufficient numbers of B10pro and B10 cells for adoptive transfer experiments. The fourth method is for isolating B cells that are secreting IL-10. This technique generates small numbers of pure IL-10+ B cells, but the B10 cell population is viable and can be used for functional, RNA, or biochemical studies. In both cases, spleens from multiple mice will need to be pooled to have sufficient numbers of cells for the envisioned studies; e.g., 1 × 106 IL-10+ B cells can be isolated from spleens of five C57BL/6 mice. 3.1. Analysis of B10 Cells

1. Resuspend single-cell mononuclear cell preparations (2 × 106 cells/ml) in prewarmed culture medium alone (as a control) or culture medium containing PMA (50 ng/ml, final concentration), ionomycin (500 ng/ml final), LPS (10 mg/ml final), and monensin (2 mM final) to induce IL-10 production. 2. Culture the cells at 37°C in a tissue culture incubator with 5% CO2 atmosphere for 5 h. 3. Harvest the media containing the cultured cells and place in an appropriately sized container on wet ice to keep the cells at ~4°C. Dispense 2 × 106 cells into one 5 ml tube for each analysis to be carried out. Fill each FACS tube up to ~4 ml with cold PBS and resuspend the cells by shaking or vortexing each tube. 4. Centrifuge the tubes (300 × g, 5 min) at 4°C to pellet the cells. Carefully remove all media from the tube by aspiration without disrupting the cell pellet and resuspend the cells in PBS. Repeat this wash procedure and then aspirate the media from the cell pellet. 5. Add 50 ml of live/dead solution (working dilution of the solution is 1:3,000–5,000) (see Notes 1–3).

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6. Add 50 ml of Fc block (working dilution of the solution is 1:50) and then vortex the tube briefly (see Note 4). 7. Incubate the cell suspension for 15 min on wet ice (~4°C) in the dark. 8. Add ice-cold PBS to each tube to resuspend the cells, pellet the cells (centrifuge 300 × g, at 4°C for 5 min) to wash, and remove supernatant fluid by aspiration. 9. Add 100 ml of PE-Cy5 CD19 mAb (general working dilution is ~1:500) and then vortex briefly (see Notes 2– 4). 10. Incubate for 20–30 min on wet ice in the dark. 11. Wash the cells twice with PBS, centrifuge (centrifuge 300 × g, at 4°C for 5 min), and aspirate the supernatant fluid from the cell pellet. 12. Thoroughly resuspend the cell pellet by vortexing and add 250 ml of BD Cytofix/Cytoperm™ solution to each tube. Resuspend the cells and incubate on wet ice in the dark for 20 min. 13. Wash the cells twice with 1× BD Perm/Wash™ buffer. Pellet the cells (centrifuge 300 × g, at 4°C for 5 min) and remove the supernatant fluid by aspiration. 14. Thoroughly resuspend the permeabilized cells in 100 ml of ice-cold BD Perm/Wash™ buffer containing PE-labeled antiIL-10 mAb (general working dilution is 1:100) or PE-labeled Isotype control Ab (see Note 5). 15. Incubate the cell mixture on wet ice for 30 min in the dark. 16. Wash the cells twice with ice-cold 1× BD Perm/Wash™ buffer and resuspend the cell pellet in 250 ml of ice-cold 1.5% formaldehyde fixative. Mix the samples thoroughly by vortexing. 17. Keep the cells on ice or refrigerated at 4°C before analyzing the stained cells by flow cytometry (see Fig. 1). It is recommended that the time between staining of the cells and flow cytometry analysis be kept to a minimum. 3.2. Analysis of B10pro and B10 Cells

1. Resuspend single-cell mononuclear cell preparations (2 × 106 cells/ml) in sterile prewarmed culture medium alone (as a control) or sterile culture medium containing CD40 mAb (1 mg/ml). 2. Culture the cells at 37°C in a tissue culture incubator with 5% CO2 atmosphere for 48 h. 3. During the final 5 h of culture, add PMA (50 ng/ml final concentration), ionomycin (500 ng/ml final), LPS (10 mg/ml final), and monensin (2 mM final) to the appropriate cell cultures to induce IL-10 expression. It may be advantageous to keep an aliquot of CD40 stimulated cells that have not been cultured with L+PIM for controls.

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Fig. 1. Isolation and purification of mouse spleen B10 cells. Splenocytes from wild type or IL-10−/− C57BL/6 mice were cultured with LPS, PMA, ionomycin, and monensin (L+PIM) for 5 h to evaluate IL-10+ B10 cell frequencies. (a) Gating strategy and flow cytometry analysis for identifying B10 cells. Gate G1 defines single cells as determined by singlet gating using a forward scatter area (FSC-A) vs. height (FSC-H) plot. The cells outside or below the gate are cell doublets, which should be excluded. Gate G2 defines the predominant lymphocyte population by forward light scatter (FSC) and side scatter (SSC) properties. It is important to not draw the gates too small as some larger B10 cells will be excluded. It is often useful to identify IL-10 producing B cell and then back-gate to optimize the FSC and SSC gates to include all B10 cells. Gate G3 defines the live cells and excludes the dead cells that are positive for Live/Dead staining. Representative cell surface CD19 and cytoplasmic IL-10 staining by viable single cells is shown in the dot-plot histograms (right panels). The percentages indicate the normal frequencies of IL-10-producing B cells within the indicated gates among total CD19+ B cells. IL-10−/− mice were used as optimal negative controls for cytoplasmic IL-10 staining (see Note 9). It is normal to see some non-B cells (CD19−) expressing IL-10 in these assays. (b) Analysis of spleen B10pro cells plus B10 cells after in vitro maturation. Splenocytes were cultured with CD40 mAb for 48 h, with L+PIM added during the final 5 h of culture. The staining, gating, and analysis strategy was the same as in (a). (c) The cell surface phenotype of IL-10+ CD19+ B cells (heavy lines) is compared with IL-10− B cells (thin lines) or control mAb staining (gray histograms) after L+PIM stimulation of cells for 5 h as in (a).

4. Harvest the activated cells, stain for cell surface CD19 and cytoplasmic IL-10, and analyze by flow cytometry as outlined in steps 3–17 in Subheading 3.1. Analysis of B10 cells. 3.3. Isolation of CD1d hiCD5+ B Cells

1. Isolate single-cell splenocyte suspensions using aseptic techniques with all procedures carried out in a biohazard hood to maintain cell sterility. Isolate CD19+ B cells from single-cell splenocyte suspensions by sterile MACS sorting or other similar techniques to enrich for B cells. (We recommend

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using CD19 or B220 mAb-based positive selection techniques since some B10 cells express the CD43 cell surface molecule and many B cell negative selection kits are designed to deplete CD43+ cells). Enriching the cell population will increase the recovery of CD1dhiCD5+ B cells and reduce the amount of cell sorting time required. 2. Using sterile 5 ml tubes with caps, resuspend 2 × 108 B cells in 2 ml of sterile staining buffer containing FITC-conjugated CD19 mAb (general working dilution is ~1:100 final concentration), PE-labeled CD1d mAb (general working dilution is ~1:250 final), and PE-Cy5-conjugated CD5 mAb (general working dilution is ~1:100 final). 3. Thoroughly resuspend the cells by vortexing and incubate the cell suspension on wet ice for 30 min in the dark. Add 2 ml of cold staining buffer to the tubes, cap the tube, and gently mix the cells. 4. Pellet the cells (centrifuge 300 × g at 4°C for 5 min) and remove the supernatant fluid by aspiration. Resuspend the cells to 107 cells/ml in sterile culture medium containing DNase I (10 mg/ml) and filter the cell suspension through a sterile nylon mesh into sterile 5 ml tubes with caps for cell sorting (see Note 6). 5. Analyze and sort cells using a FACS (see Fig. 2).

Fig. 2. Spleen IL-10-competent B10 cells are predominantly found within the CD1dhiCD5+ B cell subset. Spleen B cells were isolated by MACS, and stained for cell surface CD1d, CD5, and CD19 before flow cytometry analysis. The CD1dhiCD5+CD19+ and CD1dloCD5− CD19+ B cell populations were purified by cell sorting and cultured with L+PIM for 5 h in order to evaluate IL-10+ B10 cell frequencies. Alternatively, the sorted cell populations were cultured with agonistic CD40 mAb for 48 h, with L+PIM added during the final 5 h of culture to enumerate B10pro plus B10 cell numbers. The cultured cells were stained for viability, permeabilized, and stained for cytoplasmic IL-10 expression, with live single cells analyzed by flow cytometry as shown in Fig. 1. As shown, the frequency of CD1dhiCD5+ B cells that express IL-10+ increases significantly after 48 h of stimulation, reflecting the in vitro maturation of B10pro cells into IL-10 competent B10 cells.

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3.4. Isolation of IL-10-Secreting B Cells

1. Isolate single-cell splenocyte suspensions using aseptic techniques and enrich for CD19+ B cells as outlined in Subheading 3.3. 2. Resuspend the cells at 2 × 106 B cells in sterile culture medium containing PMA (50 ng/ml final concentration), ionomycin (500 ng/ml final), LPS (10 mg/ml final), and DNase I (20 mg/ml final). 3. Culture the cells at 37°C in a tissue culture incubator with 5% CO2 atmosphere for 4 h. 4. Transfer the cells to sterile 5 ml tubes with caps for cell sorting and wash the cells twice with cold-MACS buffer, centrifuge (300 × g at 4°C for 10 min) to pellet the cells, and aspirate the supernatant fluid completely. 5. Add 80 ml of ice-cold culture medium per 107 cells and resuspend the cell pellet. 6. Add 20 ml of IL-10 capture reagents (component of Mouse IL-10 Secretion Assay – Detection Kit) per 107 cells, mix, and incubate on wet ice for 5 min. 7. Add warm tissue culture medium containing LPS (10 mg/ml final concentration), PMA (50 ng/ml final), ionomycin (500 ng/ml final), and DNase I (20 mg/ml final) to dilute the cells to 2 × 106 cells/ml, cap the tubes, and mix gently. Incubate the cell suspension in capped tubes for 45 min at 37°C under slow continuous rotation or invert the tubes every 5 min to resuspend the cells (see Note 7). 8. Wash the cells twice with cold-MACS buffer, pellet the cells (centrifuge 300 × g at 4°C for 10 min), and aspirate the supernatant fluid completely. 9. Add 80 ml of ice-cold culture medium to the tubes for each 107 cells. 10. Add 20 ml of PE-conjugated IL-10 detection antibody per 107 cells, mix well, and incubate the cell suspension on ice for 10 min. 11. Wash the cells with ice-cold sterile MACS buffer, pellet the cells (centrifuge 300 × g at 4°C for 10 min), and aspirate the supernatant fluid completely. 12. Add 50 ml of sterile live/dead working solution to the cell pellet of each tube (general working dilution is ~1:5,000) (see Notes 2 and 3). 13. Add 50 ml of sterile Fc block (general working dilution is ~1:50 final) to each tube and then vortex briefly to resuspend the cell pellet (see Note 4). Incubate the capped tube at 4°C in the dark for 15 min.

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14. Wash the cells with ice-cold sterile PBS, pellet the cells (centrifuge 300 × g at 4°C for 5 min), and aspirate the supernatant fluid completely. 15. Add 100 ml of sterile PE-Cy5-conjugated CD19 mAb (general working dilution is ~1:500 final concentration) and then vortex the cell suspension briefly (see Note 4). 16. Incubate the cell suspension on wet ice for 10 min. 17. Resuspend the cells at 107 cells/ml in ice-cold sterile culture medium containing DNase I (10 mg/ml) and filter through nylon mesh into sterile 5 ml tubes with caps as used for cell sorting (see Note 6). 18. Analyze and isolate the IL-10+ CD19+ cells using a FACS (see Fig. 3) (see Note 8).

Fig. 3. Isolation of IL-10-secreting B cells. (a) Splenic CD19+ cells isolated from C57BL/6 mice were cultured with L+PI (LPS, PMA, ionomycin) for 5 h before staining for cell surface CD19 expression and cytokine capture to visualize IL-10-secreting cells. Gate G1 defines singlet cells. G2 defines the main lymphocyte population. G3 defines live cells and excludes Live/Dead staining dead cells (upper panels). (b) Flow cytometry analysis of viable single cells that were secreting IL-10 compared with unstimulated cells. Secreted IL-10+ and IL-10− B cells were then isolated by two rounds of cell sorting using the indicated gates, with subsequent reassessment of IL-10 secretion and CD19 expression. The percentages indicate the relative frequencies of cells within the indicated gates.

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4. Notes 1. Each lot of Live/dead cell working solution must be titrated for optimal dilution to use. 2. All media, solutions, and tubes should be sterile or as near-sterile as practical. For cell cultures, all work must be carried out in a tissue culture hood to maintain the sterility of the cell preparations. For cell surface staining, the cells should be kept cold to prevent the capping and endocytosis of cross-linked receptors. 3. Live/Dead cell working solution, Fc block, and CD19 mAb must be diluted in PBS without protein. (We do this for Live/ Dead stain, but most of us usually have carrier protein present during incubations with Fc Block and CD19 mAb.) 4. Immediately before use, diluted mAb reagents should be centrifuged in a microcentrifuge for 1 min to pellet any potential debris. 5. It is critical that the anti-IL-10 mAb is diluted in BD Perm/ Wash™ Buffer rather than staining buffer to maintain the cells in an optimally permeabilized state for intracellular staining. 6. The release of DNA from dead cells is the biggest cause of nozzle clogs during the course of cell sorting since DNA is rigid and causes cell aggregates. DNAse reduces this problem. 7. This step is important to reduce intercellular contact and thereby avoid cross-contamination of cells that do not secrete IL-10. 8. Due to the rarity of B10 cells, it is recommended that the sorted IL-10+ cells undergo two rounds of sorting to yield IL-10-secreting B cells at high purities. 9. Spleen B cells from IL-10−/− mice (B6.129P2-Il10tmlCgn/J, ref. 22) are the optimal negative control for cytoplasmic IL-10 staining and analysis since they provide background staining levels.

Acknowledgments This work was supported by grants from the National Institutes of Health CA105001, CA96547, AI56363, and AI057157. We thank Drs. Guang Yang, Regina Lin, Kathleen Candando, and David DiLillo for their input and review of this manuscript.

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References 1. LeBien, T. W., and Tedder, T. F. (2008) B-lymphocytes: How they develop and function. Blood 112, 1570–1579. 2. Liu, Y., Wu, Y., Ramarathinam, L., Guo, Y., Huszar, D., Trounstine, M., et al. (1995) Gene-targeted B-deficient mice reveal a critical role for B cells in the CD4 T cell response. Int. Immunol. 7, 1353–1362. 3. O’Neill, S. K., Cao, Y., Hamel, K. M., Doodes, P. D., Hutas, G., and Finnegan, A. (2007) Expression of CD80/86 on B cells is essential for autoreactive T cell activation and the development of arthritis. J. Immunol. 179, 5109–5116. 4. Linton, P. J., Bautista, B., Biederman, E., Bradley, E. S., Harbertson, J., Kondrack, R. M., et al. (2003) Costimulation via OX40L expressed by B cells is sufficient to determine the extent of primary CD4 cell expansion and Th2 cytokine secretion in vivo. J. Exp. Med. 197, 875–883. 5. Linton, P. J., Harbertson, J., and Bradley, L. M. (2000) A critical role for B cells in the development of memory CD4 cells. J. Immunol. 165, 5558–5565. 6. Crawford, A., Macleod, M., Schumacher, T., Corlett, L., and Gray, D. (2006) Primary T cell expansion and differentiation in vivo requires antigen presentation by B cells. J. Immunol. 176, 3498–3506. 7. Bouaziz, J. D., Yanaba, K., Venturi, G. M., Wang, Y., Tisch, R. M., Poe, J. C., et al. (2007) Therapeutic B cell depletion impairs adaptive and autoreactive CD4+ T cell activation in mice. Proc. Natl. Acad. Sci. U. S. A. 104, 20882–20887. 8. Homann, D., Tishon, A., Berger, D. P., Weigle, W. O., von Herrath, M. G., and Oldstone, M. B. (1998) Evidence for an underlying CD4 helper and CD8 T-cell defect in B-cell-deficient mice: failure to clear persistent virus infection after adoptive immunotherapy with virus-specific memory cells from mMT/mMT mice. J. Virol. 72, 9208–9216. 9. Bergmann, C. C., Ramakrishna, C., Kornacki, M., and Stohlman, S. A. (2001) Impaired T cell immunity in B cell-deficient mice following viral central nervous system infection. J. Immunol. 167, 1575–1583. 10. Katz, S. I., Parker, D., and Turk, J. L. (1974) B-cell suppression of delayed hypersensitivity reactions. Nature 251, 550–551.

11. Neta, R., and Salvin, S. B. (1974) Specific suppression of delayed hypersensitivity: the possible presence of a suppressor B cell in the regulation of delayed hypersensitivity. J. Immunol. 113, 1716–1725. 12. Mizoguchi, A., and Bhan, A. K. (2006) A case for regulatory B cells. J. Immunol. 176, 705–710. 13. Serra, P., and Santamaria, P. (2006) To ‘B’ regulated: B cells as members of the regulatory workforce. Trends Immunol. 27, 7–10. 14. Mauri, C., and Ehrenstein, M. R. (2008) The ‘short’ history of regulatory B cells. Trends Immunol. 29, 34–40. 15. Lund, F. E. (2008) Cytokine-producing B lymphocytes-key regulators of immunity. Curr. Opin. Immunol. 20, 1–7. 16. Bouaziz, J.-D., Yanaba, K., and Tedder, T. F. (2008) Regulatory B cells as inhibitors of immune responses and inflammation. Immunol. Rev. 224, 201–214. 17. Yanaba, K., Bouaziz, J. D., Haas, K. M., Poe, J. C., Fujimoto, M., and Tedder, T. F. (2008) A regulatory B cell subset with a unique CD1dhiCD5+ phenotype controls T celldependent inflammatory responses. Immunity 28, 639–650. 18. Matsushita, T., Yanaba, K., Bouaziz, J. D., Fujimoto, M., and Tedder, T. F. (2008) Regulatory B cells inhibit EAE initiation in mice while other B cells promote disease progression. J. Clin. Invest. 118, 3420–3430. 19. Harris, D. P., Haynes, L., Sayles, P. C., Duso, D. K., Eaton, S. M., Lepak, N. M., et al. (2000) Reciprocal regulation of polarized cytokine production by effector B and T cells. Nat. Immunol.1, 475–482. 20. Yanaba, K., Bouaziz, J.-D., Matsushita, T., Tasubata, T., and Tedder, T. F. (2009) The development and function of regulatory B cells expressing IL-10 (B10 cells) requires antigen receptor diversity and TLR signals. J. Immunol. 182, 7459–7472. 21. DiLillo, D. J., Matsushita, T., and Tedder, T. F. (2010) B10 and regulatory B cells balance immune responses during inflammation, autoimmunity and cancer. Ann. New York Acad. Sci. 1183, 38–57. 22. Kuhn, R., Lohler, J., Rennick, D., Rajewsky, K., and Muller, W. (1993) Interleukin-10deficient mice develop chronic enterocolitis. Cell 75, 263–274.

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Chapter 8 DCs in Immune Tolerance in Steady-State Conditions Tomohiro Fukaya, Hideaki Takagi, Honami Taya, and Katsuaki Sato Abstract Dendritic cells (DCs) are antigen-presenting cells (APCs) characterized by a unique capacity to stimulate naïve T cells and initiate primary immune responses. Recent studies suggest that DCs are also involved in the induction of immunological tolerance in peripheral tissues under steady-state conditions by maintaining the homeostasis of self-reactive CD4+Foxp3+naturally occurring thymic-derived regulatory T cells (nTregs) and de novo generation of antigen-specific CD4+Foxp3+inducible regulatory T cells (iTregs). We demonstrate here the impact of CD11+DCs on the antigen-specific differentiation of CD4+Foxp3+iTregs from CD4+Foxp3−T cells under steady-state and inflammatory conditions. CD11c+DCs promoted the transforming growth factor (TGF)-b1-mediated conversion of CD4+Foxp3−T cells into CD4+Foxp3+iTregs in vitro, while stimulation of CD11c+DCs with CpG oligodeoxynucleotide (ODN) abrogated this conversion. Furthermore, antigen-specific generation of CD4+Foxp3+iTregs required the function of CD11+DCs under steady-state conditions, whereas such conversion was severely abolished under inflammatory conditions. Thus, these results suggest the crucial role of DCs in the antigen-specific de novo conversion of CD4+Foxp3−T cells into CD4+Foxp3+iTregs under steady-state conditions, thereby leading to the establishment of peripheral immune tolerance. Key words: Dendritic cells, Tolerance, Regulatory T cells, Steady-state conditions, Inflammatory conditions

1. Introduction Dendritic cells (DCs) are essential APCs that initiate primary immune responses. DCs consist of heterogeneous subsets, including conventional DCs (cDCs) and plasmacytoid DCs (pDCs), distinguishable by surface and intracellular phenotypic markers, immunologic function, and anatomic distribution (1). Immature DCs (iDCs) serve as sentinels in peripheral tissues, recognizing the presence of invading pathogens through various patternrecognition receptors, and become mature DCs (mDCs) with

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_8, © Springer Science+Business Media, LLC 2011

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the upregulated expression of major histocompatibility complex (MHC) molecules and costimulatory molecules under inflammatory conditions (1). Consequently, mDCs move via the afferent lymphatics into the T-cell area of secondary lymphoid tissues, where they prime rare antigen-specific naive T cells for differentiation into various effector T cells (1, 2). DCs thereby play a crucial role in the link between innate and adaptive immunity (1). Conversely, iDCs could also be important for the induction of immunological tolerance under steady-state conditions in peripheral tissues (3–5), and the mechanisms involved include clonal deletion and anergy, as well as active immune suppression by CD4+ Foxp3+regulatory T cells (6–8) that encompass self-reactive naturally occurring thymic-derived Tregs (nTregs) and inducible Tregs (iTregs) generated from antigen-specific CD4+Foxp3−T cells, a function of likely importance in self-tolerance as well as immune disorders and transplant rejection. We demonstrated that antigenic peptide-pulsed splenic CD11c+DCs induced the antigen-specific conversion of CD4+Foxp3− T cells into CD4+Foxp3+iTregs in the presence of transforming growth factor (TGF)-b1 although antigenic peptide-pulsed splenic CD11c+DCs alone failed to do so in vitro. In addition, proinflammatory stimulation of antigenic peptide-pulsed splenic CD11c+DCs with CpG ODN abrogated the antigen-specific generation of CD4+Foxp3+iTregs in vitro. On the other hand, the adoptive transfer of antigen-specific CD4+Foxp3−T cells plus antigenic peptide into mice generated CD4+Foxp3+iTregs in spleen, whereas the blockade of the function of CD11c+DCs with anti-CD11c monoclonal antibody (mAb) largely impaired this generation under steady-state conditions. In contrast, the in vivo application of CpG ODN markedly abrogated the generation of CD4+Foxp3+iTregs in mice. Collectively, our experimental models showing the crucial role of DCs in the antigen-specific generation of CD4+Foxp3+iTregs under steady-state and inflammatory conditions are useful for a better understanding of the nature of CD4+Foxp3+iTregs-mediated peripheral immune tolerance.

2. Materials 2.1. Mice

1. BALB/c mice are purchased from Charles River Laboratories (Portage, MI). 2. Ovalbumin (OVA)-specific T-cell receptor (TCR; KJ1-26 clonotype) transgenic DO11.10 BALB/c mice are purchased from The Jackson Laboratory (Bar Harbor, ME). 3. DO11.10 BALB/c mice are bred with Rag2−/−BALB/c mice (Jackson Laboratory) to generate Rag2−/− DO11.10

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BALB/c mice (see Note 1), which lack KJ1-26+CD4+ Foxp3+nTregs (6, 9, 10). 4. All mice are used between 6 and 10 weeks of age and maintained in specific pathogen-free conditions and in accordance with guidelines of the Institutional Animal Care Committee. 2.2. Cell Culture Medium and Buffer

1. RPMI 1640 medium with L-Glutamin (nakalai tesque, Kyoto, Japan) supplemented with 10% fetal bovine serum (FBS, HyClone, Ogden, UT) and Antibiotic-Antimycotic Mixed Stock Solution (nakalai tesque). Culture medium is stored at 4°C. 2. Dulbecco’s phosphate-buffered saline (D-PBS) without Calcium and Magnesium(nakalai tesque). D-PBS(−) is stored at 4°C. 3. AutoMACS Running Buffer (Miltenyi Biotec, Bergisch Gladbach, Germany). Stored at 4°C. 4. AutoMACS Rinsing Solution (Miltenyi Biotec). Stored at 4°C. 5. 10× BD IMag buffer (BD Biosciences, San Jose, CA). Stored at 4°C. 1× BD IMag buffer is prepared by diluting of 10× BD IMag buffer with sterile distilled water(1:10) and stored at 4°C. 6. Foxp3 staining buffer set (eBioscience, San Diego, CA). Stored at 4°C.

2.3. Reagents

1. Collagenase type III (Worthington Biochemical, Lakewood, NJ) is dissolved in D-PBS(−) at 4,000 U/ml, stored in singleuse aliquots at −30°C, and added to cell culture as required. 2. Mouse CD11c (N418) Microbeads (Miltenyi Biotec). Stored at 4°C. 3. Mouse CD4 T lymphocyte Enrichment Set-DM (BD Biosciences). Stored at 4°C. 4. Commercially synthesized OVA323–339 peptide (OVAp, ISQAVHAAHAEINEAGR; >90% pure) is dissolved in D-PBS(−) at 1 mM, stored in single-use aliquots at −30°C, and added to cell cultures as required. 5. Recombinant mouse interleukin (IL)-2 (Wako Pure Chemicals, Osaka, Japan) is dissolved in distilled water DNAse, RNAse free (Gibco/BRL, Bethesda, MD) at 1 mg/ml, stored in single-use aliquots at −30°C, and added to cell cultures as required. 6. Recombinant human TGF-b1 (Wako Pure Chemicals) is dissolved in Distilled Water DNAse, RNAse Free at 1 mg/ml, stored in single-use aliquots at −30°C, and added to cell cultures as required.

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7. Antimouse IFN-g mAb (clone R4-6A2, 0.5 mg/ml) is purchased from BD Biosciences, stored at 4°C, and added to cell cultures as required. 8. Antimouse IL-4 mAb (clone 11B11, 0.5 mg/ml) is purchased from BD Biosciences, stored at 4°C, and added to cell cultures as required. 9. Hamster IgG1 (clone A19-3, 0.5 mg/ml) is purchased from BD Biosciences, stored at 4°C, and injected into mice as required. 10. Antimouse CD11c mAb (clone N418, 0.5 mg/ml) is purchased from eBioscience, stored at 4°C, and injected into mice as required. 11. Antimouse CD16/CD32 mAb (clone 2.4G2, 0.5 mg/ml) is purchased from BD Biosciences, stored at 4°C, and added to cell suspensions as required. 12. R-Phycoerythrin (R-PE)-conjugated antimouse I-A/I-E mAb (clone M5/114.15.2, 0.2 mg/ml) is purchased from BD Biosciences, stored at 4°C, and added to cell suspensions as required. 13. APC-conjugated antimouse CD11c mAb (clone HL3, 0.2 mg/ml) is purchased from BD Biosciences, stored at 4°C, and added to cell suspensions as required. 14. Fluorescein isothiocyanate (FITC)-conjugated antimouse DO11.10 TCR mAb (clone KJ1-26, 0.1 mg/ml) is purchased from Invitrogen (Carlsbad, CA), stored at 4°C, and added to cell suspensions as required. 15. Allophycocyanin (APC)-conjugated anti-mouse/rat Foxp3 mAb (clone FJK-16s, 0.2 mg/ml) is purchased from eBioscience, stored at 4°C, and added to cell suspensions as required. 16. Propidium iodide (PI; Sigma-Aldrich, St Louis, MO) is dissolved in D-PBS(−) at 25 mg/ml, stored in single-use aliquots at −30°C, and added to cell suspensions as required. 17. CpG ODN 1668 (InvivoGen, San Diego, CA) is dissolved in Distilled Water DNAse, RNAse Free at 636 mg/ml (100 mM), stored in single-use aliquots at −30°C, and added to cell cultures as required. 2.4. Apparatus

1. Scissors. 2. Pincette. 3. 1-ml tuberculin syringe (Terumo, Tokyo, Japan). 4. 27G3/4 needle (Terumo). 5. 5-ml tuberculin syringe (Terumo).

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6. 35 mm/Tissue culture dish (Iwaki, Tokyo, Japan). 7. 100-mm cell strainer (BD Bioscience). 8. 50-ml polypropylene conical tube (BD Bioscience). 9. 5-ml polystyrene round-bottom tube (BD Bioscience). 10. 5-ml polystyrene round-bottom tube with cell-strainer cap (BD Bioscience). 11. 96-well flat-bottom plate (BD Bioscience). 12. 1.5-ml snaplock microtube (Axygen, Union City, CA). 13. AutoMACS Separator (Miltenyi Biotec). 14. AutoMACS Separation column (Miltenyi Biotec). 15. BD IMagnet (BD Biosciences). 16. Fluorescence-activated cell sorting (FACS)Calibur flow cytometer (BD Biosciences). 17. CELLQuest Software (BD Biosciences). 18. High-speed refrigerated microcentrifuge. 19. Refrigerated centrifuge. 20. Water-jacket-type CO2 incubator.

3. Methods 3.1. Preparation of Single-Cell Suspensions from Spleen

1. Spleen is removed from CO2-sacrificed BALB/c mice or Rag2−/− DO11.10 BALB/c mice. 2. Spleen is kept in 3 ml of culture medium containing collagenase type III (final concentration; 400 U/ml) on a 35 mm/Tissue culture dish. 3. Spleen is injected with a total volume of about 300 ml of culture medium containing collagenase type III (final concentration; 400 U/ml) at several sites using a 1-ml tuberculin syringe with a 27G3/4 needle, and incubated for 20 min at 37°C in a Water-jacket-type CO2 incubator. 4. Digested spleen is set on a 100-mm cell strainer in culture medium containing collagenase type III (final concentration; 400 U/ml) on a 35 mm/Tissue culture dish and ground by the plunger of a 5-ml tuberculin syringe. 5. Single-cell suspensions are obtained by forcing through a 100-mm cell strainer on 35 mm/Tissue culture dishes (see Note 2). 6. Single-cell suspensions are collected in 50-ml polypropylene conical tubes, and 30 ml of D-PBS(−) added to the tubes.

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7. Cells are recovered by centrifugation (780 × g, 5 min, 4°C), and all the supernatants are carefully discarded. 8. Cell pellets are resuspended in 10 ml of culture medium. 9. Cells are counted and kept on ice (see Note 3). 3.2. Purification of CD11c+DCs from Single-Cell Suspensions

CD11c+DCs are positively selected with Mouse CD11c (N418) Microbeads and the AutoMACS Separator from splenic single-cell suspensions obtained from BALB/c mice according to the manufacturer’s instructions (see Note 4) with some modifications. 1. Splenic single-cell suspensions obtained from BALB/c mice are washed with a 10× excess volume of AutoMACS Running Buffer in 50 ml polypropylene conical tubes and centrifuged (780 × g, 5 min, 4°C), and all the supernatants are carefully discarded. 2. Resuspend cell pellets in 400 ml of AutoMACS Running Buffer per 108 cells. 3. Add 100 ml of Mouse CD11c (N418) Microbeads per 108 cells. 4. Mix well and incubate for 15 min at 4°C in the refrigerator. 5. Wash the labeled cells by adding 1–2 ml of AutoMACS Running Buffer per 107 total cells and centrifuge (780 × g, 5 min, 4°C). Carefully discard all the supernatants. 6. Resuspend up to 108 cells in 500 ml of AutoMACS Running Buffer, and filter through a 5-ml polystyrene round-bottom tube with a cell-strainer cap. 7. Prepare and prime the AutoMACS Separator (see Note 5). 8. Apply the tube containing the sample and provide tubes (50-ml polypropylene conical tubes) for collecting the labeled and unlabeled cell fractions. 9. Place sample tube at the uptake port and the fraction collection tubes at port neg1 and port pos2. The positive fraction contains CD11c+DCs with Mouse CD11c (N418) Microbeads. 10. Recover CD11c+DCs by centrifugation (780 × g, 5 min, 4°C), and carefully discard all the supernatants. 11. Cell pellets are resuspended in 10 ml of culture medium. 12. Cells are counted and kept on ice (see Note 6).

3.3. Purification of CD4+T Cells from Single-Cell Suspensions

CD4+T cells are negatively selected with mouse CD4 T lymphocyte Enrichment Set-DM and BD IMagnet from splenic singlecell suspensions obtained from BALB/c mice or Rag2−/−DO11.10 BALB/c mice (Rag2−/−KJ1-26+T cells) according to the manufacturer’s instructions (see Note 7) with some modifications.

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1. Splenic single-cell suspensions obtained from BALB/c mice or Rag2−/−DO11.10 BALB/c mice are washed with a 10× excess volume of 1× BD IMag buffer in 50-ml polypropylene conical tubes and centrifuged (780 × g, 5 min, 4°C), and all the supernatants are carefully discarded. 2. Cell pellets are resuspended in 1× BD IMag buffer at a concentration of 20 × 106 cells/ml. 3. Add the Biotinylated Mouse CD4 T Lymphocyte Enrichment Cocktail at 5 ml per 1 × 106 cells and incubate on ice for 15 min. 4. Wash the labeled cells with a 10× excess volume of 1× BD IMag buffer, centrifuge (780 × g, 5 min, 4°C), and carefully discard all the supernatants. 5. Vortex the BD™ IMag Streptavidin Particles Plus – DM thoroughly, and add 5 ml of particles for every 1 × 106 cells. 6. Mix thoroughly. Refrigerate for 30 min at 4°C. 7. Bring the labeling volume up to 20 to 106 cells/ml with 1× BD IMag buffer. 8. Transfer the labeled cells to a 5-ml polystyrene round-bottom tube, maximum volume added not to exceed 3.0 ml. 9. Place this positive-fraction tube on the BD IMagnet (horizontal position) for 6 min. 10. With the tube on the BD IMagnet and using a 1 ml-pipetman, carefully aspirate the supernatant (enriched fraction) and place in a new sterile tube. 11. Remove the positive-fraction tube from the BD IMagnet, and add 1× BD IMag buffer to the same volume as in step 7. Resuspend the positive fraction well by pipetting up and down 15 times, and place the tube back on the BD IMagnet for 6 min. 12. Using a 1 ml-pipetman, carefully aspirate the supernatant and combine with the enriched fraction from step 10 above. 13. Repeat steps 11 and 12. The combined enriched fraction contains CD4+T cells with no bound antibodies or magnetic particles. 14. CD4+T cells are recovered by centrifugation (780 × g, 5 min, 4°C), and all the supernatants are carefully discarded. 15. Cell pellets are resuspended in 10 ml of culture medium and filtered through a 5-ml polystyrene round-bottom tubes with cell-strainer cap. 16. Cells are counted and kept on ice (see Note 8).

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3.4. In Vitro Conversion Assay

1. Prepare an appropriate volume of culture medium containing OVAp (final concentration; 1 mM), anti-IFN-g mAb (final concentration; 10 mg/ml), anti-IL-4 mAb (final concentration; 10 mg/ml), and recombinant mouse IL-2 (final concentration; 0.2 ng/ml) under neutral conditions in the presence or absence of recombinant human TGF-b1 (final concentration; 10 ng/ml) and/or CpG ODN 1668 (final concentration; 0.1 mM). 2. Add Rag2−/−KJ1-26+T cells (2 × 105 in 500 ml) and CD11c+DCs (2 × 104 in 500 ml) into a 1.5-ml snaplock microtube, spin down (151 × 100g, 1 min, 4°C), and carefully discard all the supernatants. 3. Resuspend cell pellets in 250 ml of culture medium as prepared in step 1 with pulse vortexing and add them per well to a 96-well flat-bottom plate. 4. Culture for 3 days at 37°C in a Water-jacket type CO2 incubator. 5. Collect the culture from each well in a 1.5-ml snaplock microtube, wash once by adding 1 ml of D-PBS(−). Spin down (151 × 100 × g, 1 min, 4°C) and carefully discard all the supernatants. 6. Resuspend cell pellets in 100 ml of D-PBS(−) and keep on ice.

3.5. Adoptive Transfer

1. Prepare 500 ml/mouse of D-PBS(−) containing OVAp (5 mg) in the presence or absence of CpG ODN 1668 (50 mg/ mouse) in a 1.5-ml snaplock microtube. 2. Move Rag2−/−KJ1-26+T cells (5x106 in 250 ml) into 1.5-ml snaplock microtube, wash once by adding 1 ml of D-PBS(−). Spin down (151 × 100 × g, 1 min, 4°C) and carefully discard all the supernatants. 3. Resuspend cell pellets in 500 ml of D-PBS(−) containing OVAp (5 mg) in the presence or absence of CpG ODN 1668 (50 mg/mouse) as prepared in step 1 with pulse vortexing. 4. Inject BALB/c mice intravenously through a tail vein with Rag2−/−KJ1-26+T cells (5 × 106/mouse) alone or Rag2−/−KJ1-26+T cells (5 × 106/mouse) plus OVAp (5 mg/ mouse) in the presence or absence of CpG ODN 1668 (50 mg/mouse) using a 1-ml-tuberculin syringe with a 27G3/4 needle. 5. Inject BALB/c mice intraperitoneally with or without hamster IgG used as a control Ig or anti-CD11c mAb (clone N418) (each 250 mg/500 ml/mouse) using a 1-ml-tuberculin syringe with a 27G3/4 needle on day −1, 1, 3, 5, 7, 9, and 11 before and after adoptive transfer.

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6. After 12 days, remove spleens from the recipient mice, and prepare splenic single-cell suspensions as described above. 7. Recover cells by centrifugation (780 × g, 5 min, 4°C), and carefully discard all the supernatants. 8. Resuspend cell pellets in 10 ml of culture medium. 9. Count cells and keep on ice. 3.6. Flow Cytometry of Splenocytes and CD11c +DCs

1. Add single-cell suspensions of splenocytes (5 × 106) and CD11c+DCs (5 × 105) to each 1.5-ml snaplock microtube. Spin down (151 × 100 × g, 1 min, 4°C) and carefully discard all the supernatants. 2. Resuspend cell pellets in 1 ml of D-PBS(−) and spin down (151 × 100 × g, 1 min, 4°C). Carefully discard all the supernatants. 3. Resuspend cell pellets in 100 ml of D-PBS(−) and keep on ice. 4. Preincubate the cells with antimouse CD16/CD32 mAb (1 mg per 106) with pulse vortexing for 5 min on ice prior to staining. 5. Add R-PE-conjugated antimouse I-A/I-E mAb (clone M5/114.15.2; 1 mg per 106) and APC-conjugated antimouse CD11c mAb (clone HL3; 1 mg per 106) to each tube with pulse vortexing and incubate for 30 min on ice in the dark. 6. Wash the labeled cells by adding 1 ml of D-PBS(−) and spin down (151 × 100 × g, 1 min, 4°C). Carefully discard all the supernatants. 7. Resuspend stained cell pellets in 500 ml of D-PBS(−) containing PI (see Note 9) with pulse vortexing and analyze samples on a FACSCalibur flow cytometer with CELLQuest Software (see Note 9). Example results are shown in Figs. 1a and 2a.

3.7. Flow Cytometry of Splenocytes and Rag2−/−KJ1-26+T Cells

The following protocol for intracellular staining of Foxp3 is performed using APC-conjugated antimouse/rat Foxp3 mAb and Foxp3 staining buffer set according to the manufacturer’s instructions (see Note 10) with some modifications. 1. Add single-cell suspensions of CD4+T cells (5 × 106) obtained from the recipient mice and Rag2−/−KJ1-26+T cells (5 × 105) to each 1.5-ml snaplock microtube. Spin down (151 × 100 × g, 1 min, 4°C) and carefully discard all the supernatant. 2. Resuspend cell pellets in 1 ml of D-PBS(−) and spin down (151 × 100 × g, 1 min, 4°C). Carefully discard all the supernatant. 3. Resuspend cell pellets in 100 ml of D-PBS(−), and keep on ice.

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Fig. 1. The ability of CD11c+DCs to generate antigen-specific Rag2−/−KJ1-26+Foxp3+iTregs from Rag2−/−KJ1-26+Foxp3−T cells. (a) The phenotype of CD11c+DCs (left panel) and Rag2−/−KJ1-26+T cells (right panel) was analyzed by flow cytometry, and data are represented by a dot plot, and numbers represent the proportion of I-A/I-E+CD11c+cells (left panel; see Note 11) or KJ1-26+Foxp3+T cells (right panel; see Note 12) in each quadrant. A large proportion of CD11c+DCs expressed I-A/I-E. Rag2−/−KJ1-26+T cells did not express Foxp3. (b) Rag2−/−KJ1-26+T cells (2 × 105) were cultured with CD11c+DCs (2 × 104) in combination with OVAp (1 mM), anti-IFN-g mAb (10 mg/ml), anti-IL-4 mAb (10 mg/ml), and IL-2 (0.2 ng/ml) in the presence or absence of TGF-b1 (10 ng/ml) or CpG ODN 1668 (0.1 mM) for 3 days. The expression of Foxp3 among gated KJ1-26+T cells was analyzed by flow cytometry. Data are represented by a dot plot, and numbers represent the proportion of KJ1-26+Foxp3+T cells (see Note 12) in each quadrant. OVAppulsed CD11c+DCs induced the conversion of Rag2−/−KJ1-26+Foxp3−T cells into Rag2−/−KJ1-26+Foxp3+T cells in the presence of TGF-b1, whereas OVAp-pulsed CD11c+DCs alone failed to induce this conversion. On the other hand, stimulation of OVAp-pulsed CD11c+DCs with CpG ODN 1668 suppressed the TGF-b1-mediated generation of Rag2−/−KJ1-26+Foxp3+T cells.

4. Preincubate the cells with antimouse CD16/CD32 mAb (1 mg per 106) with pulse vortexing for 5 min on ice prior to staining. 5. Add FITC-conjugated antimouse DO11.10 TCR mAb (0.1 mg per 106) to each tube with pulse vortexing and incubate for 30 min on ice in the dark. 6. Wash the labeled cells by adding 1 ml of D-PBS(−) and spin down (151 × 100 × g, 1 min, 4°C). Carefully discard all the supernatants. 7. Resuspend cell pellets with 1 ml of freshly prepared Fixation/ Permeabilization working solution. 8. Refrigerate for 2 h at 4°C in the dark.

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Fig. 2. Antigen-specific de novo generation of Rag2−/−KJ1-26+Foxp3+iTregs from Rag2−/−KJ1-26+Foxp3−T cells. Rag2−/−KJ126+T cells (5 × 106/mouse) alone or Rag2−/−KJ1-26+T cells (5 × 106/mouse) plus OVAp (5 mg/mouse) in the presence or absence of CpG ODN 1668 (50 mg/mouse) were transferred into BALB/c mice that had been treated with control Ig or anti-CD11c mAb (each 250 mg/mouse), and the spleen was removed from the recipient BALB/c mice on day 12 after the adoptive transfer. (a) The expression of I-A/I-E and CD11c on splenocytes was analyzed by flow cytometry, and data are represented by a dot plot. Numbers represent the proportion of I-A/I-E+CD11c+cells (see Note 11) in each quadrant. Repetitive injections of anti-CD11c mAb blocked the staining of CD11c on I-A/I-E+CD11c+cells in spleen (see Note 13). (b) The frequency of KJ1-26+T cells among CD4+T cells was analyzed by flow cytometry. Data are represented by a histogram, and numbers represent the proportion of KJ1-26+T cells (see Note 11) in each quadrant. The transplanted Rag2−/−KJ1-26+T cells were detected in the splenic CD4+T cells in recipient BALB/c mice. (c) The expression of Foxp3 among gated KJ1-26+T cells was analyzed by flow cytometry. Data are represented by a dot plot, and numbers represent the proportion of KJ1-26+Foxp3+T cells (see Note 12) in each quadrant. Adoptive transfer of Rag2−/−KJ1-26+Foxp3−T cells plus OVAp generated Rag2−/−KJ1-26+Foxp3+T cells, whereas adoptive transfer of Rag2−/−KJ1-26+T cells alone did not induce them. The blockade of the function of I-A/I-E+CD11c+cells abrogated the antigen-specific generation of Rag2−/−KJ126+Foxp3+T cells from Rag2−/−KJ1-26+Foxp3−T cells (see Note 13). The injection of CpG ODN 1668 suppressed the antigen-specific generation of Rag2−/−KJ1-26+Foxp3+T cells from Rag2−/−KJ1-26+Foxp3−T cells.

9. Spin down (151 × 100 × g, 1 min, 4°C) and carefully discard all the supernatants. 10. Resuspend cell pellets in 1 ml of D-PBS(−) and spin down (151 × 100 × g, 1 min, 4°C). Carefully discard all the supernatants. 11. Resuspend cell pellets in 1 ml of 1× Permeabilization Buffer (made from 10× Permeabilization Buffer diluted with sterile

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distilled water) followed by spin down (151 × 100 × g, 1 min, 4°C) and careful decanting of supernatants. 12. Resuspend cell pellets in 100 ml of 1× Permeabilization Buffer. 13. Preincubate the cells with antimouse CD16/CD32 mAb (1 mg per 106) with pulse vortexing for 5 min on ice prior to staining. 14. Add APC-conjugated antimouse/rat Foxp3 mAb (0.5 mg per ~5 × 106) to each tube with pulse vortexing and incubate for 30 min on ice in the dark. 15. Wash the labeled cells by adding 1 ml of 1× Permeabilization Buffer and spin down (151 × 100 × g, 1 min, 4°C). Carefully discard all the supernatants. 16. Resuspend stained cell pellets in 500 ml of D-PBS(−) with pulse vortexing and analyze samples on the FACSCalibur flow cytometer with CELLQuest Software (see Note 9). Example results are shown in Figs. 1a, b, and 2b, c.

4. Notes 1. Rag2−/− DO11.10 BALB/c mice are obtained from elsewhere through a material transfer agreement (MTA). 2. Undigested fibrous material is removed by filtration through a 100-mm cell strainer. 3. The number of total splenocytes obtained from BALB/c mice and Rag2−/− DO11.10 BALB/c mice is approximately 1.5 × 108/mouse and 5 × 107/mouse, respectively. 4. Technical data sheet is available at http://www.miltenyibiotec. com/download/datasheets_en/70/DS130-052-001.pdf. 5. AutoMACS Separator program is “Posseld2.” 6. The number of CD11c+DCs obtained from total splenocytes (1.5 × 108/mouse) of BALB/c mice is approximately 1.5 × 106/ mouse. 7. Technical data sheet is available at http://www.bdbiosciences. com/external_files/pm/doc/tds/cell_sep/live/web_ enabled/558131.pdf. 8. The number of CD4+T cells obtained from total splenocytes (1.5 × 108/mouse) of BALB/c mice and Rag2−/−KJ1-26+T cells obtained from total splenocytes (5 × 107/mouse) of Rag2−/− DO11.10 BALB/c mice is approximately 107/mouse and 3.5 × 106/mouse, respectively. 9. PI-stained cells are excluded from the analysis.

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10. Technical data sheet is available at http://www.ebioscience. com/ebioscience/appls/FCI.htm#foxp3. 11. Data are the results of cell surface staining. 12. Data are the results of intracellular staining. 13. Our preliminary experiments showed that the injection with anti-CD11c mAb (clone N418) caused a minimal depletion of CD11c+cells in spleen (less than 20% of total CD11c+cells) when CD11c-diptheria toxin (DT) receptor (DTR) transgenic (B6.FVB-Tg Itgax-DTR/GFP 57Lan/J) mice (11) were used. On the other hand, anti-CD11c mAb (clone N418) blocked the binding of CD11c to various cell adhesion molecules in vitro, and the treatment of mice with antiCD11c mAb (clone N418) suppressed the sheep red blood cells (SRBC)-induced delayed-type hypersensitivity (DTH) (12). Therefore, the suppressive effect of anti-CD11c mAb (clone N418) on the generation of Rag2−/−KJ1-26+Foxp3+T cells in vivo might be due to the blockade of the function of CD11c+cells rather than the depletion of CD11c+cells.

Acknowledgements The author would like to thank Yumiko Sato, Kaori Sato, Kawori Eizumi, and Naomi Uchimura for excellent technical assistance. This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Science, and Culture of Japan (c) 17790334 and 19590505 (K.S.). References 1. Sato, K. and Fujita, S. (2007) Dendritic cellsnature and classification. Allergol. Int. 56, 183–191. 2. Mucida, D., Park, Y., Kim, G., Turovskaya, O., Scott, I., Kronenberg, M., et al. (2007) Reciprocal TH17 and regulatory T cell differentiation mediated by retinoic acid. Science 317, 256–260. 3. Kretschmer, K., Apostolou, I., Hawiger, D., Khazaie, K., Nussenzweig, M. C., and von Boehmer, H. (2005) Inducing and expanding regulatory T cell populations by foreign antigen. Nat. Immunol. 6, 1219–1227. 4. Birnberg, T., Bar-On, L., Sapoznikov, A., Caton, M. L., Cervantes-Barragán, L., Makia, D., et al. (2008) Lack of conventional dendritic cells is compatible with normal development and T cell homeostasis, but causes

5.

6.

7.

8.

myeloid proliferative syndrome. Immunity 29, 986–997. Ohnmacht, C., Pullner, A., King, S. B., Drexler, I., Meier, S., Brocker, T., et al. (2009) Constitutive ablation of dendritic cells breaks self-tolerance of CD4 T cells and results in spontaneous fatal autoimmunity. J. Exp. Med. 206, 549–559. Hori, S., Nomura, T., and Sakaguchi, S. (2003) Control of regulatory T cell development by the transcription factor Foxp3. Science 299, 1057–1061. Curotto de Lafaille, M. A. and Lafaille, J. J. (2009) Natural and adaptive Foxp3+ regulatory T cells: More of the same or a division of labor? Immunity 30, 626–635. Zhou, L., Chong, M. M., and Littman, D. R. (2009) Plasticity of CD4+ T cell lineage differentiation. Immunity 30, 646–655.

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9. Fujita, S., Sato, Y., Sato, K., Eizumi, K., Fukaya, T., Kubo, M., et al. (2007) Regulatory dendritic cells protect against cutaneous chronic graft-versus-host disease mediated through CD4+CD25+Foxp3+T cells. Blood 110, 3793–3803. 10. Sato, K., Eizumi, K., Fukaya, T., Fujita, S., Sato, Y., Takagi, H., et al. (2009) Naturally occurring regulatory dendritic cells regulate murine cutaneous chronic graft-versus-host disease. Blood 113, 4780–4789.

11. Jung, S., Unutmaz, D., Wong, P., Sano, G., De Los Santos, K., Sparwasser, T., et al. (2002) In vivo depletion of CD11c+ dendritic cells abrogates priming of CD8+ T cells by exogenous cell-associated antigens. Immunity 17, 211–220. 12. Sadhu, C., Ting, H.J., Lipsky, B., Hensley, K., Garcia-Martinez, L.F., Simon, S.I., et al. (2007) CD11c/CD18: novel ligands and a role in delayed-type hypersensitivity. J. Leukoc. Biol. 81, 1395–1403.

Chapter 9 Plasmacytoid Dendritic Cells in Tolerance Eric Gehrie, William Van der Touw, Jonathan S. Bromberg, and Jordi C. Ochando Abstract Dendritic cells (DC) are professional antigen-presenting cells (APCs) that modulate the outcome of the immune response toward immunity or tolerance. There are a large variety of DC subsets according to surface phenotype, function, and tissue distribution. Murine plasmacytoid DC (pDC) represent a distinctive DC population and are characterized by the expression of CD11c, B220, Gr-1, CD45RA, Ly49Q, BST2, and siglec-H on the cell surface. PDC act as immunogenic cell sentinels by secreting large amounts of type I interferon (IFN) in the lymph nodes in response to viral stimulation. PDC also act as tolerogenic cells when expressing the inducible tolerogenic enzyme indoleamine 2,3-dioxygenase (IDO), the inducible costimulator ligand (ICOS-L), and/or the programmed death 1 ligand (PD-L1), which mediate regulatory T-cell (Treg) development and suppression of self- and alloreactive cells. The PDC ability to induce Treg development is associated with capture and presentation of antigenic peptides associated with major histocompatibility complex (MHC) class I and II. Here, we provide the tools to study PDC development from bone marrow cultures, their antigen presentation properties, and their interactions with Treg under a tolerogenic setting of sterile inflammation. Key words: Plasmacytoid dendritic cells, Antigen presentation, Regulatory T cells

1. Introduction Dendritic cells (DC) are responsible for the initiation and orchestration of the adaptive immune response. To do so, DC patrol throughout the body looking for antigen that may be present within the tissues or in circulation. DC are highly efficient antigenpresenting cells (APCs) on the major histocompatibility complex (MHC), and following antigen uptake, DC migrate to secondary lymphoid organs (SLO) and present peptide/MHC complexes to antigen-specific CD4+ and CD8+ T cells. These interactions are

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_9, © Springer Science+Business Media, LLC 2011

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key factors in the establishment of either immunity or tolerance, and many DC subsets are involved in these processes. The DC family involves a large variety of subsets according to their cell surface phenotype and tissue distribution. In the mouse, all DC subsets described express the integrin CD11c. Murine plasmacytoid DC (PDC) are a DC subset that are generated from hematopoietic precursors in the bone marrow (1) and constantly circulate in blood and SLO in a quiescent immature state expressing significative levels of CD11c, B220, Gr-1, CD45RA, Ly49Q, BST2, and siglec-H (2–7). Upon Ag recognition, PDC undergo a maturation process, upregulate their expression of MHC-II on the cell surface, and migrate to T-cell areas of secondary lymphoid tissues where they can either induce activation and proliferation of immunogenic T lymphocytes, or the development of tolerogenic regulatory T cells (Treg). Tolerogenic PDC have been shown to induce T-cell unresponsiveness under steady state conditions (8). Tolerogenic PDC cells have also been described to suppress the immune response in several inflammatory disorders such as acute graft-versus-host disease (GVHD) (9), autoimmune arthritis (10), oral tolerance (11), and a lower frequency of PDC is associated with Type 1 diabetes (12, 13) and diabetes in mice (14). As a result, protolerogenic interaction between PDC and Treg has already been suggested (8, 15) which seems to be responsible for antigenspecific suppression of CD4+ and CD8+ T-cell responses (8, 16). The mechanisms by which PDC may promote Treg development have been investigated. PDC induce in vivo differentiation of naïve CD4+ T cells into Treg through indoleamine 2,3-dioxygenase (IDO) (17, 18). Treg can also be induced by PD-L1 (19), and PDC expressing higher levels of PD-L1 also correlate with an elevated frequency of Treg in tolerized recipient (20); PDC poised to express ICOS-L upon maturation leads to the generation of IL-10-producing T regulatory cells (21). IDO expression by PDC has been reported in tumor-draining lymph nodes (22), which directly activate mature Treg through PD-L1 (23). These studies suggested tumor antigen presentation by PDC in the lymph nodes, consistent with Jung and colleagues, who reported that antigen-bearing PDCs prime naïve CD4 T cell in lymph nodes, but not in the spleen (24). Although PDC are generally considered poor presenters of exogenous antigens, some in vivo studies addressing the contribution of PDC to antigen presentation suggested that PDC are able to present antigen and prime CD4 T cells during infection with Toxoplasma gondii (25) and during sterile inflammatory conditions of GVHD (26). Activated murine PDC are characterized by a sustained expression of MHC-II on the cell surface, which is accompanied by prolonged antigen uptake, processing, and presentation (27, 28). It is possible that prolonged antigen presentation by PDC to

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CD4 T cells may promote antigen-specific Treg development, since MHC-II expression by DC is required to maintain Treg cell homeostasis (29). In this chapter, we describe the protocol to follow alloantigen presentation of MHC-II-derived peptides by PDC in a model of sterile inflammation. Induction of transplantation tolerance can be achieved in mice following costimulatory blockade with anti-CD40L mAb, and interactions of alloantigen-presenting PDC with Foxp3 expressing Treg can be monitored (30). Owing to their infrequency, PDC can be very challenging to study, making up only 0.1–0.5% of all peripheral blood mononuclear cells (PBMCs). For this reason, culturing total BM in vitro in the presence of different cytokines allows the generation of a large number of PDC that can be further used for molecular or cell biology studies. Pioneering research from Maraskovsky and colleagues described that in vivo treatment with FLT3-ligand (FLT3-L) expanded the pool of murine and human DC (31, 32), including PDC, which were characterized by their phenotype of CD4+CD11c−CD45RA+IL3-R+MHC-II+. Similar results were obtained in vitro by adding FLT3-L to bone marrow cultures, with the generation of two populations of CD11c+CD86+MHC-II+ myeloid and lymphoid DC, that differ in their expression of CD11b (33). Once PDC were identified in mice by their expression of CD11c+B220+Gr-1+ (2), several groups reported their specific development from FLT3-L in vitro bone marrow cultures (34, 35). Here, we describe culture conditions for the in vitro generation of PDC with FLT3-L and investigate their potential to induce Treg. In these protocols, we set out to differentiate PDC from BM precursors supplemented with FLT3-L in in vitro cultures. Treatment with FLT3-L rapidly differentiates BM cells into two distinct population of DC, which can be further identified by their expression of siglec-H. In this chapter, we describe different systems used to differentiate BM cells into PDC, and their development into protolerogenic cells following stimulation with FLT3-L, since repetitive injections of FLT3-L results in peripheral Treg expansion (36).

2. Materials 2.1. In Vitro Generation of PDC from Murine Bone Marrow Cells 2.1.1. Mouse Handling

(Experiment time: 3–5 h + 10 day incubation).

1. BALB/c mice. 2. Ketamine HCl (100 mg/mL or 50 mg/mL). 3. Acepromazine (10 mg/mL).

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4. 70% Ethanol, 100 mL. 5. Styrofoam board, ×1. 6. Surgical scissors. 7. Surgical forceps. 8. 18-Gauge 1.5″ needles, ×6. 2.1.2. Cell Cultures

The following are required for the culture of BM cells: 1. Sterile PBS, 500 mL bottle. 2. RBC lysis solution. 3. 70 mM Cell strainer, ×3. 4. 50 mL Centrifuge tubes, ×3. 5. 1.5 mL Microcentrifuge tubes, ×1. 6. Hemocytometer, ×1. 7. 0.4% Trypan blue. 8. Petri dishes, ×4. 9. 10 mL Glass pipette, ×1. 10. 10 mL Syringes, ×1 (per mouse). 11. 1 mL Syringe, ×1. 12. 3 mL Syringe, ×1. 13. 30-Gauge needles, ×1. 14. PDC Medium, see Appendix 2. 15. FLT3-L, see Appendix 3.

2.2. Sorting Murine Bone Marrow Cultures to Achieve Pure Populations of PDC 2.2.1. PDC Cell Sorting (see Fig. 1)

Experiment time: 2–3 h preparation + sorting time.

1. Mature bone marrow cultures (see Subheading 2.1). 2. 70 mM Cell strainer, ×3. 3. 5 mL Pipette, ×2. 4. 10% Fetal Bovine Serum RPMI 1640 Complete Medium. 5. Staining buffer: phosphate-buffered saline (PBS) supplemented with 2% FCS (see Note 1). 6. DAPI, 1 mL of 1 mg/mL solution. 7. Unconjugated anti-Fc receptor CD16/CD32 antibody. 8. PerCP Cy5.5-conjugated anti-mouse-CD11c antibody. 9. PerCP Cy5.5-conjugated Armenian Hamster IgG Isotype Control. 10. APC-conjugated anti-mouse-B220 antibody.

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Fig. 1. Plasmacytoid dendritic cell phenotype. (a) BM cells were isolated from BALB/c mice and cultured for 10 days in the presence of FLT3-L. Data can be analyzed using FlowJo software (TriStar) and BM of BALB/c mice can be divided into two major subpopulations, defined as PDC and mDC. In order to gate on PDCs, 100,000 events should be acquired for each sample. The first step in the analysis is to draw a gate looking at side scatter (SSC) versus forward scatter (FSC). This gate will be gate 1. Cell duplets are excluded in gate 2 (SSC-A versus SSC-H). Dead cells are excluded in gate 3 (SSC-A versus DAPI). Making a drill down of gate 3, we can then identify PDCs within the BM culture, by making a new gate looking at CD11c intermediate versus B220 high. This cell population should be CD11b negative. The PDC population also can be gated on from BM cultures on surface staining for Siglec-H and mPDCA-1.

11. APC-conjugated Rat IgG2a Isotype Control. 12. FITC-conjugated anti-mouse-CD11b antibody. 13. FITC-conjugated Rat IgG2b Isotype Control. 14. PE-conjugated anti-mouse-Gr-1 antibody. 15. 4 mL Falcon tube (see Note 2). 16. 50 mL Falcon tube, ×6–7.

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2.3. Harvesting and Sorting Naïve T Cells from Tea Mouse Spleens

1. BALB/c mice.

2.3.1. Mouse Handling

5. Surgical scissors.

2. Ketamine HCl (100 mg/mL). 3. Acepromazine (10 mg/mL). 4. 70% Ethanol, 100 mL. 6. Surgical forceps. 7. RPMI 1640 complete medium supplemented with 10% FCS. 8. 50 mL Centrifuge tubes, ×3.

2.3.2. T-Cell Sorting

1. APC-CD4. 2. FITC-CD25. 3. PE-Foxp3. 4. 500 mL Sterile PBS. 5. RPMI 1640 complete medium supplemented with 10% FCS. 6. RBC lysis solution. 7. 70 mM Sterile cell strainer, ×3.

2.4. Culturing Bone Marrow-Derived PDC with Naïve T Cells

1. 96-well, round-bottom culture plates.

2.4.1. PDC and Naïve T Cell Co-culture

4. Human TGFb1.

2.5. Staining for Surface Markers and Intracellular Foxp3 for FACS Analysis 2.5.1. Surface Staining for CD4 and CD25 with Intracellular Foxp3 FACS Staining (see Fig. 2)

2. Murine IL-2. 3. Murine anti-CD3 mAb. 5. RPMI 1640 complete medium. 1. PBS, 500 mL. 2. Distilled water 500 mL. 3. Fixation/permeabiliziation concentrate and dilution solution, 1 mL/sample (see Note 3). 4. 3 mL of prepared permeabilization solution and isotype control (see Note 4). 5. FITC-labeled anti-mouse CD25 antibody and isotype control, 1 mL/sample. 6. APC-labeled anti-mouse CD4 antibody and isotype control, 1 mL/sample. 7. PE-labeled anti-mouse Foxp3 antibody, 1 mL/sample. 8. 1.5 mL Microcentrifuge tubes. 9. Tubes suitable for flow cytometry.

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Fig. 2. Induction of regulatory T cells with BM-derived plasmacytoid dendritic cells. Freshly sorted BM-derived PDC cells (2 × 104 cells/well) isolated from BALB/c mice for 10 days in the presence of FLT3-L, were cultured with CD4+CD25− T cells (5 × 104 cells/well) for 96 h with 150 ng/mL of anti-CD3 mAb, 10 ng/mL IL-2, and 10 ng/mL TGFb. Flow cytometry data indicates percentage of Foxp3+ CD4+ T cells induced.

2.6. Identification of AlloantigenExpressing PDC

1. Biotin mouse IgG2b isotype control.

2.6.1. PDC-YAe FACS Staining (see Fig. 3)

4. Biotin YAe mAb.

2. Rat serum. 3. Unconjugated anti-Fc receptor CD16/CD32 antibody. 5. APC anti-siglec-H mAb. 6. Streptavidin–PE.

2.6.2. PDC-YAe IHC Staining (see Fig. 4)

1. Biotin mouse IgG2b isotype control (see Note 6). 2. Rat serum. 3. Unconjugated anti-Fc receptor CD16/CD32 antibody. 4. Biotin YAe mAb. 5. Anti-siglec-H mAb. 6. Anti-Foxp3 mAb.

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Fig. 3. Alloantigen-presenting PDCs circulate systemically through blood. Percentage of alloantigen-presenting YAe+ cells from the blood of tolerant mice are shown. Flow cytometry results indicate that only PDC are presenting alloantigen systemically, since only YAe+ cells express mPDCA-1. Other cell types on freshly isolated blood do not express YAe in their cell surface. Data are representative results from three independent experiments (n = 3).

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Fig. 4. Alloantigen-presenting PDCs are present in secondary lymphoid organs. Images of total alloantigen-presenting YAe+ cells from the lymph nodes of tolerized mice at day 5 posttransplant are shown. IHC results indicate that only PDC are presenting alloantigen, since most YAe+ cells express mPDCA-1. Data are representative results from three independent experiments (n = 3). Original magnification, ×200.

Fig. 5. PDCs are next to Foxp3 expressing cells in the lymph nodes of tolerized mice. IHC results indicate that Foxp3 expressing cells are in proximity of PDC in the lymph nodes of tolerized mice at day 5 posttransplant. Data are representative results from three independent experiments (n = 3). Original magnification, ×200.

7. Biotinyl Tyramide Kit. 8. Amplification diluent. 9. Tyramide reagent. 10. Strepavidin–HRP. 11. Peroxidase blocking reagent. 12. Cy3-Streptavidin and Cy2 anti-rat mAb. 2.7. Visualization of PDC and Foxp3 Cells

1. Rat serum.

2.7.1. PDC-Foxp3 IHC Staining (see Fig. 5)

4. Anti-siglec-H mAb.

2. Unconjugated anti-Fc receptor CD16/CD32 antibody. 3. Anti-Foxp3 mAb. 5. Anti-Foxp3 mAb. 6. Cy3 anti-rabbit and Cy2 anti-rat mAb.

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3. Methods 3.1. In Vitro Generation of PDC from Murine Bone Marrow Cells 3.1.1. Mouse Handling

1. Make a sterile solution of 2% ketamine and 0.02% acepromazine in PBS (see Appendix 1). 2. Fill a sterile 1 mL syringe with 200 mL of ketamine/acepromazine solution (per mouse). Attach a sterile 30-gauge needle to the syringe. 3. Inject 200 mL of ketamine/acepromazine solution (per mouse) intraperitoneally (IP). Wait a few minutes for the anesthesia to take effect and sacrifice the anesthetized mice by cervical dislocation. 4. Spray the sacrificed mice with a 70% ethanol solution to reduce the chance of microbial contamination. 5. Take a Styrofoam board and cover it with a clean paper towel. 6. Place the mouse in a supine position on the Styrofoam board. Use 18-gauge needles to pin the mouse by its four paws so that the abdomen is stretched and the mouse is secured to the Styrofoam board. 7. Using a clean pair of forceps and surgical scissors, make a small midline opening in the inferior abdomen. Make the opening as superficial as possible, being careful to avoid opening the peritoneum. 8. Insert the scissors into the inferior aspect of the opening and cut down the anterior aspect of the leg, in an inferior-lateral direction, all the way to the paw. Perform this bilaterally. 9. The incision should resemble an inverted “Y” shape. Using your gloved fingers, separate the peritoneum from the skin on both sides of the mouse and use additional 18-gauge needles to pin the abdominal skin to the Styrofoam tray. Also, using your gloved fingers and the forceps, peel the skin off of the legs. 10. Use the scissors to remove the legs from the mouse. In order to maximize yield of bone marrow and to avoid contamination, it is best for the legs to be removed whole, with the femoral head intact. Cutting through the tissue in the upper leg region carefully will eventually bring the femoral head into view. Cutting through the tissue just superior and lateral to the femoral head will dislocate the femur from the acetabulum. Place the legs, with muscle tissue still attached, into a Petri dish that is placed on ice. 11. Replace the paper towel on the Styrofoam board with a clean one. Dispose of the needles in the sharps container. Remove the muscle from each leg to the greatest extent necessary. It is recommended to start by cutting off the hind paw, inferior to the ankle joint. Then, use the scissors to cut all of the tendons around the inferior tibia and use the forceps to peel

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off the leg muscle in the superior direction. Remove the rest of the tissue using the scissors and forceps, working along the natural tissue planes as much as possible. 12. As the legs are cleaned of muscle, place them one by one into a new Petri dish, and keep them on ice until all of the legs are clean. 13. Place the Petri dish containing the cleaned legs inside the tissue culture hood. Fill the Petri dish with ~40 mL of 70% ethanol solution. Make sure that all of the legs are completely submerged in the ethanol solution and leave them for 90 s. 14. Once the 90 s have elapsed, transfer the bones to a new Petri dish containing about 40 mL of RPMI 1640 solution. This will rinse off the alcohol. Discard the Petri dish containing the ethanol. 15. Set up a new Petri dish, placing two sterile 70 mM cell strainers inside it. 16. Take a few mL of RPMI 1640 and moisten the bottom of each cell strainer in the new Petri dish. 17. Take a 10 mL syringe and fill it to capacity with the RPMI. Attach a 27.5-gauge needle to each 10 mL syringe. 18. Using your forceps, pick up each leg and hold it over one of the cell strainers. Use the scissors to cut off the ends of each bone. Flush the bone marrow cavity with RPMI 1640 using the prepared syringes. When the bone marrow has been fully removed, the bones turn bright white. 19. Take a 3 mL syringe and remove the plunger. Discard the rest of the syringe. Use the rubber end of the plunger to force the harvested bone marrow through the strainer, being forceful enough to disrupt the bone marrow plugs but gentle enough to avoid damaging the cells. Then, use the rubber plunger to mash up the bone ends contained in the other 70 mM strainer. 20. Remove the 70 mM cell strainers from the Petri dishes, and place them on top of sterile 50 mL centrifuge tubes. Use an automatic pipette to suck up all of the RPMI in the Petri dish and transfer it through the cell strainers, into the 50 mL centrifuge tubes. Serially rinse the Petri dish with RPMI 1640, leaving behind as few of the bone marrow cells as possible. 21. Spin the 50 mL tubes at 1,300 rpm for 6 min at 4°C. 22. When the centrifugation is complete, pour off the supernatant, leaving the pellet undisturbed, and add 5 mL of RBC lysis solution. Gently, flick the tube so that the pellet is dislodged and the solution is homogeneous. Let the tube sit undisturbed for 4 min at room temperature. 23. Pour the solution into a new 50-mL centrifuge tube, filtering the contents through a new 70 mM cell strainer during the

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transfer. Then, rinse the empty 50-mL centrifuge tube with ~45 mL of RPMI 1640 and pour the rinsate through the strainer and into the new tube. Now, the RBC lysis solution has been adequately diluted. The centrifuge tube used for the RBC lysis can be discarded. 24. Centrifuge the neutralized RBC lysis solution at 1,300 rpm for 6 min at 4°C. 25. At the end of the spin, pour off the supernatant and resuspend the pellet in 1 mL of PDC growth medium (see Appendix 2). 3.1.2. Cell Cultures

1. Count cells using Trypan Blue and hemocytometer. Resuspend them at a final concentration of 1.5 million cells/mL in PDC growth medium. 2. Add FLT3-L to achieve a final concentration of 100 ng/mL (see Appendix 3 for dilution and storage instructions). 3. Mix the solution well, and plate 3.5 mL of FLT3-L supplemented bone marrow cell solution per well in 6-well, flatbottom plates. Incubate at 36°C 5% CO2 incubator for 10 days. 4. On day 5, remove 1.5 mL of medium from each well and replace with 1.5 mL of fresh PDC medium, supplemented with 100 ng/mL FLT3-L. Try to remove the medium from each well without disturbing the cultures (i.e., do not mix up and down). 5. After 10 days, the culture is complete (see Subheading 3.2).

3.2. Sorting Murine Bone Marrow Cultures to Achieve Pure Populations of PDC 3.2.1. PDC Cell Sorting

The protocol is performed at 4°C unless otherwise stated. 1. On the morning of the 11th day after starting the bone marrow cultures, remove the cultures from the incubator and examine the wells through the microscope. The medium will still retain most of its original orange color. The culture will be highly cellular with many cells floating freely in the medium. Some fibroblastic growth will be evident on the bottom of the well. 2. Using a 5 mL glass pipette, carefully mix the contents of the culture around by pipetting up and down several times. Avoid generating bubbles to the greatest extent possible, as bubbles may generate surface tension and possibly damage or kill the cells. Pool the well-mixed cultures in a 50-mL centrifuge tube, filtering the culture medium through a sterile 70 mM cell strainer as it is added to the centrifuge tube. 3. Rinse the culture wells with fresh RPMI 1640 complete medium, and add to a 50 mL centrifuge tube, filtering through the 70 mM cell strainer from step 2.

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4. Centrifuge for 6 min at 1,300 rpm at 4°C. 5. Discard the supernatant and resuspend the cells in 1 mL of staining buffer. 6. Count the cells. 7. Take five aliquots of 106 unstained cells and bring them to a final volume of 150 mL in 1.5-mL microcentrifuge tubes. Label one of the tubes “isotype controls” and the others “unstained cells,” “FITC only,” “PerCP Cy 5.5 only,” and “APC only.” 8. Stain single cell suspensions in staining buffer with different combinations of mAbs by adding 1 mL of FITC antiCD11b, PerCP Cy 5.5 anti-CD11c, and APC anti-B220 per million cells to the remaining cultured cells. Also add 2 mL of each isotype control antibody to the “isotype control” tube, and 2 mL of each stain to the corresponding single stain tube. 9. Flick the tubes to mix the antibody, and place all five 1.5-mL tubes (unstained, isotype control, and single stains) as well as the 50 mL centrifuge tube on ice for 25 min. Be sure to cover the ice so that light does not bleach out any of the fluorescent labels. 10. After 25 min, add 1 mL of 1 mg/mL DAPI solution to the tubes. Flick the tube with your finger to mix the DAPI solution around. Allow the sample tubes to incubate in the dark for 5 min. 11. Centrifuge the tubes for 5 min at 1,300 rpm. 12. Discard the supernatant from each tube and resuspend the pellet in 1 mL of staining buffer. Then, pipette up and down to break up the pellet. 13. Resuspend the cells to a final concentration of 20 million cells/mL, with a minimum of 1.5 mL medium (e.g., if there are six million cells, resuspend in 1.5 mL of medium) (Fig. 1). 3.3. Harvesting and Sorting Naïve T Cells from Tea Mouse Spleens 3.3.1. Mouse Handling Procedure 3.3.2. T Cell Sorting

1. Sacrifice mice as described in Subheading 3.1, steps 1–4. Using scissors and forceps, make a horizontal incision just under the rib cage anteriorly on the left side. Identify and remove mouse spleens using a sterile technique. Place spleens in a centrifuge tube containing 15 mL RPMI 1640 complete medium supplemented with 10% FCS. 1. In a tissue culture hood, pour the medium containing the spleens into a sterile Petri dish. Then, place the spleens inside a 70 mM cell strainer and use the plunger of a 3-mL syringe to strain the spleen through the cell strainer.

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2. Place the cell strainer over a sterile 50-mL centrifuge tube and pipette the cells from the Petri dish into the 50-mL tube through the strainer. Use RPMI 1640 medium to perform serial rinses of the Petri dish, making sure to leave as few cells behind as possible. 3. Once the Petri dish is thoroughly rinsed, cap the centrifuge tube, and centrifuge at 1,400 rpm for 5 min at 4°C. Note the red pellet that forms at the bottom of the tube after centrifugation. Discard the cell strainer and the Petri dish. 4. In the tissue culture hood, pour out and discard all the supernatant, leaving the pellet undisturbed in the bottom of the centrifuge tube. 5. Add 4 mL of RBC lysis buffer per spleen to the centrifuge tube containing the pellet. Gently, flick the side of the tube until well mixed. Place on ice for 5 min. 6. Pour the mixture from step 6 into a sterile 50-mL centrifuge tube, pouring the solution through a sterile 70-mM cell strainer. This will eliminate some splenic debris from the solution. Then, add ~30 mL of RPMI 1640 complete medium supplemented with 10% FCS to neutralize the RBC lysis buffer. 7. Centrifuge at 1,400 rpm for 5 min at 4°C. 8. Discard the supernatant and resuspend the pellet in 1 mL of sterile staining buffer. 9. Add 10 mL of FITC-anti-CD25, 10 mL of APC-anti-CD4, and 10 mL of PE-anti-CD8a. Gently, flick the tube with your finger to mix the antibodies, and place all tubes on ice and protected from light for 25 min. 10. After 25 min, add 30 mL of sterile staining buffer to the centrifuge tube. 11. Pour the contents of the centrifuge tube into a sterile 50-mL centrifuge tube, passing the contents through a sterile 70-mM strainer. 12. Centrifuge at 1,400 rpm for 5 min at 4°C. 13. Pour off supernatant, leaving the gray-white pellet undisturbed. 14. Resuspend the cells in sterile staining buffer diluted to 20 million cells/mL. Filter the cells into a sterile 50-mL centrifuge tube through a 70-mM cell strainer. 3.4. Culturing Bone Marrow-Derived PDC with Naïve T Cells

1. Culture freshly sorted BALB/c BM-derived PDCs (104 cells/ well) with CD4+CD25− T cells (5 × 104 cells/well) in 96-well round-bottom plates.

3.4.1. PDC and Naïve T Cell Co-culture

2. Add anti-CD3 mAb (150 ng/mL), 10 ng/mL IL-2, and 10 ng/mL TGFb. 3. Culture cells for 96 h, with no medium changes or additional cytokines.

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1. Remove cells from culture plates. Place cells in collecting tube and add 2 mL of staining buffer. 2. Centrifuge at 2,000 rpm for 4 min at 4°C. 3. Discard the supernatant, leaving ~100 mL in the bottom of the centrifuge tube. 4. Take five aliquots of 106 unstained cells from the positive control and bring them to a final volume of 100 mL in 1.5-mL microcentrifuge tubes. Label one of the tubes “isotype control” and the others “PE-single stain,” “FITC-single stain,” “APC-single stain,” and “unstained cells.” 5. Add 1 mL of APC-anti-CD4 and 1 mL FITC-anti-CD25 mAb to each experimental sample tube, including the APC and FITC single stain controls. Flick the tube with your finger to mix the antibodies and disrupt the pellet, and place all tubes on ice for 25 min. Cover the ice so that light does not bleach out any of the fluorescent labels. 6. Add 1 mL of staining buffer and centrifuge at 2,000 rpm for 4 min at 4°C. 7. Discard the supernatant, preserving the pellet at the bottom of the tube and add 1 mL of fixation solution. Leave on ice overnight in a refrigerator. 8. The next morning, centrifuge at 2,000 rpm for 4 min at 4°C and remove supernatant. The pellet may no longer be visible. 9. Add 1 mL of ice-cold PBS to each tube. 10. Centrifuge at 2,000 rpm for 4 min at 4°C. 11. Discard the supernatant and add 1 mL of diluted permeabilization buffer. 12. Centrifuge at 2,000 rpm for 4 min at 4°C. 13. Discard the supernatant and add 1 mL of diluted permeabilization buffer. 14. Centrifuge at 2,000 rpm for 4 min at 4°C. 15. Discard the supernatant, leaving ~100 mL of liquid in each tube. Then, add 1 mL anti-Foxp3 mAb to each experimental sample tube, including the PE single stain control from step 4. Flick each tube with your finger to mix the antibody and dislodge the pellet, and place all tubes on ice for 25 min. Be sure to cover the ice so that light does not bleach out any of the fluorescent labels. 16. Add 1 mL diluted permeabilization solution. 17. Centrifuge at 1,400 rpm for 5 min at 4°C. 18. Discard the supernatant and add 1 mL of PBS. 19. Repeat steps 17 and 18. 20. Transfer samples to tubes used for flow cytometry analysis and proceed to flow cytometry (Fig. 2) (see Note 5).

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3.6. Identification of AlloantigenExpressing PDC 3.6.1. PDC-YAe FACS Staining

1. Resuspend 106 cells in 2 mL of ice-cold PBS. 2. Centrifuge at 1,400 rpm for 5 min at 4°C. 3. Discard the supernatant and block with rat serum 5% for 20 min. 4. Add 2 mL of ice-cold PBS and centrifuge at 1,400 rpm for 5 min at 4°C. 5. Discard the supernatant and block Fc receptor with antiCD16/32 (5 mg/mL) and biotin mouse IgG2b isotype (2.5 mg/mL) mAb for 20 min. 6. Add YAe mAb (2.5 mg/mL) for 30 min 4°C. 7. Add 2 mL of ice-cold PBS and centrifuge at 1,400 rpm for 5 min at 4°C. 8. Add Streptavidin–PE for 30 min 4°C. 9. Add 2 mL of ice-cold PBS and centrifuge at 1,400 rpm for 5 min at 4°C. 10. Add fluorescent-conjugated mAb to other cell surface molecules for 20 min 4°C. 11. Add 2 mL of ice-cold PBS and centrifuge at 1,400 rpm for 5 min at 4°C. 12. Add 1 mL of 1 mg/mL DAPI solution to the tube for 5 min. 13. Centrifuge the tubes for 5 min at 1,300 rpm at 4°C. 14. Discard the supernatant and resuspend the cells in 0.4 mL of staining buffer (Fig. 3).

3.6.2. PDC-YAe IHC Staining

1. Cut sections 8 mm thick in the cryostat. 2. Air-dry tissue sections for 30 min at room temperature. 3. Fix the tissue sections for 10 min in 3% paraformaldehyde at 4°C. 4. Wash three times with PBS for 5 min. 5. Block with peroxidase for 10 min. 6. Wash three times with PBS for 5 min. 7. Block slides with 5% rat serum plus anti-CD16/CD32 (5 mg/mL) and biotin mouse IgG2b isotype control (2.5 mg/ mL) for 30 min in a humidified chamber. 8. Drain off excess liquid. 9. Add biotin-YAe (2.5 mg/mL) or isotype control and incubate at room temperature for 1 h. 10. Wash three times with PBS on shaker for 5 min. 11. Add strepavidin–HRP (1:100 dilution) and incubate for 30 min at room temperature.

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12. Wash three times with PBS on shaker for 5 min. 13. Add biotinyl tyramide working solution (1:50 dilution in Amplification diluent) to each slide and incubate 3 min on shaker at room temperature. 14. Wash three times with PBS on shaker for 5 min. 15. Add Cy3-strepavidin (1/800) and incubate for 30 min at room temperature. 16. Wash three times with PBS for 5 min. 17. Add rat anti-mouse siglec H mAb (1/100) and incubate for 30 min at room temperature. 18. Wash three times with PBS for 5 min. 19. Add Cy2 anti-rat mAb (1/600) and incubate for 30 min at room temperature. 20. Wash two times with PBS and one in water for 5 min. 21. Dry the slides at room temperature and then mount slides with DAPI mounting medium. 22. Visualize slides using fluorescence microscopy (Fig. 4). 3.7. Visualization of PDC and Foxp3 Cells 3.7.1. PDC-Foxp3 IHC Staining

1. Cut sections 8 mm thick in the cryostat. 2. Air-dry tissue sections for 30 min at room temperature. 3. Fix the tissue sections for 10 min in acetone at −20°C. 4. Wash three times with PBS for 5 min. 5. Block slides with 5% rat serum plus anti-CD16/CD32 (5 mg/mL) for 30 min in a humidified chamber. 6. Wash three times with PBS for 5 min. 7. Add rat anti-mouse siglec H mAb (1/100), and incubate 30 min at room temperature. 8. Wash three times with PBS for 5 min. 9. Add Cy2 anti-rat mAb (1/600) and incubate for 30 min at room temperature. 10. Wash three times with PBS for 5 min. 11. Block the slides with blocking serum for 20 min (normal rat serum diluted 1/5 in 10% BSA/Tris 1×). 12. Add rabbit anti-mouse Foxp3 mAb 5 mg/mL in blocking serum and incubate at room temperature in humid chamber for 60 min (cover the section completely 15–50 mL). 13. Wash three times with PBS on shaker for 5 min. 14. Add Cy3 anti-rabbit antibody diluted in PBS (1/800) and incubate at room temperature in humid chamber for 30 min. 15. Wash two times with PBS for 5 min.

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16. Wash for 10 min in PBS at room temperature. 17. Dry the slides at room temperature and then mount slides with DAPI mounting medium. 18. Visualize slides using fluorescence microscopy (Fig. 5).

4. Notes 1. FCS must be endotoxin-free to prevent DC maturation. 2. In PDC processing, all tubes used during the staining or sorting purposes should be made from polypropylene to prevent PDC from adhering to the plastic. 3. After addition of permeabilization buffer, cells will be slippery. Take caution when dumping supernatant. If loss of cells becomes a problem, the supernatants may be aspirated after centrifugation. Also, the cells are fragile after permeabilization. The cell pellet should be disrupted gently. 4. Surface staining and intracellular staining may be performed consecutively; however, cells must be fixed overnight before being permeabilized. 5. Cell cultures without PDC do generate Treg in fairly high percentage numbers, but those cultures do not proliferate. The cultures only proliferate well when APCs are present in the cultures. Thus, in FACS, the percent of Treg seems to be high, but in fact the gross number is low because the cultures do not proliferate. 6. To monitor alloantigen presentation of the Ea52-68 peptide bound to the I-Ab/MHC Class II PDC with the YAe mAb, BALB/c heart allografts were transplanted into C56BL/6 recipients.

Acknowledgments We acknowledge Dr. Tamar Hermesh for technical assistance. We also wish to acknowledge the efforts of the Mount Sinai Sorting Core Facility. This work was supported by the Programa Ramón y Cajal RYC-2006-1588, Ministerio de Educación y Ciencia SAF2007-63579, Programa José Castillejo JC2008-00065, and Programa de Investigación de Grupos Emergentes del ISCIII (to JCO), NIH R01 AI-41428, AI-72039 (to JSB), and Howard Hughes Medical Institute (to EG).

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Appendix 1: 10 mL Solution of 2% Ketamine, 0.02% Acepromazine Solution

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2 mL Ketamine HCl (100 mg/mL) – Use 4 mL of 50 mg/mL ketamine HCl. 0.2 mL Acepromazine (10 mg/mL). 7.8 mL Sterile PBS.

Appendix 2: 500 mL of PDC Culture Medium

435 mL RPMI 1640 supplemented with L-Glutamine. 50 mL FCS (10%). 5 mL 1 M HEPES (10 mM). 10 mL (5,000 IU/mg) Penicillin/streptomycin (100 IU/mL penicillin and 100 mg/mL streptomycin). Filter and store at 4°C. Immediately before using, add b-mercaptoethanol (2-ME) to a concentration of 50 mM and cold filter a second time.

Appendix 3: FLT3-L Preparation

Use lyophilized FLT3-L (R&D Systems, Minneapolis, MN) to prepare 50 ng/mL aliquots: 1. Add 480 mL of PBS to 25 mg of lyophilized FLT3-L. 2. Add 20 mL of 2.5% fatty-acid-free BSA in PBS. 3. Wait 10 min for the powder to dissolve. Do not vortex to avoid bubbles. It takes about 10 min to dissolve. 4. Pipet up and down to mix the solution. Avoid adding bubbles. 5. Dispense into aliquots. 35 mL aliquot to be most efficient for 2-mouse experiments. Store at −20°C for up to 3 months.

References 1. Onai N, Obata-Onai A, Schmid MA, Ohteki T, Jarrossay D, Manz MG: Identification of clonogenic common Flt3+M-CSFR+ plasmacytoid and conventional dendritic cell progenitors in mouse bone marrow. Nat Immunol 2007, 8:1207–1216. 2. Nakano H, Yanagita M, Gunn MD: CD11c(+) B220(+)Gr-1(+) cells in mouse lymph nodes and spleen display characteristics of plasmacytoid

dendritic cells. J Exp Med 2001, 194: 1171–1178. 3. Naik SH, Sathe P, Park HY, Metcalf D, Proietto AI, Dakic A, Carotta S, O’Keeffe M, Bahlo M, Papenfuss A, et al.: Development of plasmacytoid and conventional dendritic cell subtypes from single precursor cells derived in vitro and in vivo. Nat Immunol 2007, 8:1217–1226.

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4. Omatsu Y, Iyoda T, Kimura Y, Maki A, Ishimori M, Toyama-Sorimachi N, Inaba K: Development of murine plasmacytoid dendritic cells defined by increased expression of an inhibitory NK receptor, Ly49Q. J Immunol 2005, 174:6657–6662. 5. Zhang J, Raper A, Sugita N, Hingorani R, Salio M, Palmowski MJ, Cerundolo V, Crocker PR: Characterization of Siglec-H as a novel endocytic receptor expressed on murine plasmacytoid dendritic cell precursors. Blood 2006, 107:3600–3608. 6. Blasius A, Vermi W, Krug A, Facchetti F, Cella M, Colonna M: A cell-surface molecule selectively expressed on murine natural interferon-producing cells that blocks secretion of interferon-alpha. Blood 2004, 103: 4201–4206. 7. Blasius AL, Giurisato E, Cella M, Schreiber RD, Shaw AS, Colonna M: Bone marrow stromal cell antigen 2 is a specific marker of type I IFN-producing cells in the naive mouse, but a promiscuous cell surface antigen following IFN stimulation. J Immunol 2006, 177: 3260–3265. 8. Martin P, Del Hoyo GM, Anjuere F, Arias CF, Vargas HH, Fernandez LA, Parrillas V, Ardavin C: Characterization of a new subpopulation of mouse CD8alpha+ B220+ dendritic cells endowed with type 1 interferon production capacity and tolerogenic potential. Blood 2002, 100:383–390. 9. Hadeiba H, Sato T, Habtezion A, Oderup C, Pan J, Butcher EC: CCR9 expression defines tolerogenic plasmacytoid dendritic cells able to suppress acute graft-versus-host disease. Nat Immunol 2008, 9:1253–1260. 10. Jongbloed SL, Benson RA, Nickdel MB, Garside P, McInnes IB, Brewer JM: Plasmacytoid dendritic cells regulate breach of selftolerance in autoimmune arthritis. J Immunol 2009, 182:963–968. 11. Dubois B, Joubert G, Gomez de Aguero M, Gouanvic M, Goubier A, Kaiserlian D: Sequential role of plasmacytoid dendritic cells and regulatory T cells in oral tolerance. Gastroenterology 2009, 137:1019–1028. 12. Chen X, Makala LH, Jin Y, Hopkins D, Muir A, Garge N, Podolsky RH, She JX: Type 1 diabetes patients have significantly lower frequency of plasmacytoid dendritic cells in the peripheral blood. Clin Immunol 2008, 129:413–418. 13. Hinkmann C, Knerr I, Hahn EG, Lohmann T, Seifarth CC: Reduced frequency of peripheral plasmacytoid dendritic cells in type 1 diabetes. Horm Metab Res 2008, 40:767–771.

14. Saxena V, Ondr JK, Magnusen AF, Munn DH, Katz JD: The countervailing actions of myeloid and plasmacytoid dendritic cells control autoimmune diabetes in the nonobese diabetic mouse. J Immunol 2007, 179: 5041–5053. 15. Tokita D, Sumpter TL, Raimondi G, Zahorchak AF, Wang Z, Nakao A, Mazariegos GV, Abe M, Thomson AW: Poor allostimulatory function of liver plasmacytoid DC is associated with pro-apoptotic activity, dependent on regulatory T cells. J Hepatol 2008, 49: 1008–1018. 16. Goubier A, Dubois B, Gheit H, Joubert G, Villard-Truc F, Asselin-Paturel C, Trinchieri G, Kaiserlian D: Plasmacytoid dendritic cells mediate oral tolerance. Immunity 2008, 29: 464–475. 17. Manches O, Munn D, Fallahi A, Lifson J, Chaperot L, Plumas J, Bhardwaj N: HIVactivated human plasmacytoid DCs induce Tregs through an indoleamine 2,3-dioxygenase-dependent mechanism. J Clin Invest 2008, 118:3431–3439. 18. Puccetti P, Fallarino F: Generation of T cell regulatory activity by plasmacytoid dendritic cells and tryptophan catabolism. Blood Cells Mol Dis 2008, 40:101–105. 19. Wang L, Pino-Lagos K, de Vries VC, Guleria I, Sayegh MH, Noelle RJ: Programmed death 1 ligand signaling regulates the generation of adaptive Foxp3+CD4+ regulatory T cells. Proc Natl Acad Sci U S A 2008, 105:9331–9336. 20. Tokita D, Mazariegos GV, Zahorchak AF, Chien N, Abe M, Raimondi G, Thomson AW: High PD-L1/CD86 ratio on plasmacytoid dendritic cells correlates with elevated T-regulatory cells in liver transplant tolerance. Transplantation 2008, 85:369–377. 21. Ito T, Yang M, Wang YH, Lande R, Gregorio J, Perng OA, Qin XF, Liu YJ, Gilliet M: Plasmacytoid dendritic cells prime IL-10-producing T regulatory cells by inducible costimulator ligand. J Exp Med 2007, 204:105–115. 22. Munn DH, Sharma MD, Hou D, Baban B, Lee JR, Antonia SJ, Messina JL, Chandler P, Koni PA, Mellor AL: Expression of indoleamine 2,3-dioxygenase by plasmacytoid dendritic cells in tumor-draining lymph nodes. J Clin Invest 2004, 114:280–290. 23. Sharma MD, Baban B, Chandler P, Hou DY, Singh N, Yagita H, Azuma M, Blazar BR, Mellor AL, Munn DH: Plasmacytoid dendritic cells from mouse tumor-draining lymph nodes directly activate mature Tregs via indoleamine 2,3-dioxygenase. J Clin Invest 2007, 117: 2570–2582.

Plasmacytoid Dendritic Cells in Tolerance 24. Sapoznikov A, Fischer JA, Zaft T, Krauthgamer R, Dzionek A, Jung S: Organ-dependent in vivo priming of naive CD4+, but not CD8+, T cells by plasmacytoid dendritic cells. J Exp Med 2007, 204:1923–1933. 25. Pepper M, Dzierszinski F, Wilson E, Tait E, Fang Q, Yarovinsky F, Laufer TM, Roos D, Hunter CA: Plasmacytoid dendritic cells are activated by Toxoplasma gondii to present antigen and produce cytokines. J Immunol 2008, 180:6229–6236. 26. Koyama M, Hashimoto D, Aoyama K, Matsuoka K, Karube K, Niiro H, Harada M, Tanimoto M, Akashi K, Teshima T: Plasmacytoid dendritic cells prime alloreactive T cells to mediate graft-versus-host disease as antigenpresenting cells. Blood 2009, 113:2088–2095. 27. Sadaka C, Marloie-Provost MA, Soumelis V, Benaroch P: Developmental regulation of MHC II expression and transport in human plasmacytoid-derived dendritic cells. Blood 2009, 113:2127–2135. 28. Young LJ, Wilson NS, Schnorrer P, Proietto A, ten Broeke T, Matsuki Y, Mount AM, Belz GT, O’Keeffe M, Ohmura-Hoshino M, et al.: Differential MHC class II synthesis and ubiquitination confers distinct antigenpresenting properties on conventional and plasmacytoid dendritic cells. Nat Immunol 2008, 9:1244–1252. 29. Darrasse-Jeze G, Deroubaix S, Mouquet H, Victora GD, Eisenreich T, Yao KH, Masilamani RF, Dustin ML, Rudensky A, Liu K, et al.: Feedback control of regulatory T cell homeostasis by dendritic cells in vivo. J Exp Med 2009, 206:1853–1862. 30. Ochando JC, Homma C, Yang Y, Hidalgo A, Garin A, Tacke F, Angeli V, Li Y, Boros P,

31.

32.

33.

34.

35.

36.

147

Ding Y, et al.: Alloantigen-presenting plasmacytoid dendritic cells mediate tolerance to vascularized grafts. Nat Immunol 2006, 7:652–662. Maraskovsky E, Brasel K, Teepe M, Roux ER, Lyman SD, Shortman K, McKenna HJ: Dramatic increase in the numbers of functionally mature dendritic cells in Flt3 ligandtreated mice: multiple dendritic cell subpopulations identified. J Exp Med 1996, 184:1953–1962. Maraskovsky E, Daro E, Roux E, Teepe M, Maliszewski CR, Hoek J, Caron D, Lebsack ME, McKenna HJ: In vivo generation of human dendritic cell subsets by Flt3 ligand. Blood 2000, 96:878–884. Brasel K, De Smedt T, Smith JL, Maliszewski CR: Generation of murine dendritic cells from flt3-ligand-supplemented bone marrow cultures. Blood 2000, 96:3029–3039. Gilliet M, Boonstra A, Paturel C, Antonenko S, Xu XL, Trinchieri G, O’Garra A, Liu YJ: The development of murine plasmacytoid dendritic cell precursors is differentially regulated by FLT3-ligand and granulocyte/macrophage colony-stimulating factor. J Exp Med 2002, 195:953–958. Brawand P, Fitzpatrick DR, Greenfield BW, Brasel K, Maliszewski CR, De Smedt T: Murine plasmacytoid pre-dendritic cells generated from Flt3 ligand-supplemented bone marrow cultures are immature APCs. J Immunol 2002, 169:6711–6719. Swee LK, Bosco N, Malissen B, Ceredig R, Rolink A: Expansion of peripheral naturally occurring T regulatory cells by Fms-like tyrosine kinase 3 ligand treatment. Blood 2009, 113: 6277–6287.

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Chapter 10 In Vitro-Generated DC with Tolerogenic Functions: Perspectives for In Vivo Cellular Therapy Cees van Kooten and Kyra A. Gelderman Abstract Dendritic cells (DCs) have a central role in immune regulation and serve as an essential link between innate and adaptive immunity. Their broad range of powerful immune stimulatory as well as regulatory functions has made DCs a target for vaccine development strategies. One approach to promote the tolerogenicity of DCs is to suppress their maturation by pharmacological agents, including the glucocorticoid dexamethasone. In this chapter, we describe methods to generate tolerogenic Dex-DC derived from either human peripheral blood monocytes or rat bone marrow cells. Key words: Dendritic cells, Tolerance, Dexamethasone, Human, Rat

1. Introduction Dendritic cells (DCs) are bone marrow-derived cells that have a central role in immune regulation, ranging from tolerance induction and the prevention of autoimmunity to the induction of antitumor immunity and the protection against infectious agents (1, 2). This broad range of powerful immune stimulatory as well as regulatory functions has made DCs a target for vaccine development strategies. These include cellular vaccination for treatment of cancer or infectious diseases, as well as “negative vaccination” for the treatment of autoimmune diseases and prevention of allograft rejection. The latter should be accomplished by inhibiting the immunostimulatory capacity of DCs or, more importantly, by exploiting tolerogenic DCs to silence specific immune responses.

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_10, © Springer Science+Business Media, LLC 2011

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The functional characteristics of DCs are determined by at least two different factors. First, different subsets of DCs can be recognized which have specific and specialized functions, such as the IFN-a production by plasmacytoid DCs. Second, DCs exist in different activation stages that for a large part determine their functional capacities. In a nonactivated, immature stage, DCs are specialized in antigen uptake, whereas activation results in a shift towards the capacity to present antigen and to activate T cells. Overall this process is indicated by a single term “maturation,” but should be considered as a collection of different processes which, based on the activation status, determine the strength and the quality of the immune response (3). It has been shown both in vitro and in vivo that DCs in an immature or partially activated (semimature) form have tolerogenic functions (4, 5). Based on this, tolerogenic DCs have the potential to be used in “negative vaccination” strategies to suppress specific immune responses like in autoimmunity or transplantation. However, controlling the maturation stage of the DC is of critical importance, not only to promote the tolerogenic capacity but also to prevent the risk of further activation after administration, which could shift the cells toward a more immunogenic function. Several strategies have recently been described to suppress DC maturation, including the use of genetic engineering, anti-inflammatory cytokines (IL-10, TGF-b), immunosuppressive drugs, and other pharmacological agents (i.e., rapamycin, vitamin D3, and dexamethasone) (6–9). Several groups have described the strong immune-modulating capacity of glucocortocoids, including dexamethasone (Dex) (10). Dex interferes with the development of immunocompetent monocyte-derived DC. Upon stimulation of Dex-DCs, these cells are strongly hampered in their upregulation of costimulatory and MHC molecules (11, 12), and show a reduced production of proinflammatory cytokines, including IL-6, TNF-a, IL-1b, and IL-12 (11–14). Importantly, the same conditions result in an increased production of the anti-inflammatory cytokine IL-10. Similar effects have been described using mouse or rat bone-marrow derived DC (10). As a consequence, Dex-DCs are poor stimulators of T cells, resulting in T cell anergy and/or the induction of regulatory T cells (15, 16). More importantly, using several experimental models of transplantation, it has been shown that Dex-DCs have the potential to modulate the recipient immune response and to prolong allograft survival (17–19). In this chapter, we describe the procedures to generate DexDCs derived from either human peripheral blood monocytes or rat bone marrow cells.

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2. Materials 2.1. Isolation of Human Monocytes

1. Single buffy coat of healthy blood donor (obtained with informed consent from local blood bank; Sanquin, Leiden, The Netherlands). 2. Phosphate-buffered saline (PBS); sterile. 3. Sterile centrifuge tubes, 50 and 15 ml (Greiner – Bio One). 4. Ficoll-Isopaque solution, density 1.077 (local pharmacy, LUMC). 5. Temperature-controlled centrifuge (Spinchron DLX; Beckman Coulter, Woerden, The Netherlands). 6. Inverted microscope (Zeiss, Sliedrecht, The Netherlands). 7. MACS separation buffer (PBS, 0.5% bovine serum albumin BSA (Sigma), 2 mM EDTA) 4°C. 8. CD14 microbeads (Myltenyi Biotech GmBH, Bergisch Gladbach, Germany). 9. LS+ columns (Myltenyi Biotech GmBH). 10. QuadroMACS (Myltenyi Biotech GmBH).

2.2. Culture of Human Tolerogenic Dendritic Cells

1. RPMI 1640 (Gibco Invitrogen, Breda, The Netherlands). 2. Heat inactivated Fetal Calf Serum (DFCS; BioWhittaker, Vervier, Belgium). 3. Penicillin/streptomycin solution (Gibco Invitrogen). 4. Fungizone (optional; Gibco Invitrogen). 5. Recombinant human GM-CSF (Biosource Invitrogen). 6. Recombinant human IL-4 (Biosource Invitrogen). 7. Dexamethasone (local pharmacy). 8. Trypsin EDTA solution (Sigma, Zwijndrecht, The Netherlands). 9. Cell scraper (Costar, Corning, NY). 10. 6-well culture plates (Costar). 11. Incubator (Thermo Scientific).

2.3. Culture of Rat Tolerogenic Dendritic Cells

1. Tibias and femurs from rats. 2. Surgical scissor/bone cutting scissors, forceps. 3. Erythrocyte lysis buffer (155 mM NH4Cl and 10 mM KHCO3). 4. 70-mm cell strainer (BD Falcon, Breda, The Netherlands). 5. Culture medium RPMI 1640 (Gibco Invitrogen) supplemented with 10% heat-inactivated DFCS (BioWhittaker), pen/ strep (Gibco, Invitrogen), fungizone (Gibco, Invitrogen),

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b-mercaptoethanol (50 mM; Merck, Darmstadt, Germany), and L-glutamine (2 mM; Gibco, Invitrogen). 6. Growth factors: Recombinant rat GM-CSF (Biosource Invitrogen), recombinant rat IL-4 (Biosource Invitrogen), recombinant human Flt3L (Amgen, Seattle, WA). 7. 6-well culture plates (Costar). 8. Centrifuge (Spinchron DLX; Beckman Coulter). 9. Incubator (Thermo Scientific).

3. Methods 3.1. Isolation of Human Monocytes

1. Monocytes are purified from human mononuclear cells (MNC) isolated from single buffy coats obtained from healthy blood donors. 2. All procedures are performed under sterile conditions in a flow cabinet and with sterile reagents. 3. Divide the buffy coat over four tubes and add PBS to a final volume of 50 ml. 4. Divide each tube over two tubes, making a total of eight tubes. 5. Carefully layer 12 ml of Ficoll solution under these cells using a 10-ml sterile plastic pipet. 6. Centrifuge the tubes at room temperature for 20 min at 500g without acceleration and without brake. 7. Carefully harvest the mononuclear cells from the interphase using a sterile pipet and transfer them to a new 50-ml tube (two interphases/tube). 8. Add PBS until a final volume of 50 ml. 9. Centrifuge for 10 min at 500g to remove Ficoll. High speed is required to prevent cell loss, since the remaining Ficoll increases the density of the solution. Now, centrifuge brake and acceleration can be switched on. 10. Discard supernatant, resuspend the cell pellet, and add PBS until a volume of 50 ml. 11. Centrifuge for 10 min at 100g to remove thrombocytes. This low speed is essential since this separation is based on the relatively low density of thrombocytes. 12. Repeat steps 10–11 until the supernatant is clear (mostly 2–3 times). 13. Pool the cells and count to determine the total number.

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14. Continue with the purification of monocytes. Depending on the planned experiments, continue with all isolated cells or take a lower number (see Note 1). 15. Centrifuge the cells at 200g for 8 min, remove the supernatant completely, and resuspend the cells in a concentration of 107 MNC in 95 ml of MACS buffer. 16. Add 5 ml of CD14 Microbeads per 107 MNC. 17. Mix well and incubate for 15 min in refrigerator (4–10°C). 18. Wash the cells by adding at least a 20-fold excess of MACS buffer. 19. Centrifuge for 8 min at 200g. 20. Discard the supernatant and repeat steps 18–19. 21. Remove supernatant completely and resuspend the cell pellet in MACS buffer at a concentration of 108 cells/500 ml. 22. Place LS+ column(s) in the MACS separator and rinse it with 3 ml of MACS buffer. 23. Apply the cell suspension onto the column(s) and let the negative cells pass through. The maximum amount of cells that can be applied on one column is 2 × 109 total cells or 108 monocytes. Typically, we use two LS+ columns for one single buffy coat. 24. Rinse each column three times with 3 ml of MACS buffer. 25. Remove the column from the separator and place it on a 15-ml tube. 26. Pipette 5 ml of buffer on to the column and flush out the cells using the plunger supplied with the column. 27. Centrifuge for 8 min at 200g. 28. Remove all supernatant and resuspend the cells in 10 ml of culture medium (see Subheading 3.2). 29. Count the cells. At this stage, purified CD14+ monocytes (typically over 90% pure) are directly used for cell culture; alternatively, they may be frozen down. In our experience, cell yields were higher when starting DC cultures with fresh cells (see Note 2). 3.2. Culture of Human Tolerogenic Dendritic Cells

1. Purified CD14+ monocytes are brought to a concentration of 0.75 × 106 cells/ml in culture medium, consisting of RPMI 1640, 10% heat-inactivated DFCS, Pen/Strep, human IL-4 (10 ng/ml), and human GM-CSF (5 ng/ml) (see Note 3). 2. For the generation of tolerogenic DC (Dex-DC), dexamethasone (final concentration 10−6 M) is added to the medium from the start of the culture (see Notes 4 and 5).

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3. Cells are seeded in 6-well plates, with 2 ml/well (final amount of cells 1.5 × 106/well). 4. Put cells in 37°C CO2 incubator. 5. At day 2/3, 1 ml of culture medium is added to every well. In this case, growth factors and dexamethasone are included 3× concentrated (30 ng/ml IL-4, 15 ng/ml GM-CSF, and 3 × 10−6 M dex) (see Note 6). 6. At day 4/5, 1 ml of culture medium is carefully removed from the top of every well. Next, 1 ml of culture medium is added as in step 5. 7. After day 6, monocytes get fully differentiated into tolerogenic Dex-DC orcontrol immature monocyte-derived DC in the case that dex is omitted in steps 2, 5, and 6 (see Note 7). 8. Culture of cells can be further prolonged, but then, at day 6/7, step 6 should be repeated. 9. To obtain tolerogenic Dex-DC for experiments, cells are harvested from culture plates and used for experiments. Since Dex-DCs are more adherent than normal immature moDC, cells should be harvested by brief trypsinization and/or scraping with a cell scraper. 10. For trypsinization, supernatants and nonadherent cells are harvested and collected in a 50-ml tube. 11. Plates are washed with PBS. 12. Plates are incubated at 37°C with 200 ml/well trypsin solution and monitored occasionally through the microscope for detachment of cells. When they start to round up (after a few minutes), cells are harvested by pipetting or gentle scraping. 13. Cells are collected in a 50-ml tube. 14. Trypsin is inactivated by the addition of equal volume RPMI 1640 with 10% DFCS. 15. Centrifuge for 8 min at 200g, together with the cells from step 10. 16. Discard the supernatant. 17. Resuspend the pellets in culture medium, pool the samples, and count the cells using Trypan blue staining to determine the viable cell count (see Notes 8 and 9). 3.3. Culture of Rat Tolerogenic Dendritic Cells

1. Bone Marrow (BM) cells are obtained from tibia and femur of adult (>8 weeks) rats. We have successfully generated tolerogenic Dex-DC from all rat strains investigated, including Brown Norway (BN), Albino Oxford (AO), Dark Agouti (DA), and Lewis (LEW) (see Note 10). 2. Whole femurs and tibias are obtained and kept in PBS until isolation of BM.

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3. Bone ends are cut off and bones are flushed with RPMI++ (RPMI 1640 with FCS and pen/strep) using a syringe with a 23-G needle. 4. BM cells are collected in a 70-mm mesh sieve placed on a 50-ml tube and gently pushed through with the plunger of a syringe. Rinse with RPMI++ to collect all cells in the tube. 5. Centrifuge for 8 min at 200g. 6. Resuspend the cell pellet in 10 ml of lysis buffer to remove red blood cells. 7. Centrifuge for 8 min at 200g and resuspend the pellet in medium. 8. Count the viable cells. Typically, one rat gives a cell yield of 108 BM cells. 9. BM cells can directly be used for cell culture or frozen down in aliquots in liquid nitrogen, depending on the required number of (tolerogenic) DC. 10. Cells are brought to a concentration of 0.5 × 106 cells/ml and supplemented with growth factors rat GM-CSF (2 ng/ml), rat IL-4 (5 ng/ml), and human Flt3L (50 ng/ml). 11. Cells are cultured in 6-well plates, with 3 ml of cell suspension per well, in a 37°C CO2 incubator. 12. At day 2, culture medium is carefully harvested, supernatant centrifuged, and remaining cells are returned in culture. Medium is replaced by 3 ml of fresh culture medium, including growth factors. 13. At day 4, the procedure of step 13 is repeated. 14. For the generation of tolerogenic DC, dexamethasone is added at day 4 to the cultures at a final concentration of 10−6 M (see Note 11). 15. At day 7, nonadherent and semiadherent cells are harvested and collected (see Note 12). 16. Count the cells and determine the viable cell number. Cells can be used for functional characterization (see Note 13).

4. Notes 1. Positive selection using microbeads is a widely used method to purify peripheral blood monocytes. However, also other methods of purification can be used, including selection through negative selection, elutriation, selection using Percoll density gradients or plastic adherence. 2. This population of purified monocytes can also be used for the generation of other myeloid populations, including

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macrophage subsets. It has become clear that monocytes have an enormous flexibility in the differentiation towards specific cell subsets (20, 21). By the use of different growth factors, defined cell populations can be generated from the same source, allowing a direct comparison of these subsets. However, this flexibility also imposes that culture conditions have to be followed strictly to prevent unwanted variations. 3. The cell density of plated monocytes directly affects the differentiation towards mo-DC. We have observed that at higher cell density, phenotypic differentiation toward DC (loss of CD14 expression) is hampered. Therefore, we prefer starting the cultures with purified monocytes over selection by plastic adherence, since the latter will result in a variable cell density. 4. Routinely, we use 1 mM of dexamethasone for the generation of Dex-DC. Dose response experiments have shown that inhibition of differentiation is maximal at 100 nM. We have seen that the same inhibition can be obtained by other glucocorticoids including prednisolon. 5. Routinely, we add dexamethasone at the beginning of the culture and maintain its presence until cells are harvested at day 7. We have previously demonstrated that exposure during the first 48 hours is sufficient to generate the regulatory phenotype (12). Others have demonstrated that dexamethasone has a regulatory function when added to already differentiated DC, although there have been some conflicting results (11, 14, 22, 23). 6. Based on the original protocol, we still follow the procedure of medium refreshment at days 2/3 and 4/5. However, it has been shown that when lower cell concentrations are used, growth factors added at the start of culture should be sufficient. This is an advantage for protocols where mo-DC are generated for clinical purposes, and where reduced handling reduces risk of contamination (24–26). 7. This protocol focuses on the use of dexamethasone for the generation of tolerogenic DCs. As mentioned in the introduction, several other agents have been identified that have tolerogenic potential, like IL-10 or VitD3 (9). In many cases, protocols to generate other tolerogenic DCs can be easily adapted from the present protocol, allowing a direct comparison between the different cells. 8. Dex-DCs retain a monocyte/macrophage-like phenotype with CD14 expression and absence of the DC marker CD1a. However, these cells also lack expression of the typical macrophage marker CD68, and do express the DC marker DC-SIGN (15).

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9. In most cases, it is advised that the phenotype and function of the cells are controlled. Since dexamethasone affects many aspects of DC biology, a full characterization including phenotype, cytokine production, and T cell stimulatory capacity will not always be possible. For us, the primary readout for the successful generation of tolerogenic Dex-DC is the complete absence of IL-12 production (both IL-12p70 as well as IL-12p40) upon activation by LPS or CD40L. 10. Tolerogenic Dex-DCs have also been generated from mouse bone marrow (17, 27, 28) based on established protocols to generate mouse BM-DC. In many cases, only GM-CSF is used as a growth factor, although also combination with IL-4 or with TNF have been described. The use of dexamethasone is comparable to the method described for rat DCs. 11. For the generation of rat tolerogenic Dex-DC, dexamethasone is added at day 4 of the bone marrow cultures. Addition at day 0, comparable to the monocyte system described in Subheading 3.2, resulted in low cell yields. This is in line with our observations using human CD34+ precursor cells from cord blood, where early addition of dexamethasone resulted in a specific apoptosis induction (29). 12. At this stage, we ignore the adherent cells, which may be macrophage-like cells. It has been shown that adherent cells in comparable rat bone marrow cultures also have regulatory functions and have been used to prolong allograft survival (30). 13. For the validation of the successful generation of tolerogenic rat DC, we have also used the absence of production of IL-12 upon activation. It is important to note that for rat BMDC, production of IL-12p70 is very low and at the limit of what is detectable by available ELISA system. We, therefore, prefer the measurement of rat IL-12p40 (31).

Acknowledgments These studies have been supported by grants from the Dutch Kidney Foundation, NWO, and the EU (FP6-RISET). References 1. Banchereau, J. and Steinman, R. M. (1998) Dendritic cells and the control of immunity, Nature 392, 245–252. 2. Steinman, R. M. and Banchereau, J. (2007) Taking dendritic cells into medicine, Nature 449, 419–426.

3. Reis, E. and Sousa, C. (2006) Dendritic cells in a mature age, Nat. Rev. Immunol. 6, 476–483. 4. Lutz, M. and Schuler, G. (2002) Immature, semi-mature and fully mature dendritic cells: which signals induce tolerance or immunity? Trends Immunol. 23, 445.

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5. Steinman, R. M., Hawiger, D., and Nussenzweig, M. C. (2003) Tolerogenic dendritic cells, Annu. Rev. Immunol. 21, 685–711. 6. Hackstein, H. and Thomson, A. W. (2004) Dendritic cells: emerging pharmacological targets of immunosuppressive drugs, Nat. Rev. Immunol. 4, 24–35. 7. Woltman, A. M. and van Kooten, C. (2003) Functional modulation of dendritic cells to suppress adaptive immune responses, J. Leukoc. Biol. 73, 428–441. 8. Adorini, L., Giarratana, N., and Penna, G. (2004) Pharmacological induction of tolerogenic dendritic cells and regulatory T cells, Semin. Immunol. 16, 127–134. 9. Morelli, A. E. and Thomson, A. W. (2007) Tolerogenic dendritic cells and the quest for transplant tolerance, Nat. Rev. Immunol. 7, 610–621. 10. van Kooten, C., Stax, A. S., Woltman, A. M., and Gelderman, K. A. (2009) Handbook of experimental pharmacology “dendritic cells”: the use of dexamethasone in the induction of tolerogenic DCs, Handb. Exp. Pharmacol. 188, 233–249. 11. Piemonti, L., Monti, P., Allavena, P., Sironi, M., Soldini, L., Leone, B. E., Socci, C., and Di Catlo, V. (1999) Glucocorticoids affect human dendritic cell differentiation and maturation, J. Immunol. 162, 6473–6481. 12. Woltman, A. M., De Fijter, J. W., Kamerling, S. W., Paul, L. C., Daha, M. R., and van Kooten, C. (2000) The effect of calcineurin inhibitors and corticosteroids on the differentiation of human dendritic cells, Eur. J. Immunol. 30, 1807–1812. 13. Vieira, P. L., Kalinski, P., Wierenga, E. A., Kapsenberg, M. L., and de Jong, E. C. (1998) Glucocorticoids inhibit bioactive IL-12p70 production by in vitro-generated human dendritic cells without affecting their T cell stimulatory potential, J. Immunol. 161, 5245–5251. 14. Rea, D., van Kooten, C., Van Meijgaarden, K. E., Melief, C. J. M., and Offringa, R. (2000) Glucocorticoids transform CD40triggering of dendritic cells into an alternative activation pathway resulting in antigen presenting cells that secrete IL-10, Blood 95, 3162–3167. 15. Woltman, A. M., Van der Kooij, S. W., De Fijter, J. W., and van Kooten, C. (2006) Maturation-resistant dendritic cells induce hyporesponsiveness in alloreactive CD45RA+ and CD45RO+ T-cell populations, Am. J. Transplant. 6, 2580–2591.

16. Fazekasova, H., Golshayan, D., Read, J., Tsallios, A., Tsang, J. Y., Dorling, A., George, A. J., Lechler, R. I., Lombardi, G., and Mirenda, V. (2009) Regulation of rat and human T-cell immune response by pharmacologically modified dendritic cells, Transplantation 87, 1617–1628. 17. Roelen, D. L., Schuurhuis, D. H., van den Boogaardt, D. E., Koekkoek, K., van Miert, P. P., van Schip, J. J., Laban, S., Rea, D., Melief, C. J., Offringa, R., Ossendorp, F., and Claas, F. H. (2003) Prolongation of skin graft survival by modulation of the alloimmune response with alternatively activated dendritic cells, Transplantation 76, 1608–1615. 18. Mirenda, V., Berton, I., Read, J., Cook, T., Smith, J., Dorling, A., and Lechler, R. I. (2004) Modified dendritic cells coexpressing self and allogeneic major histocompatability complex molecules: an efficient way to induce indirect pathway regulation, J. Am. Soc. Nephrol. 15, 987–997. 19. Stax, A. M., Gelderman, K. A., Schlagwein, N., Essers, M. C., Kamerling, S. W., Woltman, A. M., and van Kooten, C. (2008) Induction of donor-specific T-cell hyporesponsiveness using dexamethasone-treated dendritic cells in two fully mismatched rat kidney transplantation models, Transplantation 86, 1275–1282. 20. Gordon, S. and Taylor, P. R. (2005) Monocyte and macrophage heterogeneity, Nat. Rev. Immunol. 5, 953–964. 21. Mosser, D. M. and Edwards, J. P. (2008) Exploring the full spectrum of macrophage activation, Nat. Rev. Immunol. 8, 958–969. 22. Vanderheyde, N., Verhasselt, V., Goldman, M., and Willems, F. (1999) Inhibition of human dendritic cell functions by methylprednisolone, Transplantation 67, 1342–1347. 23. de Jong, E. C., Vieira, P. L., Kalinski, P., and Kapsenberg, M. L. (1999) Corticosteroids inhibit the production of inflammatory mediators in immature monocyte-derived DC and induce the development of tolerogenic DC3, J. Leukoc. Biol. 66, 201–204. 24. de Vries, I, Lesterhuis, W. J., Scharenborg, N. M., Engelen, L. P., Ruiter, D. J., Gerritsen, M. J., Croockewit, S., Britten, C. M., Torensma, R., Adema, G. J., Figdor, C. G., and Punt, C. J. (2003) Maturation of dendritic cells is a prerequisite for inducing immune responses in advanced melanoma patients, Clin. Cancer Res. 9, 5091–5100. 25. Palucka, A. K., Ueno, H., Connolly, J., Kerneis-Norvell, F., Blanck, J. P., Johnston, D. A., Fay, J., and Banchereau, J. (2006)

In Vitro-Generated DC with Tolerogenic Functions Dendritic cells loaded with killed allogeneic melanoma cells can induce objective clinical responses and MART-1 specific CD8+ T-cell immunity, J. Immunother. 29, 545–557. 26. Nestle, F. O., Alijagic, S., Gilliet, M., Sun, Y., Grabbe, S., Dummer, R., Burg, G., and Schadendorf, D. (1998) Vaccination of melanoma patients with peptide- or tumor lysatepulsed dendritic cells, Nat. Med. 4, 328–332. 27. Emmer, P. M., van der Vlag, J., Adema, G. J., and Hilbrands, L. B. (2006) Dendritic cells activated by lipopolysaccharide after dexamethasone treatment induce donor-specific allograft hyporesponsiveness, Transplantation 81, 1451–1459. 28. van Duivenvoorde, L. M., Han, W. G., Bakker, A. M., Louis-Plence, P., Charbonnier, L. M., Apparailly, F., van der Voort, E. I., Jorgensen, C., Huizinga, T. W., and Toes, R. E. (2007) Immunomodulatory dendritic cells inhibit

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Th1 responses and arthritis via different mechanisms, J. Immunol. 179, 1506–1515. 29. Woltman, A. M., Massacrier, C., De Fijter, J. W., Caux, C., and van Kooten, C. (2002) Corticosteroids prevent generation of CD34+derived dermal dendritic cells but do not inhibit Langerhans cell development, J. Immunol. 168, 6181–6188. 30. Peche, H., Trinite, B., Martinet, B., and Cuturi, M. C. (2005) Prolongation of heart allograft survival by immature dendritic cells generated from recipient type bone marrow progenitors, Am. J. Transplant. 5, 255–267. 31. Stax, A. M., Crul, C., Kamerling, S. W., Schlagwein, N., van der Geest, R. N., Woltman, A. M., and van Kooten, C. (2008) CD40L stimulation of rat dendritic cells specifically favors the IL-12/IL-10 ratio resulting in a strong T cell stimulatory capacity, Mol. Immunol. 45, 2641–2650.

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Chapter 11 Preparation of Mouse Bone Marrow-Derived Dendritic Cells with Immunoregulatory Properties Mercedes Segovia, Maria Cristina Cuturi, and Marcelo Hill Abstract Tolerogenic dendritic cells (Tol-DCs) are critical players in physiological tolerance. Moreover, they also play a role in immune regulation both in a pathophysiological context and when used as therapeutic tools in cell therapy strategies. Here, we describe a protocol to differentiate murine Tol-DCs from bone marrow precursors in vitro. Importantly, Tol-DCs actively suppress T cells stimulated with immunogenic allogeneic DCs. Indeed, Tol-DCs generated using this approach can be useful in studying and characterising the immunoregulatory pathways of Tol-DC and in developing Tol-DC-based cell therapy protocols using in vivo models. Key words: Tolerogenic dendritic cells, Mixed leukocyte reaction (MLR), Immunoregulation, Bone marrow-derived DCs

1. Introduction Dendritic cells (DCs) are a heterogeneous population of leukocytes that orchestrate the immune response by playing a critical role at the interface between natural and adaptive immunity. In recent years, it has been proposed that DCs not only control immunity but also maintain tolerance to self-antigens (1). Indeed, tolerogenic DCs (Tol-DC) play a role in both central and peripheral tolerance mechanisms. Moreover, DCs can be manipulated by malignant cells or pathogens to avoid immune-mediated elimination. The characterisation of immune regulation by Tol-DCs, therefore, will lead to a better understanding of both self-tolerance and pathologically acquired tolerance. Moreover, the characterisation of the immunoregulatory mechanisms used by Tol-DCs could guide the development and rationalisation of immune intervention Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_11, © Springer Science+Business Media, LLC 2011

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strategies. Tol-DCs and their molecular immunoregulatory network could be targeted using therapeutic approaches. Interestingly, Tol-DCs can also be used as tools to manipulate immune responses. Thus, cell therapy using Tol-DCs or even regulatory T cells (Tregs) is a promising strategy to manipulate antigen-specific immune responses in the context of organ transplantation and autoimmunity (2, 3), as well as in immune-mediated elimination of gene therapy vectors/products. Rodent models are needed to improve protocols and adjuvant treatments when using cell therapybased protocols. Here, we describe how to generate mouse bone marrow-derived Tol-DCs with immunoregulatory properties.

2. Materials 1. RPMI medium supplemented with 10% FCS, 0.05 mM 2-ME, 2 mM L-glutamine, 1 mM sodium pyruvate, 1% HEPES buffer, 100 U/mL penicillin, and 0.1 mg/mL streptomycin (see Note 1). 2. Culture supernatant from murine GM-CSF-transfected COS cells. 3. 6- to 10-week-old male mice: two Balb/c and two C56/B6. 4. Bacteriological 10-cm Petri dishes (see Note 2). 5. PBS supplemented with 7 mM EDTA and 2% FCS (PES). 6. 0.2 mm-filtered red cell lysis solution (RCLS): 0.15 M NH4Cl, 0.01 M KHCO3 and 100 mM Na2EDTA, pH 7.4. 7. 5-mL syringe and 271/2 G needle. 8. Collagenase D (Boehringer Mannheim) (2 mg/mL) in RPMI. 9. 14.5% Nycodenz (Nycomed Pharma) in PES. 10. EDTA/FCS: 1 mL 0.1 M EDTA and 9 mL FCS. 11. 96 round-well sterile plates. 12. Sterile scissors and forceps.

3. Methods 3.1. Differentiation of DCs from Bone Marrow Precursors

We assayed different doses of GM-CSF during the differentiation period. In addition, the number of DCs was assessed at different time points for each dose (Fig. 1). The phenotype (Table 1) and suppressive capacity (Fig. 2) of the BMDCs were studied using cells harvested following 8 days of differentiation in culture.

total฀number฀of฀cells฀(x฀10 6)/dish

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GM-CSF฀ concentration฀

1.5

adherents adherents suspension suspension

1.0

0.4 ng/ml 0.1 ng/ml 0.4 ng/ml 0.1 ng/ml

0.5

0.0 d5

d6

d7

d8

Fig. 1. Cell quantification. Bone marrow cells were differentiated with the indicated doses of GM-CSF. Adherent cells and cells in suspension were harvested at the indicated days and counted. The mean ± SD of three different experiments is shown.

Table 1 BMDC phenotype at day 8 GM-CSF Supernatant cells

Adherent cells

Marker

0.4 ng/ml (%)

0.1 ng/ml (%)

0.4 ng/ml (%)

0.1 ng/ml (%)

CD80

75.5 ± 14.5

64 ± 19

81 ± 16.5

74 ± 14

CD 86

65.6 ± 14.5

52 ± 19.9

61.5 ± 10.6

48.5 ± 23

CD11c

77.5 ± 16.7

64 ± 24

79.3 ± 3

79.5 ± 10.5

MHC II

62.6 ± 12.4

54 ± 18

57.3 ± 10.9

55 ± 9.5

The phenotypes of the adherent cells and the cells in suspension cultured for 8 days in the presence of the indicated doses of GM-CSF are shown. The values represent the percentage (mean ± SD) of cells positive for the indicated markers from three independent experiments.

3.1.1. Day 0

1. Sacrifice one C57/B6 mouse by cervical dislocation (see Note 3). 2. Clean the leg bones (femurs and tibias) from the muscles and other tissues and put them into tubes containing 70% ethanol. After 10 min, thoroughly rinse away the ethanol with sterile PBS. 3. Put the bones onto 10-cm Petri dishes and cover them with about 20 mL of PES. Cut each bone into two parts using the sterile scissors and forceps. Using a 5-mL syringe and a 271/2 G needle, flush out the bone marrow. Remove the red matter from the bones and solubilise it with the syringe (see Note 4). Discard the bones fragments without bone marrow. 4. Aspirate the solubilised bone marrow and place it in a 50 mL sterile tube. Rinse the Petri dishes with another 20 mL PES

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percent of inhibition

100

Adherents Supernatants

75 50 25 0 0.40

0.10

G M -CSF concentration (ng/ml) Fig. 2. Inhibition of allogeneic MLR by Tol-DCs. The capacity of adherent BMDCs and BMDCs in suspension harvested at day 8 to regulate immune responses was studied using an in vitro assay described in Subheading 3.2. The basal MLR proliferation was 13.157 ± 6.073. One experiment representative of three is shown.

to rescue the maximum number of bone marrow precursors and add this solution to the tube containing the bone marrow solution. 5. Centrifuge the bone marrow solution at 500 × g for 7 min. 6. Aspirate the supernatant and re-suspend the pellet thoroughly in 5 mL RCLS (see Note 5). Incubate the mixture for 5–10 min at room temperature (see Note 6), then add PBS up to 50 mL. 7. Centrifuge the solution at 500 × g for 7 min. 8. Repeat steps 6 and 7. 9. Re-suspend the cell pellet in 10 mL of cold PES, then filter the solution (100-mm filter) and count the cells (see Note 7). 10. Centrifuge the solution at 500 × g for 7 min. 11. Re-suspend the cell pellet in complemented RPMI containing 0.4 or 0.1 ng/mL GM-CSF at 0.5 × 106 cells/mL. Pour 10 mL of the cell suspension into the Petri dishes (see Note 8) and incubate them at 37°C and 5% CO2. 3.1.2. Day 3

3.1.3. Day 6

1. Add 10 mL of pre-heated complemented RPMI alternatively 0.4 or 0.1 ng/mL of GM-CSF per Petri dish. 1. Aspirate 10 mL of cell suspension per Petri dish and place it in 50 mL tubes. 2. Centrifuge the tubes at 500 × g for 7 min. 3. Re-suspend the cell pellets in the same volume added before centrifugation using pre-heated, freshly-prepared complemented RPMI containing 0.8, 0.4 or 0.08 ng/mL GM-CSF. Add 10 mL of cells per Petri dish.

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Microscopic examination will reveal adherent cells and cells in suspension. We previously reported that rat (4) and non-human primate (5, 6) cultures display both populations. Both populations are harvested independently to assess their immunoregulatory properties: 1. Harvest the medium containing cells in suspension after flushing the cells two to three times. 2. Immediately add 10 mL of cold PES and incubate the cells for about 15 min (see Note 9). Pipette up and down with a 10 mL pipette until almost all cells are in suspension. 3. Centrifuge the cells at 500 × g for 7 min. 4. Re-suspend the tubes of adherent cells and cells in suspension using 5 mL of cold PES per Petri dish and count the cells. 5. Centrifuge the cells at 500 × g for 7 min. 6. Re-suspend the cells in complemented RPMI at 1 × 106/mL (see Note 10) and place them on ice while preparing the other cell types.

3.2. Co-culture Mixed Leukocyte Reaction

In order to assess whether BMDCs display active immunoregulatory properties, we study their capacity to inhibit the proliferation of syngeneic MLN cells (of the same haplotype as the BMDCs) stimulated by allogeneic antigen presenting cells.

3.2.1. Preparation of Responder MLN Cells

1. Dissect the mesenteric lymph nodes (MLN) from a C57/B6 mouse (H2kb). 2. Isolate the MLN cells by crushing the organs with a syringe tube on a 100-mm nylon strainer. Thoroughly rinse the strainer with cold PES and collect the cell suspension in a 50 mL tube. 3. Centrifuge the cells at 500 × g for 7 min. 4. Re-suspend the cell pellet in 10 mL of cold PES and count the cells. 5. Centrifuge the cells at 500 × g for 7 min. 6. Re-suspend the cell pellet in complemented RPMI at 2 × 106/ mL. Pour 1 × 105 MLN cells (50 mL/well) into round 96-well plates (see Note 11).

3.2.2. Preparation of Immunogenic Antigen Presenting Cells

1. Dissect two spleens from Balb/c mice (H2ka). 2. Place the spleens in a 10-cm Petri dish and add 5 mL of Collagenase D (2 mg/mL). Perfuse the spleens carefully until the entire organ becomes swollen. Cut the perfused spleen into about 1 mm3 pieces using a scalpel. 3. Incubate the pieces for 20 min at 37°C.

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4. Stop the collagenase reaction by adding 10 mM EDTA and incubating for 5 min. 5. Add about 30 mL of cold PES. 6. Transfer the cell suspension and spleen pieces onto a cell strainer and obtain a cell suspension by crushing the material with a syringe tube. 7. Centrifuge the cells at 500 × g for 7 min. 8. Re-suspend the cell pellet in 10 mL of 14.5% Nycodenz. 9. Add 5 mL of the cell suspension very slowly to a 15 mL tube containing 5 mL of 14.5% Nycodenz (see Note 12), using one tube per spleen. 10. Add 1 mL of EDTA/FCS and remove the interface slowly with a pipette tip. 11. Centrifuge the cells at 1,700 × g for 10 min at 4°C without the brake (see Note 13). 12. Harvest the ring containing low-density cells near the interface of the Nycodenz and the EDTA/FCS and transfer the cells to a 50 mL tube. 13. Add cold PES up to 50 mL. 14. Centrifuge the cells at 500 × g for 7 min. 15. Repeat steps 13 and 14. 16. Re-suspend the cell pellet in 5 mL of cold PES and count the cells. 17. Centrifuge the cells at 500 × g for 7 min. 18. Re-suspend the cell pellet in complemented RPMI (1 × 106/ mL). 3.2.3. Co-culture

Pour the following into 96-round well plates: (a) 50 mL of Balb/c immunogenic antigen presenting cells (5 × 104 cells) (b) 50 mL of C57/B6 Tol-DC (5 × 104 cells) (c) 100 mL of C57/B6 MLN cells (1 × 105 cells) Culture by triplicate for 5 days at 37°C with 5% CO2. Assess the proliferation of the MLN cells by studying 3H-thymidine incorporation. The mixed leukocyte reaction (MLR) basal proliferation is obtained by omitting the Tol-DCs. Instead, add 50 mL RPMI. The percentage of inhibition is obtained by the following formula:  CPM MLR + Tol - DC  1−   × 100 CPM ML  

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4. Notes 1. The choice of FCS batch is a critical issue that will determine the phenotype and functional properties of the BMDCs (7). 2. Very different results are obtained with treated cell-culture Petri dishes. 3. The differentiation of BMDCs from any mouse strain can be achieved using this protocol. Here, we chose C57/B6 to have a complete MHC mismatch with the immunogenic Balb/c antigen presenting cells in the co-culture MLR experiment. This protocol can also be performed with female mice, although the cell number obtained will be diminished as compared to males. 4. Be sure to obtain the maximum quantity of bone marrow. This will clearly impact the final quantity of cells. 5. Some protocols do not perform red cell lysis; however, we prefer to do it to avoid red cell death during the culture and the resulting release of haem. Haem is a potent inducer of the catabolising enzyme haem oxygenase-1 (HO-1), which can modulate DC biology (5, 8, 9). 6. Longer incubations will affect the final quantity of bone marrow precursors and the reproducibility of experiments. 7. Cell counting is an important issue as different cell densities during the differentiation culture can result in disparate phenotypes (7). 8. Pour 10 mL of cell suspension into all Petri dishes to maintain a constant cell density. Do not use Petri dishes with less than 10 mL. 9. This amount of time is needed to facilitate the separation of adherent cells from the plastic. 10. The FCS for the MLR assay is not necessarily the same at that used to differentiate bone marrow precursors. Different FCS batches must also be tested for the MLR assay. 11. Round wells increase the possibility of cellular contact in the MLR assay. 12. It is very important to add the cells very slowly to the Nycodenz tube. Otherwise, the density gradient will be seriously affected. 13. The brake must not be used during the centrifugation as it will affect the density gradient.

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Acknowledgments We are grateful to IMBIO, ESOT, CENTAURE, and RISET for financial support. References 1. Steinman, R.M., Hawiger, D., and Nussenzweig, M.C. (2003) Tolerogenic dendritic cells. Annu Rev Immunol 21:685–711. 2. Bluestone, J.A., Thomson, A.W., Shevach, E.M., and Weiner, H.L. (2007) What does the future hold for cell-based tolerogenic therapy? Nat Rev Immunol 7:650–654. 3. Morelli, A.E., and Thomson, A.W. (2007) Tolerogenic dendritic cells and the quest for transplant tolerance. Nat Rev Immunol 7:610–621. 4. Peche, H., Trinite, B., Martinet, B., and Cuturi, M.C. (2005) Prolongation of heart allograft survival by immature dendritic cells generated from recipient type bone marrow progenitors. Am J Transplant 5:255–267. 5. Moreau, A., Hill, M., Thebault, P., Deschamps, J.-Y., Chiffoleau, E., Chauveau, C., Moullier, P., Anegon, I., Alliot-Licht, B., and Cuturi, M.C. (2009) Tolerogenic dendritic cells actively inhibit T cells through heme oxygenase-1 in rodents and non-human primates. FASEB J 23:3070–3077. 6. Moreau, A., Chiffoleau, E., Beriou, G., Deschamps, J.Y., Heslan, M., Ashton-Chess,

J., Rolling, F., Josien, R., Moullier, P., Cuturi, M.C., et al. (2008) Superiority of bone marrow-derived dendritic cells over monocytederived ones for the expansion of regulatory T cells in the macaque. Transplantation 85:1351–1356. 7. Lutz, M.B., and Rossner, S. (2007) Factors influencing the generation of murine dendritic cells from bone marrow: the special role of fetal calf serum. Immunobiology 212:855–862. 8. Chauveau, C., Remy, S., Royer, P.J., Hill, M., Tanguy-Royer, S., Hubert, F.X., Tesson, L., Brion, R., Beriou, G., Gregoire, M., et al. (2005) Heme oxygenase-1 expression inhibits dendritic cell maturation and pro-inflammatory function but conserves IL-10 expression. Blood 106:1694–1702. 9. Remy, S., Blancou, P., Tesson, L., Tardiff, V., Brion, R., Royer, P.J., Motterlini, R., Foresti, R., Painchaut, M., Pogu, S., et al. (2009) Carbon monoxide generated by heme oxygenase activity confers tolerogenic capacity to dendritic cells. J Immunol 182:1877–1884.

Chapter 12 Myeloid-Derived Suppressor Cells: Characterization and Expansion in Models of Endotoxemia and Transplantation Nicolas Van Rompaey and Alain Le Moine Abstract CD11b+GR1+ myeloid-derived suppressor cells (MDSC) accumulate in several inflammatory conditions including cancer, infections, or trauma. MDSCs are found in bone marrow and lymphoid organs and suppress both innate and adaptive immune responses. Although mechanisms of suppression are not fully understood, they have been reported to require cell–cell contact and very often implicate L-arginine metabolism. We and others recently observed that lipopolysaccharide (LPS) administration, as other TLR ligands, induces MDSC. In this case, MDSC regulate immune response independently of L-arginine metabolism through heme oxygenase-1 activity. Manipulating MDSC as immunoregulators represents an attractive approach for cancer immunotherapy or transplantation. Herein, we describe methods for expanding and purifying MDSC, as well as in vitro and in vivo techniques to measure their suppressive functions. Key words: Myeloid-derived suppressor cells, Lipopolysaccharide, T cells, Transplantation, Heme oxygenase-1

1. Introduction Myeloid-derived suppressor cells (MDSC) constitute a relatively new population of regulatory cells. Initially described as myeloid natural suppressor, 20 years ago (1), their potential role in suppression immunity – antitumor immunity in particular– has grown over the last decade especially because they are potent inhibitor of innate and adaptive responses (2). Present transiently in the bone marrow of healthy individuals during the process of maturation of myeloid cells, they can expand dramatically under pathological conditions in which their suppressive capacities have been described. Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_12, © Springer Science+Business Media, LLC 2011

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Indeed, they are a heterogeneous population of immature cells derived from the myeloid lineage interrupted at different stages of maturation which is underlined by the fact that they can differentiate into mature cells under specific conditions in vitro as well as in vivo (3). MDSC expansion is induced by a variety of proinflammatory mediators such as GM-CSF, G-CSF, VEGF associated to cancer or other inflammatory conditions such as bacterial or viral infection, auto-immunity, trauma (2), and transplantation (4). Under chronic inflammation, MDSC accumulate in the spleen, the peripheral blood, and bone marrow (5). Even though MDSC constitute a heterogeneous population with extracellular phenotype differing from one to another experimental mouse model, they are commonly identified by the coexpression of the CD11b antigen (an adhesion molecule also known as Mac-1, mainly expressed on monocytes/macrophages and granulocytes) and the Gr-1 antigen (the epitope of the GPI-anchored glycoproteins, Ly-6G and Ly-6C, expressed transiently by monocytes and predominantly by neutrophils). In humans, MDSC are generally described as a population of cells expressing CD11b and CD33 but lacking the expression of CD14 and HLA-DR (6). Precise phenotype and mechanisms by which these cells express their immunosuppressive capacities depend on the model considered and the environmental milieu (7). Nevertheless, most studies have shown the need of close cell–cell contact to suppress activated T cells (8, 9), suggesting a role of surface receptors or short-lived soluble mediators. Initially, it has been described that MDSC mediate T-cell suppression through two key enzymes involved in L-arginine metabolism working either alone or synergistically: arginase-1, which depletes arginine of the milieu and nitric oxide synthase-2, which produces NO (3) resulting in the inhibition of T-cell proliferation and induction of their apoptosis. More recently, the release of reactive oxygen species as peroxynitrite by MDSC has also been described (9). In the context of lipopolysaccharide (LPS)-desensitization (multiple injections of low doses of LPS inducing tolerance to a normally lethal dose), we have shown that CD11b+GR1+ cells expand dramatically in the bone marrow, the spleen, and the lymph nodes. These cells showed potent direct cell–cell contact dependent mechanism of T-cell suppression in vitro and in vivo. In this setting, a new mechanism of suppression by MDSC was identified (10), namely the heme oxygenase-1 (HO-1), a powerful antiinflammatory and antioxidant enzyme involved in many immunoregulatory processes (as extensively described by Philippe Blancou and Ignacio Anegon in the discussion chapters). Targeting MDSC represents a promising approach for cancer immunotherapy for improving immune reactivity against tumor antigens, as well as in the field of transplantation by stimulation of their expansion and/or suppressive capacities. Herein, we describe

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our approach to expand and isolate LPS-induced MDSC, and how we investigated their in vitro and in vivo suppressive functions.

2. Materials 2.1. Expansion of MDSC by LPS

1. LPS from Escherichia coli 0111:B4 purified by phenol extraction dissolved in phosphate buffered saline (PBS), calcium, magnesium free, pH 7.2 to a working dilution of 250 mg/ml. Store in single use aliquots at −20°C (see Note 1). 2. One milliliter syringe with 26G needle. 3. Six- to eight-week-old C57BL/6 mice.

2.2. Flow Cytometry Analysis of MDSC on Bone Marrow, Spleen, and Lymph Nodes and Purification by Magnetic Cell Sorting

1. Roswell Park Memorial Institute Medium 1640 (RPMI1640) with 25 mM Hepes an L-glutamine. 2. Cell culture medium: RPMI-1640 supplemented with 5% heat-inactivated fetal bovine serum (FBS), 0.04 mM b-mercaptoethanol, 1 mM Na pyruvate, 1% nonessential amino acid solution, 60 mg/ml streptomycin sulfate, 60 U/ml penicillin G, 0.15 mg/ml Amphotericin B. Filter with 0.22 mm syringe filter and store at 4–8°C. 3. Red blood cell (RBC) lysis buffer (ACK buffer): 0.15 M NH4Cl, 10 mM NaHCO3, 0.1 mM Na2 ethylenediaminetetraacetic acid (EDTA) in distilled H2O. Filter and store at 4–8°C. 4. FACS buffer: PBS supplemented with 0.1% of bovine serum albumine and 1.17 mM NaN3 and 2 mM (EDTA), store at 4–8°C. 5. Rat anti-mouse FcgRIIA (CD16/CD32) (clone 2.4G2, Fc Block, BD Biosciences) diluted at 0.01 mg/ml in FACS buffer the day of the experiment. Store at 4–8°C. 6. Rat anti-mouse GR-1 phycoerythrin (PE) (clone RB6-8C5, BD Biosciences) diluted at 4 mg/ml in FACS buffer the day of the experiment. Store at 4–8°C protected from light exposure. 7. Rat anti-mouse CD11b APC (clone M1/70, BD Biosciences) diluted at 1 mg/ml in FACS buffer the day of the experiment. Store at 4–8°C protected from light exposure. 8. BD Cell Fix solution (BD Biosciences) diluted 10× in FACS buffer. 9. Dual laser flow cytometer (e.g., CyAn LX cytometer, Dako) and analysis software (e.g., Summit 4.1 Cell, DakoCytomation).

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2.3. Purification by Magnetic Cell Sorting

1. Sorting buffer: PBS containing 0.5% BSA and 2 mM EDTA, stored at 4–8°C. 2. Mouse CD11b positive selection kit (CD11b Microbeads, Miltenyi). 3. MACS LS-column (Miltenyi). 4. QuadroMACS separator (Miltenyi).

2.4. In Vitro Suppression Assays

1. Mouse anti-CD3/CD28 microbeads (Dynabeads Mouse T-Activator CD3/CD28, Invitrogen): transfer the desired volume of microbeads in a tube and wash by adding the same volume (or at least 1 ml) of PBS supplemented with 0.1% of BSA and 2 mM EDTA, place tube in a strong magnet for 1 min, discard the supernatant, and resuspend microbeads in the same initial volume. 2. Ninety-six-well U-shaped bottom culture plate. 3. Transwell chamber (Anopore insert 0.2 mm, Nunc). 4. Ninety-six-well flat bottom MicroWell™ plates (Nunc). 5. (Methyl-3H)thymidine 1 mCi/ml. 6. Scintillation liquid. 7. Filter-bottomed 96-well plates. 8. Tin protoporphyrin IX (SnPP, Enzo Lifesciences) dissolved in 0.1 mM NaOH to a concentration of 2 mg/ml (SnPP is light sensitive).

2.5. Adoptive Transfer and Skin Grafting

1. C57BL/6 male donor and female recipient mice (for minor histocompatibility disparate skin graft) (see Note 2), 6–8 weeks old. 2. Anesthetic solution: prepare 0.1% xylazine, 1% ketamine diluted in water for injection. Filter and store at 4–8°C. 3. Prepare wide surgical tape (2.5 cm) and another adhesive tape, it will be used to fix the bandages. 4. 100 × 100 mm paraffin gauze dressing. 5. 94 × 16 mm petri dishes. 6. Scalpel blades. 7. Microdissecting tools (scissor, forceps, etc.).

3. Methods As shown in our laboratory, repetitive sublethal doses of LPS can induce a reliable, strong, and predictable expansion of MDSC. Mouse mortality associated with this technique is around 5–10%,

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depending much on the age and mouse weight. Expansion of CD11b+GR1+ cells after LPS treatment can be estimated easily by flow cytometry analysis on different target organs (bone marrow, spleen, or lymph nodes). The isolation of MDSC from spleen can be achieved using a magnetic CD11b positive cell sorting technique which is very useful because it is easy, quick and gives reproducible yields of good purity cells (see Note 3). Cytospin centrifugation and Giemsa coloration on purified cells will give a morphologic insight to confirm their myeloid and immature origins. To assess their direct suppressive effect on T-cell proliferation, purified MDSC can be added to polyclonally activated T cells. T-cell activation is calculated after determining the difference in incorporation of (3H)thymidine between stimulated and control cells. Moreover, this straightforward in vitro approach offers the possibility to investigate pathways of suppression, as well as the role of cell–cell contact using a transwell semi-permeable membrane and the role of HO-1 by adding tin-protoporphyrin IX, a selective inhibitor of HO-1 (11), to suppression assays. Finally, the in vivo suppressive function of MDSC can be evaluated using an adoptive transfer of MDSC prior skin allograft transplantation. 3.1. LPS Expansion of MDSC

1. Thaw LPS solution to room temperature, vortex strongly 1 min to have a homogeneous solution. 2. For 3 days, inject intraperitoneally 2 mg/g of LPS every 24 h for the first 2 days and 16 mg/g for the last one.

3.2. Flow Cytometry Analysis of MDSC on Bone Marrow, Spleen, and Lymph Nodes

1. For flow cytometry analysis, euthanize the mice by carbon dioxide asphyxiation or other approved method and harvest spleen, lymph nodes and bone marrow to make a single cell suspension as followed. 2. Clean trunk of each mouse with 70% ethanol solution and make a median incision towards the upper and lower limbs with scissor to expose the inguinal lymph nodes, the spleen, and the femoral and tibia bones. 3. Remove the spleen and lymph node and disrupt them with the plunger of a 5 ml syringe in a petri dish filled with 5 ml culture medium. 4. To extract bone marrow, dissect closely the femoral and tibia bones, release their hip and ankle attachment, and cut the epiphysis. Flush the intramedullary compartment with culture medium through a 1 ml syringe with a 26G needle onto a petri dish filled with 5 ml culture medium. 5. Pass the cell on a 100 mm sieve to obtain a single cell suspension, centrifuge at 400 × g for 10 min at 4–8°C and remove the supernatant.

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6. Add 1 ml of ACK buffer to lyse the erythrocytes for 1 min maximum and neutralize with 10 ml of culture medium. Centrifuge at 400 × g for 10 min at 4–8°C and remove the supernatant. 7. Resuspend cells in FACS buffer to a concentration of 20 × 10E6 cells/ml, aliquot 50 ml of the cell suspension (1 × 10E6 cells) to each FACS tube, and add 50 ml (0.5 mg) of diluted Fc Block solution, incubate 10 min at 4–8°C. 8. Add 50 ml of mixed CD11b-APC, GR1-PE solution and incubate 20 min at 4–8°C in the dark. 9. Wash with 2 ml of FACS buffer and centrifuge at 400 × g for 5 min at 4–8°C. Discard supernatant and resuspend in 250 ml of diluted Cell Fix solution. 10. Set up data acquisition parameters on the flow cytometer using single-stained controls and positive and negative control samples. Acquire a minimum of 10,000 events from each sample tube. A characteristic result is shown in Fig. 1. Seven days after LPS desensitization. 1. Five to seven days after the first injection of LPS, euthanize the mice.

3.3. Purification by Magnetic Cell Sorting

2. Harvest the spleen following the same protocol as above until step 6 and resuspend the pellet of cells in 900 ml of cooled sorting buffer for 10E8 cells. Lymph node

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Fig. 1. FACS analysis of CD11b+GR1+ cells in lymph nodes, spleen, and bone marrow of mice 7 days after lipopolysaccharide (LPS) desensitization compared to untreated mice.

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3. Add 100 ml of CD11b microbeads for 10E8 cells and incubate 15 min at 4–8°C (see Note 4). 4. Wash the cells by adding 2 ml of sorting buffer for 10E7 cells and centrifuge at 400 × g for 10 min at 4–8°C. 5. Resuspend the pellet in 500 ml sorting for 10E8 cells. 6. Pass the cells on a 100 mm filter (see Note 5). 7. Place LS-column in a strong magnet and prepare LS-column by adding 3 ml of sorting buffer. 8. Load the cell suspension onto the LS column and wash three times with 3 ml of sorting buffer. 9. Remove the column from the magnet and flush out the positive fraction by flushing the column with 5 ml of sorting buffer into a new sterile 15 ml tube with the plunger supplied. 10. Expected yield should attempt 5 × 10E6 cells for purity around 80% of CD11b+GR1+ cells evaluated by flow cytometry analysis as described above (Fig. 2). 11. Morphologic aspect of purified cells can be obtained by cytospin centrifugation and a classical GIEMSA coloration (Fig. 3). 12. Further phenotype characterization can be made after cell sorting. As shown in Fig. 4, MDSC induced by LPS express very low levels of MHC-II and CD11c but similar levels of F4/80 (a macrophage maker). They also express lower levels of the IL-4 receptor alpha chain, CD31 (a marker of granulocyte/monocyte precursors) and PDL-1, compared to GR-1 negative cells. 3.4. In Vitro Suppression Assays

1. Purify total T cells using the same magnetic cell-sorting technique as above with CD90.2 positive selection kit from Miltenyi.

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Fig. 3. Cytospin preparation of enriched CD11b cells 7 days after LPS treatment demonstrating immature heterogeneous myeloid phenotypes with characteristic ring-shaped nuclei.

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Fig. 4. Expression of MHC-II, CD11c, F4/80, IL-4 receptor alpha chain, CD31, and programmed death ligand-1 was measured on gated CD11b+GR-1 positive cells (black line) or gated CD11b+GR-1 negative cells (gray line). CD11b cells were sorted from spleen of LPS-desensitized mice. Arrows indicate percentages of positive cells compared to fluorescenceminus-one (FMO) control (gray area). Percentage are expressed as mean ± SD of minimum five individual mice.

2. Resuspend cells in culture medium to obtain concentration of 1 × 10E6 cells/ml and stimulate purified T cells by adding 25 ml of washed aCD3/CD28 microbeads for 10E6 cells. 3. Add 100 ml of purified T-cell solution (1 × 10E5 cells) per well in 96-well U-shaped bottom culture plate and add

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CD11b+ cells at different ratio to assess their suppressor effects in a dose–response fashion. 4. For transwell experiments, concentrate the desired number of CD11b+ cells in maximum 60 ml of culture medium to place in the transwell insert, whereas T cells on the bottom of the MicroWell plate must be suspended in 175 ml of culture medium (see Note 6). 5. To assess the role of HO-1, add 10 ml of the solution of SnPP to obtain a final concentration of 100 mM per well (see Note 7). 6. Incubate in a humidified incubator at 37°C with 5% CO2 for 24–72 h. 7. Add 1 mCi of [3H]thymidine to each well and incubate 18–24 h in the incubator. 8. Harvest the plate on a filter-bottomed 96-well plate (see Note 8) and dry for 2–12 h in a dry incubator at 40°C, finally add 30 ml of scintillation liquid before measuring cpm in b scintillation counter. 9. ELISA test on cell culture supernatant can also be made to assess modulation of cytokines production. An example of the suppression assay results is shown in Fig. 5. 3.5. Adoptive Transfer and Skin Grafting

1. After female CD11b+ cell sorting as described above, wash the cells with PBS and resuspend them up to a concentration of 5 × 10E6 cells for 200 ml (which is the maximum iv volume that can be administered to an adult mouse). Keep cells on ice until the injection.

Anti-CD3/CD28 stimulated T cells only Anti-CD3/CD28 stimulated T cells + CD11b cells

Fig. 5. Proliferation and IFN-g production by anti-CD3/CD28-stimulated T cells (white box ) were suppressed in a dosedependent manner by the addition of CD11b+ cells from LPS-desensitized mice at increased ratio (black box ). Cell proliferation and IFN-g production were measured by thymidine incorporation and ELISA test after 72 h of culture. Results are expressed as mean ± SD of triplicate cultures.

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2. Inject the autologous cells in the tail vein the day prior to the skin transplantation (see Note 9). 3. The day of the skin transplantation, sacrifice donor mice, clean its tail with appropriate solution for disinfection and remove full-thickness tail skin by incising the base circumferentially and making a ventral midline incision, preserve in physiological solution. 4. Anesthetize recipient mice with 100–120 ml of anesthetic solution, wait until profound sleep, and wet and shave it with razor blade (see Note 10). 5. For the graft bed, practice 1 cm2 round skin incision on flank. 6. Fit the tail skin piece to the graft bed and place it gently drying it after with a gauze. 7. Place paraffin gauze on it and wrap the mouse tied up with the surgical tape and consolidate it with an adhesive tape of the same size. 8. Wait until complete recovery of the mice and see if the bandages fit well. 9. Thereafter, mice will be followed every day to prevent the take away of the bandage. 10. Remove the bandage 9 days later with a blunt-end scissor and monitor the skin graft rejection every day. 11. Consider the graft rejected when more than 75% of epithelial breakdown had occurred.

4. Notes 1. Use silanized containers since LPS can bind to plastics and certain types of glass. 2. Strain is very important for the median survival time in minorH disparate skin graft (12). 3. Better (>80%) purity can be achieved by flow cytometric cell sorting but it takes more time (impeding the viability of cells), and the yield is far poorer limiting the in vivo studies that are consuming a lot of cells. 4. Use a roller mixer to label the cells uniformly, be accurate on the timing, extended staining induce nonspecific labeling. 5. It is critical to filter the cells to prevent the columns from clogging. The cells are very concentrated and rinsing the filter with a small volume of sorting buffer will increase recovery.

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6. Place the CD11b+ cells in the transwell insert before putting it on the MicroWell plate and ensure that no air trapping occur. To prevent desiccation of the culture, seal the plate with a porous tape. 7. SnPP may impair T-cell proliferation at high concentration, if you use higher concentration, do not forget to have conditions with activated T cells with SnPP and without CD11b+ cells. 8. Waste materials are radioactive, manipulate and store with precautions. 9. Heat the mice with a heat lamp, it will dilate vein and facilitate the iv injection. 10. Manipulate the mice on a heated mattress or a heat lamp to avoid the cool down.

Acknowledgment Nicolas Van Rompaey is Research Fellow of the Fonds Erasme and Alain Le Moine is Research associate of the Fonds National de la Recherche Scientifique FNRS-Belgium. References 1. Strober, S. (1984). Natural suppressor (NS) cells, neonatal tolerance, and total lymphoid irradiation: exploring obscure relationships. Annu. Rev. Immunol. 2: 219–237. 2. Ostrand-Rosenberg, S., and P. Sinha. (2009). Myeloid-derived suppressor cells: linking inflammation and cancer. J. Immunol. 182: 4499–4506. 3. Bronte, V., and P. Zanovello. (2005). Regulation of immune responses by L-arginine metabolism. Nat. Rev. Immunol. 5: 641–654. 4. Zhang, W., S. Liang, J. Wu, and A. Horuzsko. (2008). Human inhibitory receptor immunoglobulin-like transcript 2 amplifies CD11b+Gr1+ myeloid-derived suppressor cells that promote long-term survival of allografts. Transplantation 86: 1125–1134. 5. Ezernitchi, A. V., I. Vaknin, L. Cohen-Daniel, O. Levy, E. Manaster, A. Halabi, E. Pikarsky, L. Shapira, and M. Baniyash. (2006). TCR zeta down-regulation under chronic inflammation is mediated by myeloid suppressor cells differentially distributed between

6.

7.

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various lymphatic organs. J. Immunol. 177: 4763–4772. Almand, B., J. I. Clark, E. Nikitina, J. van Beynen, N. R. English, S. C. Knight, D. P. Carbone, and D. I. Gabrilovich. (2001). Increased production of immature myeloid cells in cancer patients: a mechanism of immunosuppression in cancer. J. Immunol. 166: 678–689. Gabrilovich, D. I., and S. Nagaraj. (2009). Myeloid-derived suppressor cells as regulators of the immune system. Nat. Rev. Immunol. 9: 162–174. Mazzoni, A., V. Bronte, A. Visintin, J. H. Spitzer, E. Apolloni, P. Serafini, P. Zanovello, and D. M. Segal. (2002). Myeloid suppressor lines inhibit T cell responses by an NO-dependent mechanism. J. Immunol. 168: 689–695. Nagaraj, S., K. Gupta, V. Pisarev, L. Kinarsky, S. Sherman, L. Kang, D. L. Herber, J. Schneck, and D. I. Gabrilovich. (2007). Altered recognition of antigen is a mechanism of CD8+ T-cell tolerance in cancer. Nat. Med. 13: 828–835.

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10. De Wilde, V., N. Van Rompaey, M. Hill, J. F. Lebrun, P. Lemaitre, F. Lhomme, C. Kubjak, B. Vokaer, G. Oldenhove, L. M. Charbonnier, M. C. Cuturi, M. Goldman, and A. Le Moine. (2009). Endotoxin-induced myeloid-derived suppressor cells inhibit alloimmune responses via heme oxygenase-1. Am. J. Transplant. 9: 2034–2047.

11. Ryter, S. W., J. Alam, and A. M. Choi. (2006). Heme oxygenase-1/carbon monoxide: from basic science to therapeutic applications. Physiol Rev. 86: 583–650. 12. Gasser, D. L., and W. K. Silvers. (1972). Genetics and immunology of sex-linked antigens. Adv. Immunol. 15: 215–247.

Chapter 13 Human Regulatory Macrophages James A. Hutchinson, Paloma Riquelme, Edward K. Geissler, and Fred Fändrich Abstract Regulatory macrophages (M regs) are a novel type of suppressor macrophage which may be a particularly suitable cell for inducing tolerance of solid organ transplants. In this article, we provide a detailed description of the generation of human M regs from peripheral blood monocytes and methods for the assessment of their phenotype. The uniqueness of the human M reg is best appreciated when the M reg is compared to macrophages in other states of activation; therefore, protocols are provided for generating five comparator macrophage types which have been used as cell type-specificity controls in our work. Key words: Suppressor macrophage, Transplantation tolerance, Cellular therapy

1. Introduction Although the induction and adoptive transfer of allograft tolerance with suppressor cell preparations are common methods in Experimental Immunology, there remain very substantial obstacles to the clinical translation of these techniques. From a clinical perspective, any cellular product intended for use as a tolerancepromoting therapy must be predictably efficacious and safe for the recipient. In practical terms, this means that any cell-based therapeutic product must be consistently homogeneous in phenotype and function, and that its properties remain unaltered by administration to the recipient. Consequently, therapeutic cell preparations must be accurately defined in terms of their phenotype, yield and possible contaminants, in order that they may be properly quality controlled. In addition, even if treatment with a tolerance-promoting cellular therapy can be shown to afford a benefit without imparting unacceptable risk, it must Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_13, © Springer Science+Business Media, LLC 2011

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ultimately be shown to be cost-effective compared to current immunosuppressive treatments. The human regulatory macrophage (M reg) reflects a unique state of macrophage differentiation, distinguished from macrophages in other activation states by its particular mode of derivation, robust phenotype and potent T cell suppressive function. M regs may represent a particularly suitable cell type for use in clinical tolerance-promoting strategies, because they are easily and reliably generated from peripheral blood monocytes under GMP conditions and can be safely administered to patients by central venous infusion ((1–6) and Hutchinson, unpublished data). Experiments in vitro demonstrate that M regs are very stable in their phenotype and T cell suppressive capacity: such functional stability is a crucial property of any tolerance-promoting cell type, as conversion to an effector phenotype may cause sensitisation or GvH-like reactions. As M regs appear to be a terminally differentiated, postmitotic cell type with a finite lifespan, their potential to cause or promote malignancies may be less than that of certain alternative tolerogenic cell preparations. In addition, as M regs have been shown to promote completely mismatched allograft survival in fully immunocompetent animals, treatment with human M regs does not require lymphodepletive or myeloablative conditioning, which are essential components of some alternative cell-based tolerancepromoting strategies. This chapter describes an experimental protocol for the generation of human M regs; however, a close adaptation of this method has been used to produce M regs for administration to patients. Wherever possible, reagents for which a GMP-approved equivalent is available are recommended, although alternative products may be suitable for purely experimental purposes. A population of macrophages closely resembling human M regs can be derived from mice using analogous techniques (7).

2. Materials 2.1. Preparation of Human PBMC

1. Dulbecco’s phosphate buffered saline (DPBS) without Ca2+ and Mg2+ (Lonza, Belgium). 2. Biocoll (Ficoll separating solution) 1.077 g/ml (Biochrom AG, Germany). 3. Whole human blood or PBMC concentrates anticoagulated with heparin, EDTA, citrate, ACD-A, or CPD. 4. Neubauer chamber and Trypan blue solution or an automated cell counter.

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1. Sorting buffer: DPBS without Ca2+ and Mg2+ (Lonza, Belgium) containing 0.5% human albumin (Aventis, Germany) and 10% citrate-phosphate-dextrose solution with adenine (CPD-A) (Sigma, Germany). 2. 0.2-mm vacuum filter unit for filtration of tissue culture medium and solutions (Millipore, Germany). 3. MACS columns, MACS separator, and preseparation filters (Miltenyi Biotec, Germany). 4. CD14 MicroBeads, human (Miltenyi Biotec).

2.3. Generation of M regs in Culture

1. M reg culture medium: RPMI 1640 with phenol red (Lonza) containing 10% human AB serum (Lonza) (see Note 1), 2-mM GlutaMAX™ (Invitrogen, Germany), 100-U/ml penicillin, 100-mg/ml streptomycin (Invitrogen), and recombinant human M-CSF (R&D Systems, Germany) at a final concentration 5 ng/ml, carried on human serum albumin. After preparation, the medium should be passed through a 0.2-mm sterilisation filter. 2. Cell+ tissue culture vessels, flask, or plates (Sarstedt, Germany). 3. Human recombinant interferon-g (Chemicon, Millipore).

2.4. Recovery of Human M regs from Culture

1. DPBS without Ca2+ and Mg2+ (Lonza). 2. HyQTase (ThermoFisher Scientific, Germany). 3. Rubber cell scrapers (Sarstedt). 4. Neubauer chamber and Trypan blue solution.

2.5. Phenotypic Characterisation of Human M regs by Flow Cytometry

1. Staining buffer: DPBS without Ca2+ and Mg2+ (Lonza) containing 1% BSA and 0.1% NaN3.

2.6. Assessing the Direct Suppression of Mitogen-Stimulated T cell Proliferation by M regs

1. 1% HABS medium: RPMI 1640 with phenol red (Lonza) containing 2-mM GlutaMAX™ (Invitrogen, Karlsruhe, Germany), 100-U/ml penicillin, 100-mg/ml streptomycin (Invitrogen).

2. FcR blocking reagent (Miltenyi Biotec). 3. 7-AAD, for dead cell exclusion (BD Biosciences).

2. Human AB serum (Lonza). 3. DPBS without Ca2+ and Mg2+ (Lonza). 4. CFSE solution: CellTrace™ CFSE Cell Proliferation Kit (Invitrogen). 5. CountBright beads (Invitrogen). 6. FlowJo software (Ashland, OR, USA).

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2.7. Comparator Macrophage Types

1. Culture medium: RPMI 1640 with phenol red (Lonza) containing 2-mM GlutaMAX™ (Invitrogen, Germany), 100-U/ml penicillin, 100-mg/ml streptomycin (Invitrogen). 2. Human AB serum (Lonza). 3. Foetal calf serum (FCS) (Biochrom, Germany). 4. Recombinant human M-CSF (R&D Systems). 5. Human recombinant interferon-g (Chemicon, Millipore). 6. Lipopolysaccharide (LPS) from S. minnesota (Sigma-Aldrich). 7. Human recombinant IL-4 (R&D Systems). 8. Dexamethasone (Sigma). 9. Human immunoglobulin for intravenous infusion (IVIG): Sandoglobulin™ (CSL Behring, Germany).

3. Methods Under inflammatory conditions, classical monocytes traffic to sites of inflammation and enter the tissues, acquiring the characteristics of activated macrophages. In the absence of inflammation, classical monocytes give rise to resident monocytes, which may enter noninflamed tissues to contribute to resident macrophage populations. It is hypothesised that the generation of M regs from monocytes in vitro emulates the transition of peripheral blood monocytes to a tissue-resident macrophage type. The practical corollary of this hypothesis is that the development of M regs can be disrupted by exposure to proinflammatory stimuli, especially microbial components and other activated leucocytes. 3.1. Preparation of Human PBMC

The generation of human M regs from peripheral blood monocytes is usually very consistent; however, occasionally some preparations do not develop as expected. To minimise the rate of failed preparations, blood donors should be carefully selected to exclude any individuals with ongoing or recent illnesses, and the blood should be processed within a few hours of collection (see Note 2). 1. Whole human blood anticoagulated with heparin or citrate is diluted 1:1 with DPBS at room temperature (RT). 2. Fifteen millilitres of diluted blood are layered by pipetting onto 15-ml Ficoll in a 50 ml Falcon tube before centrifugation at 850 × g for 20 min at RT without brake. 3. The leucocyte layer at the interface is collected using a serological pipette and subsequently washed twice in DPBS at RT by centrifugation at 500 × g for 10 min, followed by a single

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wash in DPBS with centrifugation at 200 × g for 10 min to remove platelets. 4. The cell pellet is resuspended in an appropriate volume of DPBS and a sample is taken for a differential leucocyte count. 3.2. Positive Selection of CD14+ Human Monocytes

The following protocol for the isolation of monocytes using magnetic beads differs slightly from the manufacturer’s recommendations. An alternative buffer is used, which contains CPD-A instead of EDTA and human albumin instead of BSA. The purity of the isolated monocytes can be maximised by passing the cells through a second MACS column. This procedure consistently results in CD14+ monocyte purities of ³98% with ³95% cell viability, assessed by 7-AAD exclusion in flow cytometry. 1. Sorting buffer should be prepared freshly and degassed prior to use, as it acidifies when stored. 2. PBMCs are pelleted by centrifugation at 300 × g for 10 min and are resuspended in sorting buffer at a density of 1.25 × 105 cells/ml. 3. Two microlitres of CD14 MicroBeads are added per 106 PBMCs, mixed well, and incubated at 2–8°C for 15 min. 4. The cells are washed once with 0.2 ml sorting buffer per 106 cells and centrifuged at 300 × g for 10 min. 5. Resuspend up to 100 × 106 cells in 500 ml of sorting buffer. 6. Magnetic separation is performed according to the manufacturer’s recommendations. 7. The CD14+ eluted monocyte fraction should be applied to a second column to increase purity. 8. Before plating, the isolated monocytes are washed twice with DPBS.

3.3. Generation of M regs in Culture

For experimental and clinical purposes, M regs can be easily generated from purified human peripheral blood monocytes. M regs develop optimally when cultured in Cell+ coated plasticware: depending on the application, M regs can be generated in tissue culture flasks or plates. 1. Resuspend sorted CD14+ monocytes at 106 cells/3 ml in freshly prepared M reg culture medium. 2. Plate monocytes at 105 cells/cm² in Cell+ tissue culture vessels and place at 37°C and 5% CO2 in an incubator. 3. On day 6 of culture, the cells are stimulated for 18–24 h with IFN-g at a final concentration of 25 ng/ml. 4. On day 7 of culture, the M regs have obtained their final phenotype.

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3.4. Recovery of Human M regs from Culture

Human M regs adhere very tightly to plastic, which makes them difficult to recover without loss of viability. Excellent cell viability is obtained if M regs are harvested using HyQtase™ (ThermoFisher Scientific) but a substantial loss of cell surface markers is observed. When the cell surface phenotype of the cells must be preserved, M regs can be harvested by scraping into DPBS, but cell viability is then much reduced. HyQtase is not certified for use under GMP conditions, but certain trypsin preparations can be used for clinical cell production, such as TrypLE Express™ recombinant trypsin without phenol red (Invitrogen, NY, USA) with tolerable loss of cell viability. 1. M regs are washed three times with room temperature DPBS to remove nonadherent cells and culture medium. 2. (a) If using HyQtase™, the M regs are submerged in HyQTase solution (1 ml/25 cm2) and incubated at RT until they begin to detach from the plate (~30 min) at which point they can be recovered by gentle pipetting. The harvested cells should then be gently washed once (500 × g, 5 min) in medium containing 10% HABS. Any cells remaining attached to the plate can be collected by scraping. (b) If the cells are to be physically detached, use a rubber scraper to softly lift the cells into DPBS suspension. The cells inevitably form clumps that must be disaggregated by gentle pipetting. Violent pipetting or vortexing will damage the cells. 3. M regs lose viability if kept in suspension for prolonged periods. Ideally, the cells should be processed within 1 h of harvesting.

3.5. Phenotypic Characterisation of Human M regs by Flow Cytometry

Analysing human M regs by flow cytometry presents a number of technical challenges. M regs are relatively large, densely granular cells with high background autoflourescence and a high capacity for nonspecific antibody binding. During preparation for analysis by flow cytometry, these cells have an aggravating tendency to flocculate, which can make them difficult to precipitate by centrifugation and may affect the uniformity of staining. When analysing M regs by flow cytometry, it is imperative to perform dead-cell exclusion with 7-AAD, as most preparations contain nonviable cells that label nonspecifically. 1. Harvest the M regs into ice-cold staining buffer (DPBS, 1% BSA, 0.1% NaN3). 2. Wash twice in cold staining buffer by centrifugation at 4°C for 5 min at 500 × g. 3. Resuspend the M regs at 10 × 106 cells/ml in staining buffer containing 10% FcR-block and incubate the cell suspension for 30 min on ice. 4. Transfer aliquots of 106 cells into FACS tubes and add directly conjugated fluorescent antibodies.

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Live M regs CD14-/low HLA-DRint CD80low CD86+฀CD83−

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Fig. 1. Gating strategy for the analysis of human regulatory macrophages (M regs) by flow cytometry.

5. Minimise exposure to light. Incubate on ice for 60 min. 6. Add 5 ml of 7-AAD into the samples, then incubate for a further 10 min. 7. Wash three times with cold staining buffer, centrifuging for 5 min at 500 × g at 4°C and carefully aspirating the supernatant with a vacuum pump. 8. Resuspend the cells in 400 ml staining buffer for analysis with the flow cytometer. 9. An appropriate gating strategy is given in Fig. 1. 3.6. Assessing the Direct Suppression of Mitogen-Stimulated T cell Proliferation by M regs

Human M regs are profoundly suppressive of PHA-stimulated T cell proliferation in vitro, an effect which can be partly explained by the activation-dependent elimination of directly cocultured T cells. Both the suppression of T cell division and deletion of T cells can be conveniently assayed by flow cytometry using CFSElabelled responder T cells and absolute enumeration of T cells with CountBright™ beads (8). A schematic overview of the experiment is given in Fig. 2. 1. As described above, M regs are generated over 7 days in 96-well Cell+ coated tissue culture plates. 2. On the sixth day of M reg culture, PBMC should be isolated from a healthy donor according to the protocol given above and resuspended at a density of 20 × 106 cells/ml in DPBS at 37°C. 3. One millilitre fresh 2.5-mM CFSE solution is prepared in DPBS and warmed to 37°C. 4. Equal volumes of the PBMC suspension and CFSE solution are mixed and incubated for precisely 10 min in a 37°C waterbath. The labelling reaction is halted by the addition of 2-ml HABS. The cells are washed three times (500 × g, 5 min) in 10% HABS medium. 5. The labelled cells are cultured overnight in a 75 cm2 flask in 10-ml 10% HABS medium.

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Fig. 2. M reg-mediated T cell suppression and elimination can be conveniently assayed by flow cytometry. (a) A schematic overview of the generation of M regs and subsequent coculture with CFSE-labelled responder T cells. (b) An appropriate gating strategy for the absolute enumeration of CD4+ and CD8+ T cells in coculture with M regs and the concomitant analysis of their proliferation by CFSE dilution.

6. Unlabelled PBMCs are likewise cultured overnight in a 75 cm2 flask with complete medium. These cells are necessary for properly adjusting the settings of the flow cytometer. 7. On day 7 of culture, collect the nonadherent CFSE-labelled responder cells and isolate the T cell fraction using the Pan-T Cell Isolation Kit (Miltenyi) according to the manufacturer’s instructions. Adjust the density to 5 × 106 cells/ml in 1% HABS medium.

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8. Wash the adherent M regs three times with warmed 1% HABS medium, leaving the cells in a final volume of 20 ml of medium. 9. A range of M regs:T cells ratios in coculture should be tested: 1:1, 1:2, 1:4, 1:8, and 1:16. After addition of the appropriate volume of the T cell suspension to each well (i.e. 10, 20, 40, 80 or 160 ml), the volume in each well should be adjusted to 100 ml. PHA is added to each of the M reg:T cell cocultures at a final concentration of 2 mg/ml. T cells cultured without M regs, with and without PHA stimulation, are necessary controls for the interpretation of the experiment. To obtain accurate estimates of T cell numbers after coculture, each coculture should be performed in triplicate. The M regs and T cells should be cultured together for 3–5 days prior to analysis. 10. The cocultures should be harvested by scraping and vigorous pipetting. It is important to recover all the cells from a well into a single FACS tube. 11. To each sample add 20-ml FcR block, vortex, and incubate at 4°C for 30 min in darkness. 12. To each sample add 10 ml of anti-CD4 APC and 10 ml of anti-CD8 PE antibody. Incubate the samples at 4°C for 60 min in darkness. 13. To each sample add 50 ml of CountBright™ beads (see Note 3). 14. Set up the flow cytometer and adjust the compensation for a four colour staining with CFSE, PE, 7-AAD, and APC. Figure 2 illustrates a suitable gating strategy for the analysis of proliferation and absolute counting of CD4+ and CD8+ cells in M reg cocultures. 15. Five minutes prior to analysis of the sample with the flow cytometer, add 5 ml of 7-AAD to the sample and vortex. Keep the samples on ice and protected from light. In this protocol, the samples should not be washed at any point during their preparation. 16. Immediately prior to reading the sample, vortex it well. Collect at least 105 events through the “counting bead” gate. 17. The absolute number of T cells is calculated from the absolute number of beads and the ratio of T cells to counting beads. Division Index is a useful measure of T cell proliferation and can be derived from the data. 3.7. Comparator Macrophage Types

The literature describes many subtly different methods for the cultivation of particular human macrophage populations (Fig. 3). We have necessarily adapted these various published protocols to

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M1 M1 - Classical activation

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Fig. 3. States of macrophage activation. M1-polarised (classically activated) MFs are induced by IFN-g and LPS in vitro. M2a-polarised (alternatively activated) MFs are induced by IL-4 or IL-13. M2b (Type II) MFs arise through stimulation with immunoglobuincomplexes and LPS. M2c (deactivated) MFs are a heterogeneous group of MFs, induced by IL-10, TGF-b, inhibitory receptor ligation (CD200R, CD47-CD172a) or glucocorticoids.

produce macrophage populations that can be compared fairly: specifically, the comparator macrophages are all generated from positively isolated CD14+ peripheral blood monocytes and are cultured for 7 days in Cell+ coated plasticware at a density of 105 cells/cm2 (9). Resting macrophages: Culture for 7 days in RPMI supplemented with P/S, 2-mM GlutaMax, 20% FCS and 100-ng/ml M-CSF. On day 6, medium is changed to 5% FCS. M1-polarised (classically activated) macrophages: Culture for 7 days in RPMI supplemented with P/S, 2-mM GlutaMax, 20% FCS and 100-ng/ml M-CSF. On day 6, medium is changed to 5% FCS. One hundred nanograms per millilitre LPS and 20-ng/ ml IFN-g are added for the final 18 h of culture. M2a-polarised (alternatively activated) macrophages: Culture for 7 days in RPMI supplemented with P/S, 2-mM GlutaMax, 20% FCS and 100-ng/ml M-CSF. On day 6, medium is changed to 5% FCS and 20-ng/ml IL-4 added for the final 18 h of culture. M2b-polarised (immunoglobulin complex-stimulated) macrophages: A commercial human IVIG preparation (Sandoglobulin™, CSL Behring, Hattersheim am Main, Germany) is reconstituted in DPBS, extensively washed by centrifugal filtration and used to coat tissue culture surfaces by overnight incubation in a 0.1 M NaHCO3 solution. After washing the wells, monocytes are plated in RPMI containing 10% FCS and 100-ng/ml M-CSF, 2-mM GlutaMax and P/S. The cells are cultured for 6 days prior to 100-ng/ml LPS stimulation for the final 18 h of culture.

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M2c-polarised (dexamethasone-deactivated) macrophages: Culture for 7 days in RPMI supplemented with P/S, 2-mM GlutaMax, 20% FCS and 100-ng/ml M-CSF. On day 6, medium is changed to 5% FCS. Dexamethasone is added at final concentration 10 M−7 for the final 18 h of culture. IFN-g-stimulated macrophages: Culture for 7 days in RPMI supplemented with P/S, 2-mM GlutaMax, 10% FCS and 100-ng/ml M-CSF. On day 6, 20-ng/ml IFN-g is added for the final 18 h of culture.

4. Notes 1. For our experiments, male only, pooled human AB serum was sourced from Lonza. Serum is heat inactivated at 56°C for precisely 30 min before being cooled on ice. Aliquots are prepared and stored at −20°C for up to 3 months. Serum is defrosted on the day of use and any excess should not be used at a later date or refrozen. It is noted that the turbidity of human serum is much reduced by filtration of the complete M reg medium. 2. M regs have been derived from human PBMC obtained from a variety of sources, including whole blood, buffy coats and the leucocyte reduction system (LRS) chamber of the Trima SN system (Gambro BCT, Germany), which is used routinely to prepare thrombocyte concentrates for clinical purposes. When using PBMC concentrates, the dilution of the cells in DPBS prior to Ficoll separation should be appropriately adjusted. 3. The concentration of CountBright beads varies slightly between batches. It is essential to take note of the absolute number of beads used in particular experiments. 4.1. Conflict of Interest Statement

Prof. Fändrich holds the intellectual property rights to the clinical application of M regs.

References 1. Hutchinson JA, Govert F, Riquelme P et al. (2009) Administration of donor-derived transplant acceptance-inducing cells to the recipients of renal transplants from deceased donors is technically feasible. Clin Transplant 23: 140–145. 2. Hutchinson JA, Riquelme P, Brem-Exner BG et al. (2008) Transplant acceptance-inducing cells as an immune-conditioning therapy in renal transplantation. Transpl Int 21: 728–741.

3. Hutchinson JA, Brem-Exner BG, Riquelme P et al. (2008) A cell-based approach to the minimization of immunosuppression in renal transplantation. Transpl Int 21: 742–754. 4. Hutchinson JA, Roelen D, Riquelme P et al. (2008) Preoperative treatment of a presensitized kidney transplant recipient with donor-derived transplant acceptance-inducing cells. Transpl Int 21: 808–813.

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5. Chatenoud L (2008) The long and winding road towards induction of allograft tolerance in the clinic. Transpl Int 21: 725–727. 6. Warnecke G, Hutchinson JA, Riquelme P et al. (2009) Postoperative intravenous infusion of donor-derived transplant acceptance-inducing cells as an adjunct immunosuppressive therapy in a porcine pulmonary allograft model. Transpl Int 22: 332–341. 7. Brem-Exner BG, Sattler C, Hutchinson JA (2008) Macrophages driven to a novel state of

activation have anti-inflammatory properties in mice. J Immunol 180(1): 335–349. 8. Riquelme P, Govert F, Geissler EK et al. (2009) Human transplant acceptance-inducing cells suppress mitogen-stimulated T cell proliferation. Transpl Immunol 21(3): 162–165. 9. Riquelme P, Wundt J, Hutchinson JA et al. (2009) A refined characterisation of the NeoHepatocyte phenotype necessitates a reappraisal of the transdifferentiation hypothesis. Differentiation 77(3): 263–276.

Chapter 14 NKT and Tolerance Julien Diana, Lucie Beaudoin, Anne-Sophie Gautron, and Agnès Lehuen Abstract NKT cells are innate-like ab T cells that are conserved between humans and mice. They are distinct from conventional T cells as they recognize lipid antigens presented by the CD1d molecule. Most NKT cells expressed a highly restricted TCR repertoire and can be activated by a-galactosylceramide (a-GalCer) and detected by a-GalCer-loaded-CD1d tetramers. Upon activation, NKT cells respond in few hours by producing cytokines and stimulating many other cells of the innate and adaptive immune system. Over the last decade, many studies have analyzed the regulatory role of NKT cells that can either suppress or exacerbate immune functions. This chapter describes the tools and techniques required to study in vivo and in vitro the regulatory role of NKT cells in mouse as well as from human blood. Key words: NKT, a-GalCer, Immunoregulation, Suppressive, Cytokine, Dendritic cell

1. Introduction NKT cells are nonconventional ab T cells that are restricted by the monomorphic MHC class-I-like molecule CD1d presenting lipid ligands and not peptides (1, 2). A majority of NKT cells are named invariant (i) NKT cells as they expressed an invariant TCRa chain encoded by Va14-Ja18 genes in mice and Va24Ja18 genes in humans. iNKT cells exhibit a restricted Vb repertoire, Vb8, 7, and 2 in mice and Vb11 in humans. The other NKT cell subset is named as variable (v)NKT cells, since they express a more diverse TCR repertoire. Both subsets of NKT cells express markers usually associated with the NK lineage and with activated/memory phenotypes. Most of the studies on iNKT cells in the mouse have taken advantage of several mouse lines such as CD1d deficient mice that do not contain any CD1d restricted Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_14, © Springer Science+Business Media, LLC 2011

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T cells (3), Ja18 deficient mice that specifically lack iNKT cells (4) and mice transgenic for the iNKT cell TCR alpha chain (Va14-Ja18) that contain a higher frequency of iNKT cells (5, 6) as well as transgenic mice expressing the TCR of a vNKT cell hybridoma (7, 8). Another critical step has been the discovery of a specific iNKT cell agonist, a-GalCer (4). This reagent allows the specific activation of human and mouse iNKT cells and to analyze the events downstream of iNKT cell triggering such as the activation of dendritic cells (DC), NK cells, and conventional T and B cells. NKT cells are unique in their ability to release rapidly large amounts of cytokines such as IFN-g and IL-4 and they can influence a wide array of immune responses. Cytokine production by NKT and NK cells can be detected either directly by intracytoplasmic staining or indirectly by the cytokines released in the serum. Another major step in the analysis of iNKT cells has been the production of mouse (m)CD1d-aGalCer tetramer that recognizes specifically mouse and human iNKT cells (9, 10) and allows their detection and isolation from mouse lymphoid tissues as well as human blood. Regarding their function, many experimental in vivo and in vitro data as well as clinical observations support an immunoregulatory function of NKT cells. For example, increasing the frequency of iNKT cells in mouse models can prevent the development of autoimmune diseases such as type 1 diabetes and experimental autoimmune encephalomyelitis (11). The in vivo suppressive effect of iNKT cells was well characterized following the transfer of monoclonal antiislet T-cell population (12). In vitro coculture experiments further confirmed their suppressive effect and revealed the role of cell contact in this function (13). The main tools and techniques used to identify and study iNKT are described below.

2. Materials 2.1. Cell Preparation for iNKT Cell Analysis and Isolation

1. Buffer for cell isolation (CI buffer): RPMI 1640 (Gibco) containing 5% (v/v) fetal calf serum (FCS). Use at +4°C. 2. Red blood cell (RBC) lysis solution, Tris-Buffered ammonium chloride solution: 90% v/v 0.16 M NH4Cl + 10% v/v 0.17 M Tris-Hcl, pH 7.65, adjust to pH 7.2 with HCl. Prewarm at RT. 3. Collagenase P from Clostridium histolyticum (Roche diagnostics GmbH) at 1 mg/mL in DMEM (Dulbecco’s modified eagle’s medium, Gibco) (see Note 1). 4. Nylon cell strainers from BD Falcon, 40, 70, and 100 mm. 5. Ficoll: 40% stock solution of Ficoll 400 (Sigma) in Eurocolins (Fresenius Medical Care) is autoclaved 5 min at 110°C then diluted to make 23, 20, and 11% solutions and prewarm at RT before use.

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6. Cell dissociation solution nonenzymatic: (Sigma) prewarm at 37°C. 7. Heparin tube LH 170 IU (BD Vacutainer). 8. Leucosep 30 mL (Greiner-bio-one). 9. Ficoll-Paque PLUS (GE Healthcare). 10. Percoll™ (GE Healthcare), prewarm at RT. 2.2. Flow Cytometry for iNKT Cells

1. FACS buffer: phosphate-buffered saline (PBS) without calcium and magnesium pH 7.2, 2% FCS, 0.1% w/v NaN3. Use at +4°C. 2. a-Galactosylceramide (KRN7000 from Funakoshi Co or a-GalCer from Alexis) is dissolved in 0.5% (w/v) Tween-20, 0.9% NaCl at 95°C for 30 min then sonicate for 1 min in a glass tube. Store the a-GalCer stock solution (200 mg/mL) at +4°C (stable over 2 months). 3. mCD1d-a-GalCer tetramers [coupled to antigen presenting cell (APC) or PE fluorochrome for optimal detection sensitivity] 250 mg/mL in PBS 0.1% w/v NaN3, store at +4°C, stable for 4 months. 4. Phorbol 12 myristate 13-acetate (PMA) (Sigma): 1 mg/mL in ethanol. Ionomycin (Sigma): 0.5 mg/mL in DMSO. Brefeldin A (BA) (Sigma): 1 mg/mL in ethanol. All reagents are aliquoted and stored at −20°C. 5. Cytofix/Cytoperm Kit (BD Pharmingen). 6. Antimouse mAbs: anti-Fcg receptor (2.4G2); anti-TCR-b (H57), anti-NK1.1 (PK136), anti-CD4 (L3T4), anti-CD69 (H1.2F3), anti-CD44 (Ly44), anti-OX40 (OX86), anti-IFN-g (XMG1-2), anti-IL-4 (BVD6), anti-IL-10 (JES5-16E3), and anti-IL-17 (TC11-18H10) all from BD Pharmingen and antiIL-13 (Bio13a) from eBioscience. 7. Antihuman mAbs, anti-Va24, and anti-Vb11 (Beckman coulter); anti-CD3 (HIT3a) (Myltenyi); 6B11mAb, antiCD161 (DX12), anti-CD4 (SK3), anti-CD8 (SK1) (BD Pharmingen), anti-IFN-g (4S.B3), anti-IL-4 (8D4-8), and anti-IL-17 (eBio64DEC17) (eBioscience).

2.3. Mouse and Human iNKT Cell Purification and Expansion and Mouse Reconstitution

1. Buffer for cell purification (CP buffer): RPMI 1640, 5% FCS, 0.5 mM EDTA (Gibco). Use at +4°C. 2. Dynal mouse CD4 Negative Isolation Kit (Dynal Invitrogen). 3. Magnets: Dynal MPC-15 for 1–15 mL or MPC-50 for 15–50 mL samples. 4. Anti-APC microbeads (Miltenyi). 5. LS Column (Miltenyi).

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6. MACS Separator (MidimMACS, Miltenyi). 7. Anti-NK1.1 (PK136 mAb) purified and aliquots are stored sterile in PBS at −20°C. 8. Complete RPMI medium: L-glutamax RPMI 1640, 10% FCS, 1% PS, 1 mM 2-ME, 1 mM sodium pyruvate (Gibco), 1× nonessential acid amines (NEAA) (Gibco). 9. C1R LCL cells engineered with hCD1d (14) are grown in complete RPMI medium with 0.5 mg/mL G418. C1R wildtype cells are used as controls. 2.4. In Vivo and In Vitro iNKT Cell Function

1. Complete RPMI medium: L-glutamax RPMI 1640, 10% FCS, 1% PS, 1 mM 2-ME, 1 mM sodium pyruvate, 1× NEAA. 2. Complete IMDM: DMEM (Gibco), 10% FCS, 1% PS, 1 mM 2-ME, 10 mM HEPES (Gibco), 100 mM L-glutamine (Gibco). 1 mM sodium pyruvate, 1× NEAA. 3. 0.5 mg/mL PHA (Sigma), 0.3 mg/mL G418 (Invitrogen), 0.5 mg/mL Hygromycin B (Invitrogen). 4. Recombinant human (rh)IL-2 (R&D) reconstituted in PBS at 1 mg/mL crystallized bovine serum albumin (BSA) (Gibco), aliquots stored at −80°C. 5. Specific peptide 1040–51 dissolved in DMSO then diluted in PBS to 10% DMSO final solution, aliquots at 1 mg/mL stored at −80°C. 6. Mitomycin C (Sigma) dissolved in PBS at 1 mg/mL, filtered on 0.2 mm, aliquots stored at −20°C. 7. Nunc tissue culture inserts 0.2-mm Anopore membrane (NuncBrand Products). 8. Carboxyfluorescein succinimidyl ester (CFSE) (Sigma) dissolved in DMSO at 50 mM. Store aliquots at −20°C in the dark. 9. 3H-Thymidine (3H-TdR) (1 mCi/well, Ge Healthcare). Store at +4°C. 10. The mouse 58ab-hybridoma cells was generated by cotransfection with the cDNA coding for the human invariant Va24JaQ and Vb11 (15).

2.5. Mouse Strains (Fig. 1)

1. CD1d deficient mice: these mice do not express the CD1d molecule and therefore they do not contain any NKT cells, neither iNKT cells nor other CD1d-restricted T cells that could express ab or gd TCR (3). These mice are available on C57BL/6, BALB/c, and NOD backgrounds. 2. Ja18 deficient mice: these mice do not express the Va14Ja18 alpha chain and therefore do not contain any iNKT cell.

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Fig. 1. Frequency of iNKT cells in various mouse strains. Cells from spleen of adult females (8–16 weeks old) were stained with a-Gal Cer-loaded mCD1d-tetramer and then stained with anti-TCR-b and anti-CD19 mAbs. B cells were gated out to remove some background staining.

These mice are available on C57BL/6 (4) and NOD backgrounds (16). 3. Va14 transgenic mice: these mice overexpress the Va14Ja18 alpha chain as a transgene and therefore they contain a higher frequency of iNKT cells. These transgenic mice are available on C57BL/6 and NOD backgrounds (6, 17). 4. Variable TCR transgenic mice: these mice contain a high frequency of vNKT cells. These vNKT cells express Va3.2/Vb9 TCR chains isolated from a CD1d-specific T-cell hybridoma. These transgenic mice are available on C57BL/6 and NOD backgrounds (8, 11). 5. pLck-CD1d mice: these mice express the mouse CD1d as a transgene. CD1d is specifically expressed in the thymus due to the proximal Lck promoter. When crossed to CD1d−/−, these mice express CD1d exclusively in the thymus allowing iNKT cell positive selection without any CD1d expression in peripheral tissues. These mice are available on C57BL/6 (18) and NOD backgrounds (19). A similar transgenic mouse line has been generated with human CD1d transgene (20).

3. Methods 3.1. Cell Preparation for iNKT Cell Analysis and Isolation 3.1.1. Mouse Spleen and Lymph Nodes

1. Place tissue to be processed in a Petri dish (3 cm in diameter) containing 5 mL CI buffer. 2. Pass the tissue through a 40-mm cell strainer by using a plunger and collect cells in a 15-mL conical tube. 3. Wash cells by adding 9 mL of CI buffer and centrifuge at 500 × g for 5 min at +4°C. 4. Resuspend cell pellet in RBC lysis solution (1 mL/spleen or 0.5 mL/lymph nodes) and incubate 2 min at RT.

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5. Wash cells in 10 mL CI buffer and resuspend pellet in 1 mL CI buffer and count live cells. 3.1.2. Mouse Liver

1. Eliminate peripheral blood lymphocyte contamination by transcardiac perfusion of mice with ice-cold PBS. 2. Transfer liver into a 40-mm cell strainer in a six-well tissue culture plate containing 5 mL of CI buffer. 3. Grind liver three times successively in three wells and pull the 15 mL in a 50-mL Falcon tube and complete the tube to 50 mL then centrifuge at 200 × g for 5 min at +4°C. 4. Transfer the supernatant into a 50-mL Falcon tube and centrifuge at 400 × g for 10 min at +20°C. 5. Resuspend cell pellet in 14 mL of 35% Percoll diluted in RPMI, transfer into a 15-mL Falcon tube, and centrifuge at 850 × g for 25 min at +20°C with no brake. 6. Resuspend cell pellet in RBC lysis solution (2 mL) and incubate 2 min at RT. 7. Wash cells in 10 mL of CI buffer (centrifuge at 500 × g for 5 min at +4°C), resuspend the pellet in 1 mL CI buffer, and count live cells.

3.1.3. Mouse Pancreatic Islets

1. Preparation of the pancreas. The animals are sacrificed and the abdomen opened. Uncover the bile duct by displacing the duodenum and the liver. Clamp off the bile duct at its entrance to the duodenum. Free the upper part of the bile duct by dissection to the liver. Inject collagenase P solution (3 mL) through the common bile duct into the pancreas. Remove the distended pancreas. Transfer the pancreas into a 50-mL Falcon tube containing 2 mL of collagenase P solution. Incubate for 10 min at 37°C in a water bath and vigorously tap three times. 2. Isolation of islets. Add about 30 mL of CI buffer and suspend the tissue. Perform the following steps at +2 to 8°C. Centrifuge at 1,800 × g for 1 min, discard the supernatant, and wash four times with CI buffer. After the first wash, transfer the pellet in 14 mL polystyrene round-bottom Falcon tube. After the last wash, remove the supernatant completely and dry the pellet on absorbent paper. Resuspend the pellet in 1 mL Ficoll stock solution, 40%. Load on top of this Ficoll suspension a discontinuous density gradient of 23% (1.5 mL), 20% (1.5 mL), and 11% (1.5 mL) Ficoll solutions. Centrifuge 17 min at 1,065 × g with no brake at RT. The islets are harvested from both interfaces between 11 and 20% and between 20 and 23%. Wash the islets in CI buffer (centrifuge at 805 × g for 5 min at RT).

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3. Isolation of single cells from islets. Wash islets suspension with PBS (centrifuge at 805 × g for 5 min at +4°C). Resuspend the pellet in 1 mL of Cell Dissociation Solution and incubate 10 min at 37°C in a water bath. Sharply tap the tube against the palm of your hand to dislodge the cells. Add 10 mL of CI buffer and pipet repeatedly to dissociate clumps. Wash the cell suspension (centrifuge at 500 × g for 5 min at +4°C), resuspend pellet in 1 mL CI buffer, and count live cells. 3.1.4. Mouse Central Nervous System

1. Central nervous system (CNS) dissection. Eliminate peripheral blood lymphocyte contamination by transcardiac perfusion of mice with ice-cold PBS. Collect brain and spinal cord in Petri dish (3 mL of diameter) in CI buffer. 2. CNS Dissociation. Dissociate tissue by gently grinding brain and spinal cord over a 100 and 70 mm nylon cell strainer sequentially in CI buffer, collect cells in a 50-mL Falcon tube. Wash cells twice in CI buffer (centrifuge at 500 × g for 5 min at +4°C). 3. Percoll purification. Prepare 50 mL Falcon tubes containing 20 mL of 70% Percoll (diluted in RPMI). Resuspend cells in 25 mL of 30% Percoll and overlay over the 70% Percoll. Centrifuge at 1,800 × g for 20 min at RT with no brake. The tubes contain a first ring of myelin debris and a pellet of large tissue fragments. The mononuclear cells reside at the 30–70% interface. Collect 3–4 mL, transfer in a 15-mL Falcon , and wash with CI buffer (centrifuge at 805 × g for 5 min at +4°C). Wash cells in CI buffer (centrifuge at 500 × g for 5 min at +4°C), resuspend pellet in 1 mL CI buffer, and count live cells.

3.1.5. Human Blood

1. Collect peripheral blood in heparin containing tubes. 2. Put 15–20 mL of Ficoll-Paque PLUS in a Leucosep tube of 30 mL and let the Ficoll pass through the membrane by 800 × g for 2 min. 3. Remove the excess of Ficoll and put until 35 mL of blood above the membrane. 4. Centrifuge at 660 × g for 20 min at 20°C. 5. PBMCs are located in the ring over the membrane. Collect PBMCs and transfer them into a 50-mL Falcon tube. 6. Wash the cells by completing the 50 mL Falcon tube with PBS and centrifuge at 666 × g for 10 min at 20°C. 7. Count the cells.

3.2. Flow Cytometry for iNKT Cells 3.2.1. Surface Staining for Mouse iNKT Cells

1. Stain 4 × 106 cells with 20 ml a-GalCer-loaded mCD1dtetramer diluted at the optimal concentration in FACS buffer, incubate 1 h at RT protected from light. 2. Wash cells (centrifuge at 500 × g for 5 min at +4°C) and block Fcg receptors with 20 ml of 2.4G2 mAb (100 mg/mL) for 15 min at +4°C.

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3. Wash cells and stain with 20 ml of anti-TCR-b mAb and other surface molecule-specific mAbs for 15 min at +4°C. The amount of mAb is determined by tittering each of them in separate staining. 4. Wash cells and resuspend the pellet in 300 ml of FACS buffer. 1. Stain 4 × 106 cells either (a) with 20 ml a-GalCer-loaded mCD1d-tetramer at the optimal concentration in FACS buffer and incubate 1 h at RT protected from light or (b) with 20 ml anti-Vb11mAb for 15 min at +4°C or (c) with 20 ml 6B11 mAb and incubate 30 min at +4°C.

3.2.2. Surface Staining for Human iNKT Cells

2. Wash with FACS buffer, centrifuge at 500 × g for 5 min at +4°C. 3. Stain with 20 ml of anti-TCR Va24 mAb [for (a) and (b)] or anti-CD3 mAb [for (c)] by incubating cells 15 min at +4°C. 4. Wash with FACS buffer and resuspend the pellet in 500 ml FACS buffer. 5. Human circulating iNKT cells are identified as a-GalCerloaded mCD1d-tetramer+ Va24+ cells, as Va24+ Vb11+ cells or as 6B11+ CD3+ cells (see Notes 2 and 3) (Fig. 2). 1. To increase cytokine detection, mouse cells can be activated for 4 h (overnight for human iNKT cells) by PMA (100 ng/mL), ionomycin (0.5 mg/mL) in the presence of BA (10 mg/mL) at 37°C in complete RPMI.

3.2.3. Intracytoplasmic Staining

2. Wash 5 × 106 cells with FACS buffer, cells are surface stained, then wash and resuspend in 500 ml of Cytofix under vortex agitation and incubate 8 min at +4°C protected from light.

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Fig. 2. Frequency of iNKT cells in human blood sample. Human PBMC were stained either with (a) CD1d-a-GalCertetramer or (b) anti-Vb11mAb or (c) 6B11 mAb. Then cells were stained with anti-TCR Va24 mAb [for (a) and (b)] or anti-CD3 mAb (for (c)). Human circulating iNKT cells are identified as CD1d-a-GalCer-tetramer+ Va24+ cells, as Va24+ Vb11+ cells, or as 6B11+ CD3+ cells.

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4. Stain with anticytokine-specific mAb diluted in Cytoperm buffer, incubate 30 min at +4°C protected from light. 5. Wash with Cytoperm buffer and resuspend the pellet in 300 ml of FACS buffer. 3.3. Mouse and Human iNKT Cell Purification and Expansion and Mouse Reconstitution

1. Prepare a single-cell suspension from spleen or lymph nodes using standard methods.

3.3.1. Enrichment of Mouse iNKT Cells by Negative Selection

2. For isolation of mouse double negative and CD4+ T Cells, use Dynal mouse CD4 Negative Isolation Kit according to manufacturer’s instructions.

3.3.2. Positive Selection of Mouse iNKT Cells

1. Staining of iNKT cells with mCD1d-a-GalCer-APC tetramer (as described in Subheading 3.2.1).

Centrifuge cells at 500 × g for 5 min and resuspend pellet in CP buffer, count cells, and adjust to 1 × 107 cells in 100 ml.

2. Resuspend cell pellet in 90 ml of CP buffer per 107 total cells. 3. Add 10 ml/107 cells of anti-APC microbeads and incubate for 15 min at +4°C. 4. Wash cells in 2 mL/107 cells of CP buffer and resuspend in 500 ml of CP buffer. 5. Place LS column in the MACS Separator, rinse it with 3 mL of CP buffer, and apply cell suspension. 6. Wash LS column with 3 × 3 mL of CP buffer and discard unlabeled cells, which pass through. 7. Add 5 mL of CP buffer onto the LS column then remove it from the MACS Separator and place it on a collection tube. 8. Immediately flush out fraction with the magnetically labeled cells by firmly applying the plunger supplied with the column. The supernatant contains the iNKT cells. 3.3.3. Adoptive Transfer of iNKT Cells into Recipient Mice

1. In vivo depletion of NK cells. To promote the “engraftment” of iNKT cells, recipient mice were previously depleted in NK cells by treatment with PK136 mAb (see Note 4). To reconstitute iNKT cell population in young mice, 2-week-old NK1.1+/+ mice are i.p. injected with PK136 mAb for several days: day 0: 50 mg, day 2: 50 mg, day 9: 50 mg, day 17: 100 mg (for each in 100 ml PBS). iNKT cells are transferred on day 3 after the beginning of PK136 mAb treatment. When possible, iNKT cells are obtained from NK1.1-mice to avoid NK1.1 triggering on iNKT cells. To transfer iNKT cells in adult complete mice, NK cell depletion can be obtained by oneshot treatment (0.5 mg in 200 ml of PBS/mouse) 2 days prior iNKT cells transfer.

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2. For iNKT cell reconstitution, mice are i.v. injected with 1 × 106 to 5 × 106 purified iNKT cells in 100 ml PBS per mouse. 3.3.4. Generation of Human iNKT Cell Clones and Lines

1. As previously described by Abrignani et al. (21), human iNKT cells can be obtained by two methods: (1) cell sorting and cloning ex vivo; (2) expansion of iNKT polyclonal cell lines in vitro with the specific antigen a-GalCer. iNKT cell clones can be obtained by cloning these cell lines at limiting dilution. 2. iNKT cell clones or lines are propagated by restimulation every 15–20 days in complete RPMI medium with a mixture of irradiated allogeneic feeder cells at 1 × 106 cells/mL, PHA (0.5 mg/mL), and rhIL-2 (50 U/mL). 3. To verify the antigen specificity of the iNKT cell clones, 2.5 × 104 cells are plated in U-bottom 96-well plates with 5 × 104 of either C1R-CD1d or C1R wildtype cells in the presence of 50 ng/mL of a-GalCer. After 48 h, antigenspecific iNKT cell activation is determined by measuring the concentration of IFN-g and IL-4 in the culture supernatant by ELISA.

3.4. In Vivo and In Vitro iNKT Cell Function 3.4.1. In Vivo, a-GalCer Injection to Mice (see Note 5)

3.4.2. In Vitro Mouse iNKT Cell Assays

To activate iNKT cells in vivo, their specific ligand, a-GalCer (2 mg in 100 ml PBS), can be injected simultaneously i.v. and i.p. to mice. Two hours later cytokine (IL-4 and IFN-g) expression by spleen iNKT cells can be analyzed by intracytoplasmic staining without in vitro activation. Six hours later cytokines released in the serum can be measured by ELISA and IFN-g produced by NK cells can be directly evaluated by intracytoplasmic staining (see Note 6). 1. a-GalCer stimulation. Prepare a single-cell suspension using standard methods and resuspend in complete RPMI and count. Incubate 5 × 105 cells with a-GalCer (100 ng/mL) at 37°C in flat bottom 96-well plate. After 24–72 h, harvest 100 ml of supernatants to measure cytokine released by ELISA (see Note 7). Add 3 H-thymidine (1 mCi/well) for 4–10 h to evaluate cell proliferation. 2. In vitro suppressive assay. (a) APCs were obtained from Ca−/− NOD mice. Prepare a splenocyte single-cell suspension using standard methods. APC are either irradiated with 3,000 rad or treated 40 min at 37°C with mitomycin C at 70 mg/mL for 107 cells in PBS and washed four times with cold PBS. (b) Responder CD4+ T cells: Antipancreatic islet antigen BDC2.5 CD4 T cells were used as responder cells (13).

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They were isolated from the spleen of transgenic BDC2.5 Ca−/− NOD mice (as described in Subheading 3.1.1). To purify naïve BDC2.5 T cells, cells were labeled with antiCD4 and anti-CD62L mAbs and sorted as CD62L+ CD4+ cells. (c) iNKT cells (see Note 8): iNKT cells were first enriched by negative selection, then labeled (described in Subheading 3.3.2) and purified by cell-sorting as TCR-b+ and mCD1d-a-GalCer tetramer+ cells. To avoid TCR triggering, spleen iNKT cells can also be purified by cellsorting as CD5+ NK1.1+ cells (see Note 9). In vitro culture: Incubate responder T cells (5 × 104 cells/ well) with APC (5 × 104 cells/well) and variable numbers of iNKT cells (from 1.5 × 103–4 × 105) in complete IMDM for 120 h at 37°C in U-bottom 96-well culture plate. Add rhIL-2 (25 U/mL) and peptide 1040-51 (10 ng/ mL) to stimulate responder T cells. a-Galcer (50 ng/ mL), or vehicle, was added into some wells. Transwell culture: Nunc tissue culture inserts 0.2 mm Anopore membrane. Incubate in the lower chamber responder T cells (2.5 × 104) with APC (2.5 × 104) with rhIL-2 and peptide 1040–51. Place the upper chambers avoiding any bubble. In the upper chamber, add iNKT cells (1 × 105) with APC (2.5 × 104) at the beginning of the culture period. Add a-GalCer 50 ng/mL. Evaluation of cytokine production by intracytoplasmic staining: After 5 days of culture, add for 4 h PMA (0.5 mg/mL) plus ionomycin (500 ng/mL) and BA (10 mg/mL) in culture (Fig. 3). Then perform anti-IFN-g intracytoplasmic staining as described in Subheading 3.2.3.

IFN–g FSC Peptide

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+

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iNKT cells Lower Ch.

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Fig. 3. iNKT cell inhibition of T-cell differentiation is contact dependent. Sorted CD62L+ BDC2.5 responder T cells were incubated with their specific peptide (peptide 1040–51), APC, rIL-2, and a-GalCer. As indicated, iNKT cells were added either in the lower chamber in contact with APC and BDC2.5 T cells or separated in the upper chamber of the Transwell. On day 5, IFN-g production was analyzed by intracytoplasmic staining. Dot plots represent BDC2.5 T cells (gated as CD4+ Thy1.1+ cells) and the values correspond to the frequency of IFN-g producing cells.

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Evaluation of T cell proliferation by CFSE: Before the culture, responder T cells were labeled with CFSE (see Note 10). Cells at 50 × 106/mL in PBS 0.5% w/v BSA were incubated with CFSE (5 mM, final concentration), for 8 min at 37°C. To block the reaction, cold PBS 0.5% BSA was added for 5 min at +4°C and cells were washed twice in complete IMDM. 3. Coactivation of mouse DC and iNKT cells. Purified iNKT cells (see Subheading 3.3.2) can be cultured in vitro with DC to investigate the activation of both cell types. Total DC population is first positively selected by magnetic bead separation according to the manufacture instruction (Pan DC MicroBeads, Miltenyi Biotech) and then sorted as 120G8+ CD11clow cells for plasmacytoid DC (pDC) and as 120G8− CD11c+ for conventional DC. The purity of sorted cell populations must be >95%. Culture with 105 pDCs and 104 iNKT cells is performed in 96-well U-bottom plate in 200 ml of complete RPMI medium with 20 ng/mL a-GalCer or vehicle. Various times after culture, supernatants are recovered for cytokine analysis by ELISA. As example, we demonstrated that iNKT cells and pDCs cooperate in vitro through OX40–OX40Ligand interaction (16). Cocultures were performed with sorted cell populations in the presence of virus. pDC–iNKT cell interaction resulted in IFN-a production only when iNKT cells were isolated from the pancreas and not from the spleen. IFN-a production was measured by ELISA (Mouse Interferon Alpha ELISA Kit, PBL) in the supernatant after 3 days of culture. Importantly, this type I IFN production was abolished by adding anti-OX40L mAb or OX40-Ig molecule. 3.4.3. In Vitro Human iNKT Cell Assays

1. Human B lymphocyte and monocyte purification. For monocyte (CD14+ cells) purification, PBMCs are first incubated with anti-CD14 mAb conjugated with magnetic beads (Miltenyi) and positively selected, then B cells (CD19+ cells) are purified from remaining cells using anti-CD19 mAb conjugated with magnetic beads (Miltenyi). 2. Human B cell and monocyte assay. Purified B cells (105 cells) or monocytes (0.5 × 105 cells) are incubated in 0.2 mL of complete RPMI, either alone or with iNKT cell clones or hybridomas at 1:1 and 1:2 ratios, respectively, in

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U-bottom 96-well plate. Duplicates are stimulated with concentrations of a-GalCer ranging from 0 to 900 ng/mL or with equal amounts of vehicle. After 48 h of culture, IFN-g and IL-4 are detected by ELISA in the supernatants of iNKT cell clone cultures, and IL-2 is detected in the supernatants of mouse T-hybridoma cell cultures.

4. Notes 1. The optimal concentration of each Collagenase P batch has to be determined. 2. Stain at least 4 × 106 cells per individual and collect all the PBMC because of the low frequency of iNKT cells. 3. Using the Va24+ Vb11+ combination without using a-GalCerloaded mCD1d-tetramer could overestimate the frequency of iNKT cells. 4. PK136 mAb treatment depletes NK cells, whereas iNKT cells are not depleted but downregulate the NK1.1 molecule on their cell surface. Not all strains express the NK1.1 molecule. C57BL/6 are NK1.1+ as well as NK1.1+ congenic NOD and BALB/c mice. 5. The same a-GalCer stock solution is used for both in vivo and in vitro assays. 6. Cytokine production by iNKT cells measured by intracytoplasmic staining at later time points than 4 h after a-GalCer injection is not fully accurate due to the downregulation of iNKT cell TCR on cell surface and the difficulty of iNKT cell detection by tetramer-a-GalCer-mCD1d staining. 7. Usually the maximum for IL-4 and IFN-g production is observed after 24 and 72 h, respectively (for both mouse and human iNKT cells). 8. In order to get large number of iNKT cells, they were usually purified from the spleen of Va14Ca−/− transgenic mice. 9. Using mouse expressing the NK1.1 molecule, CD5+ NK1.1+ cells from spleen (but not lymph nodes) of Va14 Ca−/− transgenic mice correspond to iNKT cells (purity >90%). From wildtype mice, CD5+ NK1.1+ cell subset contain up to 60% of iNKT cells. 10. CFSE labeling usually induced some cell death and around 50% of the cells are lost.

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References 1. Kronenberg, M. and L. Gapin (2002) The unconventional lifestyle of NKT cells. Nat Rev Immunol. 2(8): p. 557–68. 2. Bendelac, A., P.B. Savage and L. Teyton (2007) The biology of NKT cells. Annu Rev Immunol. 25: p. 297–336. 3. Mendiratta, S.K., W.D. Martin, S. Hong, et al. (1997) CD1d1 mutant mice are deficient in natural T cells that promptly produce IL-4. Immunity. 6(4): p. 469–77. 4. Cui, J., T. Shin, T. Kawano, et al. (1997) Requirement for Valpha14 NKT cells in IL-12-mediated rejection of tumors. Science. 278(5343): p. 1623–6. 5. Bendelac, A., R.D. Hunziker and O. Lantz (1996) Increased interleukin 4 and immunoglobulin E production in transgenic mice overexpressing NK1 T cells. J Exp Med. 184(4): p. 1285–93. 6. Lehuen, A., O. Lantz, L. Beaudoin, et al. (1998) Overexpression of natural killer T cells protects Valpha14- Jalpha281 transgenic nonobese diabetic mice against diabetes. J Exp Med. 188(10): p. 1831–9. 7. Duarte, N., M. Stenstrom, S. Campino, et al. (2004) Prevention of diabetes in nonobese diabetic mice mediated by CD1d-restricted nonclassical NKT cells. J Immunol. 173(5): p. 3112–8. 8. Skold, M., N.N. Faizunnessa, C.R. Wang, et al. (2000) CD1d-specific NK1.1+ T cells with a transgenic variant TCR. J Immunol. 165(1): p. 168–74. 9. Benlagha, K., A. Weiss, A. Beavis, et al. (2000) In vivo identification of glycolipid antigenspecific T cells using fluorescent CD1d tetramers. J Exp Med. 191(11): p. 1895–903. 10. Matsuda, J.L., O.V. Naidenko, L. Gapin, et al. (2000) Tracking the response of natural killer T cells to a glycolipid antigen using CD1d tetramers. J Exp Med. 192(5): p. 741–54. 11. Mars, L.T., A.S. Gautron, J. Novak, et al. (2008) Invariant NKT cells regulate experimental autoimmune encephalomyelitis and infiltrate the central nervous system in a CD1d-independent manner. J Immunol. 181(4): p. 2321–9.

12. Beaudoin, L., V. Laloux, J. Novak, et al. (2002) NKT cells inhibit the onset of diabetes by impairing the development of pathogenic T cells specific for pancreatic beta cells. Immunity. 17(6): p. 725–36. 13. Novak, J., L. Beaudoin, T. Griseri, et al. (2005) Inhibition of T cell differentiation into effectors by NKT cells requires cell contacts. J Immunol. 174(4): p. 1954–61. 14. Brossay, L., M. Chioda, N. Burdin, et al. (1998) CD1d-mediated recognition of an alpha-galactosylceramide by natural killer T cells is highly conserved through mammalian evolution. J Exp Med. 188(8): p. 1521–8. 15. Casorati, G., A. Traunecker and K. Karjalainen (1993) The T cell receptor alpha beta V-J shuffling shows lack of autonomy between the combining site and the constant domain of the receptor chains. Eur J Immunol. 23(2): p. 586–9. 16. Diana, J., T. Griseri, S. Lagaye, et al. (2009) NKT Cell-plasmacytoid dendritic cell cooperation via OX40 controls viral infection in a tissuespecific manner. Immunity. 30(2): p. 289–99. 17. Laloux, V., L. Beaudoin, C. Ronet, et al. (2002) Phenotypic and functional differences between NKT cells colonizing splanchnic and peripheral lymph nodes. J Immunol. 168(7): p. 3251–8. 18. Wei, D.G., H. Lee, S.H. Park, et al. (2005) Expansion and long-range differentiation of the NKT cell lineage in mice expressing CD1d exclusively on cortical thymocytes. J Exp Med. 202(2): p. 239–48. 19. Novak, J., L. Beaudoin, S. Park, et al. (2007) Prevention of type 1 diabetes by invariant NKT cells is independent of peripheral CD1d expression. J Immunol. 178(3): p. 1332–40. 20. Schumann, J., P. Pittoni, E. Tonti, et al. (2005) Targeted expression of human CD1d in transgenic mice reveals independent roles for thymocytes and thymic APCs in positive and negative selection of Valpha14i NKT cells. J Immunol. 175(11): p. 7303–10. 21. Abrignani, S., E. Tonti, G. Casorati, et al. (2009) B cell helper assays. Methods Mol Biol. 514: p. 15–26.

Chapter 15 Antiinflammatory and Immunosuppressive Functions of Mast Cells Janet Kalesnikoff and Stephen J. Galli Abstract Through the release of biologically active products, mast cells function as important effector and immunoregulatory cells in diverse immunological reactions and other biological responses; for example, mast cells promote inflammation and other tissue changes in immunoglobulin E (IgE)-associated allergic disorders, as well as in certain innate and adaptive immune responses that are thought to be independent of IgE. Despite the mast cell’s well-deserved reputation as a promoter of inflammation, others and we have used bone marrow-derived cultured mast cell (BMCMC) engrafted mast cell-deficient c-kit-mutant mice (so-called “mast cell knock-in” mice) to show that mast cells can also have important antiinflammatory and immunosuppressive functions in vivo. An early study showed that mast cells can contribute to susceptibility to ultraviolet B (UVB)-induced immunosuppression in one model of contact hypersensitivity (CHS), through effects mediated at least in part by histamine. Subsequently, it was reported that mast cells can mediate negative immunomodulatory effects following Anopheles mosquito bites, and in peripheral tolerance to skin allografts; however, the mechanism(s) by which mast cells mediate immunosuppressive functions in these two studies remains to be elucidated. Finally, we showed that mast cells and mast cell-derived IL-10 can limit the magnitude of and promote the resolution of certain CHS responses, and suppress the inflammation and skin injury associated with innate cutaneous responses to chronic low-dose UVB irradiation. This chapter outlines the generation of BMCMCs, a powerful model system commonly used to: (1) identify potential mast cell mediators in vitro; (2) study the mechanisms of mast cell activation and mediator release in response to specific stimuli in vitro; and (3) engraft mast celldeficient mice to study the effector and immunoregulatory roles of mast cells or specific mast cell mediators in diverse immunological responses in vivo. Key words: BMMCs, BMCMCs, Mast cell-deficient mice, Engraftment, IL-10

1. Introduction Mast cells are derived from hematopoietic stem cells, but they do not ordinarily circulate in the mature form; instead, mast cells acquire their mature phenotype locally, following the migration Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_15, © Springer Science+Business Media, LLC 2011

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of their precursors to the vascularized tissues or serosal cavities in which the mast cells will ultimately reside (1–3). Mast cells participate in a wide variety of physiological and pathological processes as a result of their activation by any of an array of receptors. For example, they function as key effector cells in immunoglobulin E (IgE)-associated immune responses, including allergic disorders and certain protective immune responses to parasites. However, mast cells can be activated to perform important effector and immunoregulatory functions by many mechanisms that are independent of IgE (e.g., mast cells can be activated directly by pathogens, microbial products, endogenous and exogenous peptides, cytokines, and other inflammatory mediators) (1, 4). Activated mast cells can release a diverse array of potent biologically active products (1, 3, 4). Many of these products are considered to be proinflammatory, in that they can elicit, individually or in concert, various features of inflammation (e.g., vasodilation, plasma extravasation, and the recruitment and activation of granulocytes, T cells, B cells, dendritic cells, and monocytes) and promote tissue remodeling. However, upon appropriate activation, mast cells can release products that are known to have several antiinflammatory or immunosuppressive properties, such as IL-10 and TGF-b. Thus, mast cells have the potential to exert positive or negative immunomodulatory functions on immune cells (i.e., influence the development, recruitment, survival, phenotype, or function of immune cells) and thereby enhance or suppress the initiation, magnitude, and/ or duration of certain immune responses (1, 2). Although mice that specifically lack only mast cells have not been reported, c-kit-mutant mice that are profoundly deficient in mast cells are often used to assess the contributions of mast cells or specific mast cell products to diverse biological responses in vivo. The mast cell deficiency of these mice can be selectively “repaired” through the adoptive transfer of genetically compatible, in vitroderived mast cells [e.g., bone marrow-derived cultured mast cells (BMCMCs) from congenic wildtype mice or various transgenic or mutant mice, BMCMCs transduced with short-hairpin RNA to reduce expression of proteins of interest, embryonic stem cell-derived cultured mast cells (ESCMCs), fetal liver-derived mast cells (FLMCs), etc.]. Because c-kit mutations affect lineages other than mast cells (see Note 1), it is essential to perform experiments on these so-called “mast cell knock-in mice” to assess the extent to which differences in the biological responses of c-kit-mutant mice compared with wildtype mice are due to the lack of mast cells in the c-kit-mutant mice (1, 2, 4). Recently, transgenic mice have been generated that express Cre recombinase under the control of the promoter of the gene encoding mouse mast cell

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protease 5 (Mcpt5) (5) or the gene encoding baboon a-chymase (6). Although the phenotypic features of such mice (as expressed in mast cells and possibly other cell types) remain to be fully characterized, mice with confirmed mast cell-specific Cre expression and mice with inducible mast cell-specific Cre expression may well become powerful genetic models for investigating the contributions of mast cells or mast cell-specific products to health and disease. Mast cell knock-in mice have been used to identify antiinflammatory and immunosuppressive roles of mast cells during specific immune responses in vivo. An early study showed that mast cells, through effects mediated at least in part by histamine, contribute to the ability of ultraviolet B (UVB) irradiation of the skin to induce systemic immunosuppression of contact hypersensitivity (CHS) responses to the hapten TNCB (2,4,6-trinitrochlorobenzene) (7). Mast cells have also been shown to mediate immunosuppressive functions following Anopheles mosquito bites (8), and in peripheral tolerance to skin allografts (which requires the participation of CD4+CD25+FoxP3+ TReg cells) (9). The mechanism(s) by which mast cells mediate immunosuppressive functions in these two studies remains to be elucidated; however, IL-10 was implicated as a mechanism of immunosuppression in each of these studies, acting either in the draining lymph nodes (8) or at the sites of skin allografts (9). In another study, mast cells and mast cell-derived IL-10 were shown to limit the magnitude of and promote the resolution of CHS responses induced in response to the hapten DNFB (2,4-dinitro-1-fluorobenzene) or urushiol (the haptencontaining sap of poison ivy or poison oak), and to suppress the inflammation and tissue pathology associated with innate cutaneous responses to chronic low-dose UVB irradiation (10). Defining the mechanisms by which mast cells or specific mast cell mediators (such as IL-10 or other mediators) can suppress, rather than enhance, features of specific innate or acquired immune responses and identifying the signals that can elicit such distinct mast cell functions are important goals for future research. Human mast cell populations and mouse BMCMCs, as well as cultured mast cells derived from other hematopoietic tissues (such as fetal liver) or from embryonic stem cells, are powerful tools to investigate the mechanisms by which mast cells might influence various immunological or other biological responses in vitro. BMCMCs, which can easily be expanded to produce large numbers of mast cells in vitro, are the cell type most commonly used for in vitro studies of mast cell activation events and engraftment into c-kit-mutant mast cell-deficient mice. This chapter will describe in detail the generation and characterization of BMCMCs and current protocols for the adoptive transfer of BMCMCs into mast cell-deficient mice.

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2. Materials 2.1. Generation of BMCMCs from Mouse Bone Marrow

1. Flushing medium: Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 1% antibiotic-antimycotic solution, and 50 mM b-mercaptoethanol. Filter sterilized (0.2 mm bottle top filter).

2.1.1. Bone Marrow Extraction

2. Sterile scissors and forceps. 3. Phosphate buffered saline (PBS). 4. Six-well culture plate and 50 ml Falcon tubes. 5. 5 ml Syringe; 25 gauge needle.

2.1.2. Cell Culture

1. Culture medium: DMEM (see Note 2), supplemented with 10% FBS, 20% Wehi-3 conditioned medium (as a source of IL-3; see Note 3), 2 mM L-glutamine, 1% antibioticantimycotic solution, and 50 mM b-mercaptoethanol. Filter sterilized (0.2 mm filter).

2.2. Assessing Maturity of BMCMCs

1. Flow cytometry buffer: PBS supplemented with 2% FBS (see Note 4).

2.2.1. Flow Cytometry

2. 5 ml Polystyrene round bottom tubes (compatible with flow cytometer). 3. Purified anti-mouse CD16/32, clone 93 (eBioscience, San Diego, CA; see Note 5). 4. PE conjugated anti-mouse FceRI alpha, clone MAR-1 (eBioscience; see Note 6). 5. FITC conjugated anti-mouse CD117 (c-Kit) (2B8) (eBioscience; see Note 6). 6. Propidium iodide; prepare a 1 mg/ml stock.

2.2.2. May-Grünwald/ Giemsa Staining

1. Microscope slides, precleaned. 2. PBS. 3. PAP pen. 4. Accustain® May-Grünwald stain (MG1L; Sigma, St. Louis, MO). 5. Accustain® Giemsa stain, Modified (GS1L; Sigma). 6. Mounting medium: CytoSeal 60, DPX, or Permount.

2.3. Adoptive Transfer of BMCMCs into Mast Cell-Deficient Mice 2.3.1. Engraftment of Mast Cell-Deficient Mice

1. DMEM or PBS. 2. 1 ml Syringe; 27 or 30 gauge needle.

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1. 10% Buffered formalin (fixative): 100 ml 40% formalin w/v, 900 ml deionized water, 4 g sodium phosphate monobasic monohydrate (NaH2PO4H2O), 6.5 g sodium phosphate dibasic anhydrous (Na2HPO4). Store at room temperature or 4°C. 2. Carnoy’s solution (fixative): 3:2:1 v/v/v of ethanol, chloroform, and acetic acid. Prepare fresh. 3. Microscope slides, precleaned. 4. Toluidine blue, 1% aqueous. 5. Alcian blue/Safranin O(Csaba) stain: 900 mg Alcian blue 8GX, 45 mg Safranin O, 1.2 g ferric ammonium sulfate, 250 ml acetate buffer (pH 1.42). Acetate buffer: mix 50 ml solution A (13.6 g sodium acetate in 100 ml deionized water) with 62 ml solution B (8.5 ml 12 N HCl plus 91.5 ml deionized water); make up to 100 ml with deionized water and adjust pH to 1.42. Store Csaba stain at room temperature. 6. Ethanol (70, 95, and 100%). 7. HCl (12 and 1 N). 8. Histo-Clear (National Diagnostics, Atlanta, GA). 9. Mounting medium: CytoSeal 60, DPX, or Permount.

3. Methods BMCMCs can easily be generated and expanded in vitro by culturing mouse bone marrow cells in the presence of Interleukin (IL)-3 for a period of 4–6 weeks. Mature mast cells express FceRI (the high-affinity IgE receptor) and Kit (the receptor for stem cell factor, an important regulator of mast cell development, survival, proliferation, and activation) on their surface; these markers can be used to assess the maturity and purity of BMCMC cultures by flow cytometry. Another characteristic feature of mature mast cells is large cytoplasmic granules, which store preformed bioactive mediators of inflammation (such as histamine); BMCMCs can be monitored for metachromatic granule formation by May-Grünwald/Giemsa staining (mast cell granules stain purple). BMCMCs are often used to study the signaling pathways that are activated downstream of specific receptors (by western blot analysis or phosphoflow cytometry), as well as stimulus-induced mast cell functional activation events, including degranulation, and cytokine, chemokine, or eicosanoid lipid mediator release (by various assays including degranulation assays and ELISAs). Although BMCMCs are notoriously difficult to transfect, viruses (e.g., lentiviruses) can readily be used to overexpress or knock-down specific proteins of interest in BMCMCs and the

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resulting cells can be expanded and used for both in vitro and in vivo studies (1, 3). While in vitro studies of mouse BMCMCs can be very useful for investigating mechanisms by which mast cells might exert antiinflammatory or immunosuppressive functions in various immune responses, c-kit-mutant mast cell-deficient mice are often used to assess the in vivo relevance and biological importance of such in vitro observations. The two most commonly used models for such studies are WBB6F1-KitW/W-v or C57BL/6-KitW-sh/W-sh mice (1, 2). C57BL/6-KitW-sh/W-sh mice are gaining in popularity for such studies because they have fewer or less-severe phenotypic abnormalities than are observed in WBB6F1-KitW/W-v mice. For example, C57BL/6-KitW-sh/W-sh mice are neither anemic nor sterile and therefore can easily be crossed with mice carrying other defined genetic abnormalities of interest (11, 12). However, it is important to consider the effects of other c-kit-related phenotypic abnormalities in these mice when interpreting experimental results (see Note 1). To assess the extent to which differences in the expression of biological responses observed in mast cell-deficient and wildtype mice reflect the absence of mast cells or the reduction or absence of specific products expressed by mast cells, BMCMCs can be administered by intravenous (i.v.), intraperitoneal (i.p.), or intradermal (i.d.) injection to generate “mast cell knock-in mice” (1, 4, 11–16). When designing and interpreting experiments, it is important to recognize that the numbers, anatomical distribution, and phenotypic characteristics of adoptively transferred mast cells depend on the injection route or site and the interval of time after injection of the mast cell population into the recipient c-kit-mutant mice. Following adoptive transfer of BMCMCs, tissues can be tested for selective repair of mast cell deficiency by identifying mast cells by the staining of their cytoplasmic granules with Toluidine blue (cytoplasmic granules appear purple, defined as “metachromasia”) or Alcian blue/Safranin O (Csaba) stain; the granules of all mature mast cells stain blue with Alcian blue, but some mast cells have granules that stain red with safranin, perhaps reflecting their content of heparin. One also can measure the amounts of mast cellassociated mediators or transcripts in such tissues (17). 3.1. Generation of BMCMCs from Mouse Bone Marrow 3.1.1. Bone Marrow Extraction

1. Pour sterile PBS into six-well culture plate (one well per mouse) and place on ice. 2. Sacrifice donor animal(s) (see Note 7) and remove skin from legs using sterile forceps and scissors (spray animal(s) and/or rinse instruments with 70% ethanol as needed). 3. Remove femur and tibia; scrape away all tissue from bones using sterile scissors (or scalpel blades), discard fibula. 4. Place bones in PBS on ice until all bones are collected. Carry out all remaining steps in tissue culture hood.

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5. Using sterile instruments, pick up a bone with forceps and cut off epiphyses (bone ends) with scissors to expose medullary cavity. 6. Using a 5 ml syringe with a 25-gauge needle, flush the marrow out of each bone with flushing medium into a labeled 50 ml collection tube. Pool the femoral and tibial bone marrow from each mouse into one collection tube. Fill tube with flushing medium to wash bone marrow cells. 7. Centrifuge marrow (500 × g, 5 min, 20°C). 8. Remove supernatant and resuspend pellet in 10 ml of culture medium per mouse. Place cells in tissue culture flask of appropriate size (e.g., T25 for one mouse, T75 for two mice, etc.). 3.1.2. Cell Culture

1. One to two days after plating cells, transfer medium and suspension cells (leaving debris and adherent cells to stick to bottom of flask) to a new flask and add fresh culture medium (10 ml/mouse). 2. In subsequent weeks, feed cells every 3–4 days (add ~10 ml of culture medium per mouse). Maintain cell density between 2.5 × 105 and 1 × 106 cells/ml. Transfer suspension cells to a new flask once a week until you no longer see adherent cells in the culture flask. 3. Optional (to increase yield of BMCMCs): After 3–4 weeks of culture, centrifuge one-third to half of the culture volume, remove medium, and replace with equal volume of culture medium. Return cells in new medium to original flask, and then dilute cells as necessary. 4. Production of BMCMC populations of suitable phenotype (FceRI+Kit+ cells with prominent cytoplasmic granules) and numbers (we usually generate 2.5–10 × 107 BMCMCs per mouse input) takes 4–6 weeks. Test maturity of cells (see Subheading 3.2) before use in in vitro assays or engraftment into mast cell-deficient mice.

3.2. Assessing Maturity of BMCMCs 3.2.1. Flow Cytometry

1. Need 5 × 104 to 5 × 105 cells/condition. Wash cells with PBS + 2% FBS (see Note 4) in 5 ml polystyrene round bottom tube. Centrifuge at 500 × g, 5 min, 4°C. Aspirate. 2. Dilute anti-mouse CD16/32 (clone 93; see Note 5) monoclonal antibodies 1/200 in PBS + 2% FBS. Add 10 ml to each pellet (to block nonspecific binding of antibodies). Incubate 5 min on ice. Protect samples from direct light for all remaining steps (use lid on ice bucket). 3. Prepare staining solution: dilute PE conjugated anti-mouse FceRI alpha (clone MAR-1) 1/200 and FITC-conjugated anti-mouse CD117 (c-Kit) (2B8) 1/200 in PBS + 2% FBS

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(in the same tube; see Note 6). Add 20 ml of staining solution to desired tubes from step 2; for each BMCMC population, it is best to have one tube of cells that receives staining solution (“stained”) and one tube of cells that does not receive staining solution (“unstained”). Incubate 15–30 min on ice. 4. Wash cells with 1–3 ml PBS + 2% FBS. Centrifuge at 500 × g, 5 min, 4°C. Aspirate. 5. Dilute propidium iodide (PI) stock 1/1,000 in PBS + 2% FBS (final concentration 1 mg/ml). Resuspend cell pellets in 200–400 ml of diluted PI (to exclude dead cells). 6. Analyze cells on flow cytometer. 7. Analyze results using data analysis software (we use FlowJo software; Tree Star, Ashland, OR). An example of the results produced by flow cytometry is shown in Fig. 1. 3.2.2. May-Grünwald/ Giemsa Stain

1. To prepare a cytocentrifuge preparation of BMCMCs (1 × 105 BMCMCs/slide), wash cells once with PBS (centrifuge at 500 × g, 5 min, 20°C, then aspirate) and resuspend cells at 1 × 106 cells/ml in PBS. These instructions assume the use of a Shandon Cytospin Cytocentrifuge: label slide and place in slide holder; place slide filter on top of the slide; place sample chamber on top of filter; close slide holder clip; load 100 ml of BMCMC suspension into sample chamber. Spin at 500 rpm, 5 min. Remove slide and air dry (10 min to overnight). 2. Draw a wax circle around cells using a PAP pen. 3. Add May-Grünwald Stain to cover cells. Incubate 5 min. 4. Wash slides in PBS (2–5 min). 5. Dilute Geimsa 1/20 with deionized water. Add to cells and incubate 20 min. 6. Wash slides in tap water (1 min), then air dry slides. 7. Coverslip cells using mounting medium.

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8. Examine cells using light microscope. An example of BMCMCs after May-Grünwald/Giemsa staining is shown in Fig. 2. 3.3. Adoptive Transfer of BMCMCs into Mast Cell-Deficient Mice 3.3.1. Engraftment of Mast Cell-Deficient Mice

1. Culture a sufficient number of donor BMCMCs for the mice you wish to engraft (see Note 8). The number of BMCMCs that should be used for adoptive transfer into mast cell-deficient mice depends on the route of transfer (Table 1); the following numbers are guidelines, but can be refined, as indicated based on the purpose of the study, for each type of model under investigation. 2. Count cells and resuspend in appropriate volume cold DMEM or PBS. Keep cells on ice until ready to inject.

Fig. 2. C57BL/6J BMCMCs stained with May-Grünwald/Giemsa. Note the large number of cytoplasmic granules throughout the cytoplasm. N nucleus.

Table 1 Numbers of BMCMCs recommended for adoptive transfer into mast cell-deficient mice Number of BMCMCs

Injection volume

Needle gauge

Ear

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30 gauge

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400 ml (8 × 50 ml injections in two rows down the length of shaved back)

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100 ml (retroorbital) or 200 ml (tail vein)

30 gauge

Transfer route Intradermal (i.d.)

6

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3. Inject BMCMCs [use 27- or 30-gauge needle (see Table 1) and 1 ml syringe] into anesthetized c-kit-mutant mice; recipient mice should be 4–6 weeks of age. 4. Time needed for engraftment depends on the route of injection and should be assessed for each experiment (see Note 10). General guidelines are as follows: i.d., 6–8 weeks; i.p., 4–8 weeks; i.v., 8–18 weeks. Wait appropriate amount of time and perform desired in vivo experiment. Part of the analysis should include assessment of the numbers of mast cells that have been engrafted into the anatomical sites of interest (see Subheading 3.3.2 and Note 11). 3.3.2. Staining and Quantification of Mast Cells

1. Euthanize mice and remove tissue(s) of interest (ear pinna, back skin, lung, spleen, lymph nodes, etc.). Place each specimen in labeled histology cassette. 2. Fix specimens in 10% buffered formalin (overnight at 4°C) (see Note 12). 3. Process samples for paraffin embedding (we use an automated processing schedule on the Shandon Hypercentre XP enclosed tissue processor). 4. Embed tissues in paraffin (we use the Shandon Histocentre 2), ensuring a cross-sectional orientation of all tissues, and section tissues at 4 mm. 5. Dewax/rehydrate paraffin sections: 2× Histo-Clear (5 min each), 2× 100% ethanol (2 min each), 1× 95% ethanol (2 min), 1× 70% ethanol (2 min), 3× deionized water (2 min each). 6. Stain tissues with 0.1% toluidine blue or Alcian blue/Safranin O (Csaba) stain (see Note 12). Toluidine blue. Dilute Toluidine blue 1% aqueous stock 1/10 in 1 N HCl. Add 0.1% Toluidine blue to cover tissue sections. Incubate 1 min. Rinse immediately with tap water. Air dry, then cover slip using DPX or other suitable mounting medium.

3.3.3. Alcian blue/Safranin O (Csaba) stain

Add Alcian blue/Safranin O (Csaba) stain to cover sections. Incubate 20–30 min. Dehydrate/mount in 70% ethanol (30 s), 95% ethanol (1 min), 2× 100% ethanol (2 min each), 2× HistoClear (2 min each). Mount using Permount or other suitable mounting medium. 7. Count mast cells using light microscope. For ear pinna and back skin, average the number of mast cells counted in ten consecutive fields of 1 mm length; express as number of mast cells per horizontal field length (mm). For all other tissues, quantify mast cells according to area (mm2). Report the average number of mast cells present in a 10–25 field count and express as number of mast cells per mm2.

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1. Carefully arrange mesenteric windows onto slides. Air dry (~4 h). 2. Fix specimen in Carnoy’s solution (1 h). Air dry (overnight). After fixation, can store mesenteric windows at 4°C until ready to stain. 3. Carefully remove bowel from mesenteric window. Add Alcian blue/Safranin O (Csaba) stain to cover mesentery. Incubate 20–30 min. Wash slide in tap water. Air dry. Dip in HistoClear (optional). Mount using Cytoseal 60 or suitable mounting medium. 4. Count cells as indicated in step 7 above (report mast cells numbers per mm2 of mesentery).

4. Notes 1. In addition to lacking mast cells, c-kit-mutant mice have other phenotypic abnormalities that may reflect effects of the c-kit mutation on other cell lineages. For example, WBB6F1-KitW/W-v mice, but not C57BL/6-KitW-sh/W-sh mice, are anemic and sterile. Moreover, C57BL/6-KitW-sh/W-sh mice exhibit increased numbers of neutrophils in the spleen, bone marrow, and blood, whereas KitW/W-v mice are neutropenic (16, 18, 19). The c-kit-dependent but mast cell-independent phenotypic abnormalities in these mice should be taken into account when using this approach to assess the role of mast cells in various disease models. 2. Can also culture BMCMCs in Iscove’s Modified Dulbecco’s Medium (IMDM). 3. Can culture BMCMCs in 10 ng/ml interleukin (IL)-3 (instead of 20% WEHI-3 conditioned medium). To generate WEHI-3 conditioned medium: Culture WEHI-3 cells (ATCC: TIB 68) at 1 × 105 cells/ml in IMDM containing 10% FBS for 3–4 days until the cell concentration reaches 6 × 105–1× 106 cells/ml. Centrifuge culture (500 × g, 10 min), then centrifuge supernatant again (1,500 × g, 15 min). Filter supernatant through a 0.45 mm filter (Nalgene) and store 100% WEHI-3 conditioned medium at –20°C. 4. Can use PBS, PBS + 0.5% FBS, or PBS + 2% FBS as buffer for flow cytometry staining. 5. Anti-mouse CD16/32 clone 93 or clone 2.4G2 antibodies can be used to block Fc binding. 6. Anti-mouse FceRI alpha or CD117 (c-Kit; 2B8) antibodies can be coupled to other convenient fluorophores (APC, etc.). 7. We typically generate BMCMCs from mice that are 4–12 weeks of age.

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8. Male-derived donor BMCMCs are not suitable for engraftment of female mice. Female-derived donor BMCMCs will successfully engraft into both male and female recipients. Ideally, donor BMCMCs should be generated from mice of the same genetic background as the recipient c-kit-mutant mice. 9. Ear (i.d.) engraftment: first injection in the middle of the ear; second injection towards tip of the ear (the fluid remaining from the first injection makes the second injection much easier). 10. After the adoptive transfer of BMCMCs into c-kit-mutant mice, the numbers, anatomical distribution, and features of maturation of the adoptively transferred mast cells vary according to the route of injection, the anatomic site, and the interval after transfer (1, 11–13, 15, 20, 21). For example, 4 weeks after the i.p. transfer of BMCMCs into c-kit-mutant mice, mast cells numbers are comparable to those observed in wildtype mice (as assessed by the evaluation of periotoneal lavage fluids) and mast cells can be found in the mesentery (14, 15, 20, 22, 23). During the weeks after adoptive transfer, the transferred mast cells acquire phenotypic characteristics that, compared to those of the in vitro-derived mast cells assessed prior to in vivo injection, more closely resemble those of native peritoneal mast cells in the corresponding anatomical sites in wildtype mice (1, 13, 20). However, it is impossible to prove that such adoptively transferred mast cells are literally identical to the native mast cell populations at the corresponding anatomical sites in wildtype mice. Moreover, based on studies of adoptively transferred mast cells in the peritoneal cavity, it appears that the longer the adoptively transferred mast cells remain in a particular anatomical site, the more closely their phenotype resembles that of the corresponding native population of mast cells in wildtype mice of the same age (13). Thus, depending on which aspects of mast cell function are being assessed in each experimental model under investigation, it may be necessary to wait for longer intervals after the transfer of mast cells to initiate the experiment (as the adoptively transferred mast cells may require higher levels of maturity to perform some functions than is required for other functions). 11. For i.p. transfer, we quantify the number of mast cells present in cells washed from the peritoneal cavity and also evaluate the presence and distribution of mast cells in the mesentery of the recipient mice (11, 22, 23). For i.d. transfer, we quantify the number of mast cells per mm2 of dermis (11). For i.v. transfer to engraft mast cells in the lungs, we quantify the number of mast cells per mm2 of lung parenchyma (11, 24– 27). We find that the distribution of mast cells in the peritoneal cavity (after i.p. transfer of BMCMCs) or dermis (after i.d. transfer of BMCMCs) is very similar to that of the native

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populations of mast cells in the same anatomical sites in the corresponding wildtype mice. However, after i.v. transfer of BMCMCs, the distribution of mast cells in the respiratory tract of the recipient c-kit-mutant mice differs from that in the corresponding wildtype mice in that there are fewer mast cells in the trachea and more in the periphery of the lung in the mast cell knock-in mice than in the corresponding wildtype mice (24). Remarkably, for many features of the “asthma models” analyzed in such mast cell knock-in mice, the results obtained in the mast cell knock-in mice were very similar to (and in many cases statistically indistinguishable from) those in the wildtype mice (24–27). This may reflect the fact that many mast cell functions are mediated by the release of soluble mediators, which can have their effects at some distance from the mast cells that secreted them. 12. Toluidine blue or (Alcian blue/Safranin O (Csaba) stain can be used to quantify mast cells containing cytoplasmic granules in both paraffin and frozen sections. However, if mast cells have recently undergone extensive degranulation, identifying them using stains which bind to the cytoplasmic granules (or by using immunohistochemistry to detect constituents stored in the cytoplasmic granules) may underestimate the numbers of mast cells actually present in the tissues; this is also true when Giemsa staining is used to quantify mast cells in Eponembedded 1 mm sections [e.g., in the skin after treatment with phorbol 12-myristate 13-acetate (PMA)] (28).

Acknowledgments We thank Jennifer Lilla and Chen Liu for advice regarding histology, Eon Rios and Mariola Liebersbach for sharing their expertise in flow cytometry and cell culture, respectively, and Adrian Piliponsky for technical advice. References 1. Metz M, Grimbaldeston MA, Nakae S, et al. (2007) Mast cells in the promotion and limitation of chronic inflammation. Immunol Rev 217, 304–28. 2. Galli SJ, Grimbaldeston M, Tsai M. (2008) Immunomodulatory mast cells: negative, as well as positive, regulators of immunity. Nat Rev Immunol 8, 478–86. 3. Kalesnikoff J, Galli SJ. (2008) New developments in mast cell biology. Nat Immunol 9, 1215–23.

4. Galli SJ, Kalesnikoff J, Grimbaldeston MA, et al. (2005) Mast cells as “tunable” effector and immunoregulatory cells: recent advances. Annu Rev Immunol 23, 749–86. 5. Scholten J, Hartmann K, Gerbaulet A, et al. (2008) Mast cell-specific Cre/loxP-mediated recombination in vivo. Transgenic Res 17, 307–15. 6. Musch W, Wege AK, Mannel DN, et al. (2008) Generation and characterization of alphachymase-Cre transgenic mice. Genesis 46, 163–6.

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7. Hart PH, Grimbaldeston MA, Swift GJ, et al. (1998) Dermal mast cells determine susceptibility to ultraviolet B-induced systemic suppression of contact hypersensitivity responses in mice. J Exp Med 187, 2045–53. 8. Depinay N, Hacini F, Beghdadi W, et al. (2006) Mast cell-dependent down-regulation of antigen-specific immune responses by mosquito bites. J Immunol 176, 4141–6. 9. Lu LF, Lind EF, Gondek DC, et al. (2006) Mast cells are essential intermediaries in regulatory T-cell tolerance. Nature 442, 997–1002. 10. Grimbaldeston MA, Nakae S, Kalesnikoff J, et al. (2007) Mast cell-derived interleukin 10 limits skin pathology in contact dermatitis and chronic irradiation with ultraviolet B. Nat Immunol 8, 1095–104. 11. Grimbaldeston MA, Chen CC, Piliponsky AM, et al. (2005) Mast cell-deficient W-sash c-kit mutant KitW-sh/W-sh mice as a model for investigating mast cell biology in vivo. Am J Pathol 167, 835–48. 12. Wolters PJ, Mallen-St Clair J, Lewis CC, et al. (2005) Tissue-selective mast cell reconstitution and differential lung gene expression in mast cell-deficient Kit(W-sh)/Kit(W-sh) sash mice. Clin Exp Allergy 35, 82–8. 13. Nakano T, Sonoda T, Hayashi C, et al. (1985) Fate of bone marrow-derived cultured mast cells after intracutaneous, intraperitoneal, and intravenous transfer into genetically mast celldeficient W/W v mice. Evidence that cultured mast cells can give rise to both connective tissue type and mucosal mast cells. J Exp Med 162, 1025–43. 14. Echtenacher B, Mannel DN, Hultner L. (1996) Critical protective role of mast cells in a model of acute septic peritonitis. Nature 381, 75–7. 15. Jippo T, Morii E, Ito A, et al. (2003) Effect of anatomical distribution of mast cells on their defense function against bacterial infections: demonstration using partially mast cell-deficient tg/tg mice. J Exp Med 197, 1417–25. 16. Piliponsky AM, Chen CC, Grimbaldeston MA, et al. (2010) Mast cell-derived TNF can exacerbate mortality during severe bacterial infections in C57BL/6-KitW-sh/W-sh mice Am J Pathol 176, 926–38. 17. Tsai M, Miyamoto M, Tam SY, et al. (1995) Detection of mouse mast cell-associated protease mRNA. Heparinase treatment greatly

18.

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improves RT-PCR of tissues containing mast cell heparin. Am J Pathol 146, 335–43. Zhou JS, Xing W, Friend DS, et al. (2007) Mast cell deficiency in KitW-sh mice does not impair antibody-mediated arthritis. J Exp Med 204, 2797–802. Nigrovic PA, Gray DH, Jones T, et al. (2008) Genetic inversion in mast cell-deficient Wsh mice interrupts corin and manifests as hematopoietic and cardiac aberrancy. Am J Pathol 173, 1693–701. Otsu K, Nakano T, Kanakura Y, et al. (1987) Phenotypic changes of bone marrow-derived mast cells after intraperitoneal transfer into W/Wv mice that are genetically deficient in mast cells. J Exp Med 165, 615–27. Tanzola MB, Robbie-Ryan M, Gutekunst CA, et al. (2003) Mast cells exert effects outside the central nervous system to influence experimental allergic encephalomyelitis disease course. J Immunol 171, 4385–91. Metz M, Piliponsky AM, Chen CC, et al. (2006) Mast cells can enhance resistance to snake and honeybee venoms. Science 313, 526–30. Piliponsky AM, Chen CC, Nishimura T, et al. (2008) Neurotensin increases mortality and mast cells reduce neurotensin levels in a mouse model of sepsis. Nat Med 14, 392–8. Martin TR, Takeishi T, Katz HR, et al. (1993) Mast cell activation enhances airway responsiveness to methacholine in the mouse. J Clin Invest 91, 1176–82. Williams CM, Galli SJ. (2000) Mast cells can amplify airway reactivity and features of chronic inflammation in an asthma model in mice. J Exp Med 192, 455–62. Yu M, Tsai M, Tam SY, et al. (2006) Mast cells can promote the development of multiple features of chronic asthma in mice. J Clin Invest 116, 1633–41. Nakae S, Ho LH, Yu M, et al. (2007) Mast cell-derived TNF contributes to airway hyperreactivity, inflammation, and TH2 cytokine production in an asthma model in mice. J Allergy Clin Immunol 120, 48–55. Wershil BK, Murakami T, Galli SJ. (1988) Mast cell-dependent amplification of an immunologically nonspecific inflammatory response. Mast cells are required for the full expression of cutaneous acute inflammation induced by phorbol 12-myristate 13-acetate. J Immunol 140, 2356–60.

Chapter 16 A Mesenchymal Stem Cell Potency Assay Joy Jiao, Jack M. Milwid, Martin L. Yarmush, and Biju Parekkadan Abstract Mesenchymal stem cells (MSCs) are capable of modulating the immune system and have been used to successfully treat a variety of inflammatory diseases in preclinical studies. Recent evidence has implicated paracrine signaling as the predominant mechanism of MSC therapeutic activity. We have shown in models of inflammatory organ failure that the factors secreted by MSCs are capable of enhancing survival, downregulating inflammation, and promoting endogenous repair programs that lead to the reversal of these diseases. As a marker of disease resolution, we have observed an increase in serum IL-10 when MSC-conditioned medium (MSC-CM) or lysate (MSC-Ly) is administered in vivo. Here we present an in vitro model of IL-10 release from blood cells that recapitulates this in vivo phenomenon. This assay provides a powerful tool in analyzing the potency of MSC-CM and MSC-Ly, as well as characterizing the interaction between MSC-CM and target cells in the blood. Key words: Mesenchymal stem cell, IL-10, Potency assay, Organ injury, Inflammation, Autoimmunity, Transplantation

1. Introduction Bone marrow mesenchymal stem cells (MSCs) are resident nonhematopoietic progenitor cells that possess potent immunomodulatory abilities (1–3). Allogeneic MSC transplants have been used to successfully treat hematological (1, 4), cardiovascular (5, 6), as well as neurological (7, 8) and inherited diseases (9, 10) in preclinical studies, and allogeneic transplantation of MSCs has been given expanded access as a treatment for padiatric GvHD. Recent studies involving MSC transplantation have revealed that the therapeutic activity of these grafts is independent of differentiation, and paracrine interactions of MSCs with tissue and immune cells provide the majority of therapeutic benefit (11–15).

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Fig. 1. Schematic of in vitro and in vivo potency tests of MSC-derived materials.

We have found administration of concentrated MSCconditioned medium (MSC-CM) or MSC lysate (MSC-Ly) can reproduce the effects of an MSC graft in vivo and significantly increase serum IL-10 levels in two animal models (13, 14). IL-10 is a well-known antiinflammatory cytokine that can inhibit the secretion of proinflammatory cytokines and protect cells from apoptosis and necrosis in the context of acute inflammation (16–18). We have determined that MSC-CM and lysate also substantially enhance IL-10 secretion by peripheral blood mononuclear cells (PBMCs) in vitro and have developed an assay based on this discovery. This assay allows for rapid and reproducible assessment of the potency of MSCs and MSC molecular products in a manner relevant to animal and human testing of cell therapy. Conditioned medium and cell lysate are derived from human MSC cultures and can subsequently be used for both in vitro and in vivo experimentation (see Fig. 1). Below we provide methods for an ELISA-based IL-10 assay for in vitro testing, a powerful tool to aid in determining the efficacy and underlying paracrine mechanisms of MSC transplants and MSC-based therapies.

2. Materials 2.1. Isolation of MSCs from Whole Bone Marrow and Maintenance of Culture

1. Whole bone marrow aspirate: can be obtained commercially (Lonza) or from a consented human donor. 2. Phosphate-buffered saline (PBS) (see Note 1). 3. Ficoll-Paque density 1.077 g/mL (GE Healthcare). 4. Table-top centrifuge capable of 1,500 × g. 5. Hemocytometer and microscope, or other method capable of determining cell count.

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6. Human MSC medium: a-minimum essential medium Eagle (Sigma Aldrich) supplemented with 15% (v/v) fetal bovine serum (HyClone), 2% (v/v) penicillin streptomycin solution (GIBCO), 1 mg/mL gentamicin (Sigma Aldrich), 1 ng/mL bFGF (R&D Systems). Store at 4°C. 7. 37°C incubator. 2.2. Collection, Concentration, and Storage of MSC-CM

1. Conditioning medium: Dulbecco’s modified essential medium (Sigma Aldrich) supplemented with 0.5% (w/v) bovine serum albumin and 2% (v/v) penicillin streptomycin solution (GIBCO). 2. 1× Trypsin prepared from 10 × 0.5% Trypsin-EDTA (GIBCO). 3. Hemocytometer. 4. For small volume of MSC-CM: 3 kDa centrifugal filters (Amicon Ultra Ultracel-3K, Cat No. UFC800396). Centrifuge capable of spinning at 4,000 × g. 5. For large volume of MSC-CM: Amicon pressure concentrator (no longer manufactured), pressurized nitrogen gas and 3 kDa regenerated cellulose ultrafiltration membranes (Millipore Ultracel). 6. +4°C freezer for storage.

2.3. Preparation of MSC Lysate

1. Sonicator. 2. Benchtop centrifuge capable of 1,500 × g. 3. +4°C freezer for storage.

2.4. Isolation of PBMCs from Whole Blood

1. Fresh whole blood from consenting donor. 2. PBS. 3. Ficoll-Paque density 1.077 g/mL (GE Healthcare). 4. C-10 medium: 500 mL 1× RPMI medium (GIBCO 1650), 50 mL inactivated fetal bovine serum, 6 mL MEM nonessential amino acids solution 10 mM 100× (GIBCO Cat No. 1140-050), 6 mL 100× sodium pyruvate (GIBCO 11360), 6 mL glutamate, 6 mL penicillin streptomycin solution (GIBCO 15140), 6 mL sodium bicarbonate, 3 mL betamercaptoethanol. Sterilize with corning bottle top filter or other means of sterilization. Store at 4°C. 5. Centrifuge capable of spinning at 1,500 × g. 6. 96-Well plate. 7. 37°C incubator.

2.5. Preparation of Samples and Stimulation with LPS

1. 96-Well plate seeded with 50 mL per well at 2 × 10 6 PBMCs/mL. 2. MSC-CM and lysate, 50 mL per well.

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3. LPS (Escherichia coli 0111:B4, Sigma-Aldrich L4391) at 30 mg/mL, 50 mL per well. Stock solution is diluted in C10 medium. 4. 37°C incubator. 5. −80°C freezer. 2.6. ELISA and Analysis of Results

1. BD OptEIA™ human IL-10 ELISA Set and recommended buffers and solutions, or other human IL-10 ELISA Kit. 2. BD Falcon™ Microtest™ 96-well ELISA plate or other highbinding ELISA plate. 3. Plate reader (spectrophotometer) capable of reading at indicated wavelengths in ELISA Kit (450 nm with correction at 570 nm for BD OptEIA™ Human IL-10 ELISA set). 4. Microsoft Excel or other program capable of processing data from spectrophotometer.

3. Methods Overview: MSC-CM or lysate is prepared and added to PBMCs freshly isolated from whole blood and plated in a 96-well plate. As a mock cell control, we perform the assay with CM and/or lysates from fibroblasts. The plate is incubated overnight at 37°C for 16–18 h. The PBMCs are then stimulated with LPS for 5 h, at which time the plate is centrifuged and the supernatant stored for ELISA (see Fig. 2). 3.1. Isolation of MSCs from Whole Bone Marrow

Whole bone marrow centrifuged with Ficoll results in a pattern comparable to that of centrifuged whole blood; the corresponding “buffy coat” is enriched with MSCs. This layer is collected, counted, and plated on tissue culture polystyrene. The purity of MSCs isolated from whole bone marrow is relatively high due to their ability to differentially adhere to cell culture substrates compared to other hematopoietic marrow cells. Subsequent medium changes eliminate hematopoietic cells and other nonadherent cells. The identity of MSCs can be confirmed through phenotype

Fig. 2. Summary of in vitro inflammation assay for the testing of MSC-derived factors.

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and multipotency analysis using flow cytometry or differentiation media, respectively (see Note 3). 1. Wash bone marrow with equivalent volume of PBS, thus diluting the bone marrow 1:2. Prepare 5 mL Ficoll for each 10 mL of diluted bone marrow. Add diluted whole bone marrow slowly; avoid disturbance of the boundary between Ficoll and marrow (see Note 1). 2. Spin 30 min at 1,500 × g with no brake. Collect the resulting mononuclear cell layer and wash with 5 mL PBS. 3. Spin 10 min at 1,500 × g, high brake. Resuspend pellet in 1 mL hMSC medium. Determine cell count and seed at a density of 1–10 × 103 cells/cm2. 4. Let cells adhere and grow for 10 days. Perform first medium change at day 10; the following medium change is done at day 17. Subsequent medium changes should be performed at a frequency of every 3 or 4 days. Cells are typically used between passages 1–6. 3.2. Collection, Concentration, and Storage of MSC-CM

By convention, 1× MSC-CM is defined as conditioned medium from two million cells concentrated to 1 mL of volume. MSCs are incubated for 24 h in conditioning medium and concentrated to 1×. For small volumes, centrifugal filter tubes can be used. For larger volumes, the Amicon pressure concentrator and pressurized nitrogen gas offers a more efficient method. 1. Perform medium change on MSC cultures with two PBS washes. Add conditioning medium, incubate for 24 h (see Note 4). 2. Collect MSC-CM. Trypsinize cells; determine cell count and final volume of 1× MSC-CM. For small volumes: 3. Add 4 mL PBS to centrifugal filter tube. Let centrifuge spin to 4,000 × g for a few minutes for the PBS to run through the filter to remove residual glycerin in the ultrafiltration membrane. Discard the PBS in both compartments of the centrifugal filters. 4. Add 4 mL MSC-CM under sterile conditions and spin at 4,000 × g for 5–15 min. Discard flow through and fill the centrifugal filter tube with new MSC-CM. Pipette the conditioned medium up and down to wash the filter and lessen congestion of the filter with proteins. Repeat process until the desired volume is reached. For large volumes: 3. Sterilize the ultrafiltration membrane with 70% ethanol and let dry. Assemble pressure concentrator according to the manufacturer instructions. Place pressure concentrator onto stir plate and prepare a waste bottle.

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4. Run 20 mL of PBS through the concentrator. 5. Discard waste and add MSC-CM. Monitor waste level in the bottle to determine the volume of MSC-CM. 6. Let run until desired final volume is reached. Disconnect and depressurize concentrator. 3.3. Preparation of MSC Lysate

While cell lysate can be obtained through lysate buffers and other chemical means, sonication, which causes physical disruption of the cell membrane, provides pure lysate without chemical contaminant and possible confounding factors. 1. Trypsize MSCs and pellet in tabletop centrifuge at 1,000 × g. 2. Discard supernatant and gently layer 1 mL PBS per 2 × 106 cells on top of cell pellet. To visualize lysis, do not resuspend cells into solution. 3. Sonicate cell pellet at 3 AU for 5 s with three pulses and collect in ice. 4. Centrifuge lysate at 2,000 × g for 2 min to precipitate membrane fragments. Retain the solution phase. The solution phase from this process is considered MSC-Ly.

3.4. Isolation of PBMCs from Whole Blood

1. Calculate the number of wells needed to conduct the assay to estimate the amount of blood needed, leaving extra wells for standards and controls. A total of 100,000 PBMCs will be required per well. Collect fresh whole blood from a consented donor. It is best to plate the PBMCs and add the MSC-CM or lysate on the day of the separation to ensure maximum viability (see Note 2). We have found 5 mL of blood to safely supply 10–15 million PBMCs, but this count varies from donor to donor. 2. Wash blood with equivalent volume of PBS, thus diluting the blood 1:2. Prepare 5 mL Ficoll for each 10 mL of diluted blood. Gently layer diluted blood on the Ficoll column (see Note 1). 3. Spin 30 min at 1,500 × g with no brake. Collect buffy coat and wash with 5 mL PBS. 4. Spin 10 min at 1,500 × g. Resuspend pellet in 1 mL C-10 medium and determine cell count. 5. Dilute or concentrate to two million cells/mL. Seed in 96-well plate at 50 mL per well (100,000 cells per well). Incubate at 37°C for storage if MSC-CM or lysate cannot be added immediately.

3.5. Preparation of Samples and Stimulation with LPS

1. Add 50 mL of MSC-CM or lysate to each well in the blood plate and incubate at 37°C for 16–18 h. 2. Prepare LPS solution. Vortex well before dilution. Prepare 6 mL per plate of LPS diluted in C-10. Add 50 mL to each

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well, thus yielding a final concentration of 10 mg/mL in the wells, and incubate at 37°C or 5 h. 3. Spin plates at 1,500 × g for 10 min to settle PBMCs to the bottom of the wells. Collect supernatant in a new 96-well plate and store at −80°C (see Note 5). 3.6. ELISA and Analysis of Results

1. Perform ELISA for human IL-10 according to the manufacturer instructions. Use spectrophotometer to read plates at wavelengths indicated by ELISA Kit and import raw data into program of choice. 2. Perform linear regression to generate standard curve and equation. 3. Convert raw absorbance data into concentration values using linear regression equation and generate bar graph. High IL-10 secretion by PBMCs indicates high potency of treatment.

3.7. Conclusion

MSCs are promising candidates for cell-based immunomodulatory therapy. They can be easily isolated from bone marrow aspirates, expanded 50 population doublings in 10 weeks with minimal loss in potency, and to date have not been found to cause adverse immune responses in allogenic transplantation recipients (19). Despite controversial theories regarding the primary therapeutic mechanism of action, the uses of MSC treatments have become diverse (9). Currently ongoing clinical trials exist for the use of MSC transplants in steroid refractory graft vs. host disease (20), periodontitis (21), and severe chronic myocardial ischemia (9, 22) among others. In our laboratory, we have demonstrated the effective use of MSC-CM and MSC-Ly in treating multiple organ dysfunction syndrome. To harness both the secreted and intracellular metabolism of MSCs, we have also created an MSC extracorporeal device for the treatment of organ failure (13, 14). By quantifying the potency of MSC-CM or MSC-Ly, this method enables optimization of dosage, growth, and storage conditions, as well as treatment procedures for clinical use of MSCs and MSC-based products. In future development of these products, the antiinflammatory activity that these cells possess can be measured reliably and reproducibly with this assay, providing for better consistency and more rigorous release criteria. The assay also provides a valuable tool in elucidating the mechanism underlying MSC immunomodulation. The speed at which MSC transplantation conveys therapeutic activity (~hours) is considerably faster than that of other cell transplants for regeneration purposes (~days to weeks), an inconsistency that can only be explained if other hypotheses for the mechanism of action other than engraftment and differentiation are considered. The time scale of these results could plausibly be explained by the occurrence of MSC lysis and the release of what would be paracrine factors

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during transplantation. This potency assay can account for and facilitate decomposing the “lysate effect.”

4. Notes 1. It is recommended that the bone marrow or blood be diluted 1:1 with PBS, as the larger volume provides a greater margin of error during the collection of the buffy coat. 2. We found that blood stored for prolonged periods before use tends to be activated during storage and can thereby confound the results of the assay. Freshly obtained PBMCs are preferred. 3. Phenotype and multipotency analysis. While the identities of relevant cell surface markers of human MSCs remain controversial, Table 1 offers a certain immunophenotype of MSCs. It is possible to label for certain cell surface markers and conduct flow cytometry for isolation. MSCs have been shown to be capable of differentiating into bone, cartilage, adipose cells, and myoblasts in vivo (9). Lee et al. developed differentiation media in which, under the right conditions, MSCs can be observed to differentiate into these tissues in vitro (23). Table 2 includes specialized media used to encourage differentiation of MSCs. MSC differentiation kits are also commercially available from vendors. It is also possible to perform multipotency tests in vivo through subcutaneous transplantation of MSCs. While it is beyond the scope of this paper to discuss in vivo assays, the

Table 1 Immunophenotype of human MSCs (19) Positive

Negative

Inducible

CD13, CD29, CD44, CD49a, b, c, e, f, CD51, CD54, CD58, CD71, CD73, CD90, CD102, CD105, CD106, CDw119, CD120a, CD120b, CD123, CD124, CD126, CD127, CD140a, CD166, P75, TGF-bIR, TGF-bIIR, HLA-A, B, C, SSEA-3, SSEA-4, D7, PD-L1

CD3, CD4, CD6, CD9, CD10, CD11a, CD14, CD15, CD18, CD21, CD25, CD31, CD34, CD36, CD38, CD45, CD49d, CD50, CD62E, L, S, CD80, CD86, CD95, CD117, CD133, SSEA-1, ABO

HLA-DR

CD cluster of differentiation; TGF transforming growth factor; HLA human leukocyte antigen; SSEA stage-specific embryonic antigen; ABO blood group antigens

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Table 2 Differentiation media for MSCs (23) Differentiation medium Composition Osteogenic

IMDM with 0.1 mM dexamethasone (Sigma-Aldrich, St Louis, MO), 0.2 mM ascorbic acid (AsA; Sigma-Aldrich), 10 mM-glycerol phosphate (Sigma-Aldrich)

Chondrogenic

High-glucose DMEM (Bio-fluid, Rockville, MD) with 0.1 M dexamethasone, 50 g/mL AsA, 100 g/mL sodium pyruvate(Sigma-Aldrich), 40 g/mL proline (Sigma-Aldrich), 10 ng/mL TGF-1, and 50 mg/mL ITS premix (Becton Dickinson; 6.25 g/mL insulin, 6.25 g/mL transferrin, 6.25 ng/mL selenius acid, 1.25 mg/mL bovine serum albumin (BSA), and 5.35 mg/mL linoleic acid)

Adipogenic

IMDM with 0.5 mM 3-isobutyl-1-methylxanthine (Sigma-Aldrich), 1 M hydrocortisone (Sigma-Aldrich), 0.1 mM indomethacin(Sigma-Aldrich), and 10% rabbit serum (Sigma-Aldrich)

reader may find it worthwhile to refer to the work of Bianco et al. concerning the formation of ectopic bone marrow using subcutaneous transplants of MSCs (24). 4. This volume is typically 15 mL for Corning T-175 Flasks. For other containers, a similar volume per cell ratio should be achieved, although small variations are negligible as the final definition of 1× and 10× medium is based on cell count and not initial volume. 5. Despite centrifugation, the supernatant may still be contaminated with PBMCs during the collection process. Freezing before the ELISA is recommended to lyse all remaining cells.

Acknowledgments This work was partially supported by grants from the National Institutes of Health (R01 DK43371), MIT Class of 1972 Fund, and the Shriners Hospitals for Children. References 1. Le Blanc, K., I. Rasmusson, B. Sundberg, C. Gotherstrom, et al. Treatment of severe acute graft-versus-host disease with third party haploidentical mesenchymal stem cells. The Lancet 363.9419 (2004): 1439–441. 2. Aggarwal, S., M. F. Pittenger. Human mesenchymal stem cells modulate allogeneic immune cell responses. Blood 10.4 (2005): 1815–822.

3. Bartholomew, A., C. Sturgeon, M. Siatskas, K. Ferrer, K. McIntosh, S. Patil, W. Hardy, S. Devine, D. Ucker, R. Deans, A. Moseley, R. Hoffman. Mesenchymal stem cells suppress lymphocyte proliferation in vitro and prolong skin graft survival in vivo. Experimental Hematology 30.1 (2002): 42–48. 4. Ringden, O., M. Uzunel, I. Rasmusson, M. Remberger, B. Sundberg, H. Lonnies, HU

230

5.

6.

7.

8.

9.

10.

11.

12.

Jiao et al. Marschall, A. Dlugosz, A. Szakos, Z. Hassan, B. Omazic, J. Aschan, L. Barkholt, K. Le Blanc. Mesenchymal stem cells for treatment of therapy-resistant graft-versus-host disease. Transplantation 81.10 (2006): 1390–397. Perin, E. C., H. F. Dohmann, R. Borojevic, S. A. Silva, A. L. Sousa, C. T. Mesquita, M. I. Rossi, A. C. Carvalho, H. S. Dutra, H. J. Dohmann, G. V. Silva, L. Belem, R. Vivacqua, F. O. Rangel, R. Esporcatte, Y. J. Geng, W. K. Vaughn, J. A. Assad, E. T. Mesquita, J. T. Willerson. Transendocardial, autologous bone marrow cell transplantation for severe, chronic ischemic heart failure. Circulation 107.18 (2003): 9040–42. Miyahara, Y., N. Nagaya, M. Kataoka, B. Yanagawa, K. Tanaka, H. Hao, K. Ishino, H. Ishida, T.Shimizu, K. Kangawa, S. Sano, T. Okano, S. Kitamura, H. Mori. Monolayered mesenchymal stem cells repair scarred myocardium after myocardial infarction. Nature Medicine 12 (2006): 459–65. Jin, H. K., J. E. Carter, G. W. Hungtley, E. H. Schuchman. Intracerebral transplantation of mesenchymal stem cells into acid sphingomyelinase-deficient mice delays the onset of neurological abnormalities and extends their life span. Journal of Clinical Investigation 109.9 (2002): 1183–191. Zhao, L. R., W. M Duan, M. Reyes, C. D. Keene, C. M. Verfaillie, W. C. Low. Human bone marrow stem cells exhibit neural phenotypes and ameliorate neurological deficits after grafting into the ischemic brain of rats. Experimental Neurology 174.1 (2002): 11–20. Giordano, A., U. Galderisi, I. R. Marino. From the laboratory bench to the patient’s bedside: An update on clinical trials with mesenchymal stem cells. Journal of Cellular Physiology 211.1 (2007): 27–35. Horwitz, E. M., D. J. Prockop, L. A. Fitzpatrick, W. W. Koo, P. L. Gordon, M. Neel, M. Sussman, P. Orchard, J. C. Marx, R. E. Pyeritz, M. K. Brenner. Transplantability and therapeutic effects of bone marrowderived mesenchymal cells in children with osteogenesis imperfecta. Nature Medicine 5 (1999): 309–13. Gnecchi, M., H. He, N. Noiseux, O. D. Liang, L. Zhang, F. Morello, H. Mu, L. G. Melo, R. E. Pratt, J. S. Ingwall, V. J. Dzau. Evidence supporting paracrine hypothesis for Akt-modified mesenchymal stem cell-mediated cardiac protection and functional improvement. The FASEB Journal 20 (2006): 661–69. Mirotsou, M., Z. Zhang, A. Deb, L. Zhang, M. Gnecchi, N. Noiseux, H. Mu, A.Pachori, V. Dzau. Secreted frizzled related protein 2 (Sfrp2) is the key Akt-mesenchymal stem cell-

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

released paracrine factor mediating myocardial survival and repair. PNAS 104.5 (2006): 1643–648. Kinnarid, T., E. Stabile, M. S. Burnett, C. W. Lee, S. Barr, S. Fuchs, S. E. Epstein. Marrowderived stromal cells express genes encoding a broad spectrum of arteriogenic cytokines and promote in vitro and in vivo arteriogenesis through paracrine mechanisms. Circulation Research 94 (2004): 678–85. Parekkadan, B., D. VanPoll, K. Suganuma, E. A. Carter, F. Berthiaume, A. W. Tilles, M. L. Yarmush. Mesenchymal stem cell-derived molecules reverse fulminant hepatic failure. PLoS One 2.9 (2007): e941. Van Poll, D., B. Parekkadan, C. H. Cho, F. Berthiaume, Y. Nahmias, A. W. Tilles, M. L. Yarmush. Mesenchymal stem cell-derived molecules directly modulate hepatocellular death and regeneration in vitro and in vivo. Hepatology 47.5 (2008): 1634–643. Németh, K., A. Leelahavanichkul, P. S. Yuen, B. Mayer, A. Parmelee, K. Doi, P. G. Robey, K. Leelahavanichkul, B. H. Koller, J. M. Brown, X. Hu, I. Jelinek, R. A. Star, É. Mezey. Bone marrow stromal cells attenuate sepsis via prostaglandin E2-dependent reprogramming of host macrophages to increase their interleukin-10 production. Nature Medicine 15 (2008): 42–49. De Waal Malefyt, R., J. Abrams, B. Bennett, C. G. Figdor, J. E. de Vries. Interleukin 10(IL-10) inhibits cytokine synthesis by human monocytes: an autoregulatory role of IL-10 produced by monocytes. Journal of Experimental Medicine 174 (1991): 1209–220. Deng, J., Y. Kohda, H. Chiao, Y. Wang, X. Hu, S. M. Hewitt, T. Miyaji, P. Mcleroy, B. Nibhanupudy, S. Lim, R. A. Star. Interleukin-10 inhibits ischemic and cisplatin-induced acute renal injury. Kidney International 60 (2001): 2118–128. Parekkadan, B. Cellular and molecular immunotherapeutics derived from the bone marrow stroma. Massachusetts Institute of Technology, Doctoral Thesis (2008). Lazarus, H., O. Koc, S. Devine, P. Curtin, R. Maziarz, H. Holland, E. Shpall, P. McCarthy, K. Atkinson, B. Cooper. Cotransplantation of HLA-identical sibling culture-expanded mesenchymal stem cells and hematopoietic stem cells in hematologic malignancy patients. Biology of Blood and Marrow Transplantation 11.5 (2005): 389–98. Baba, S. Clinical trials of regeneration for periodontal tissue. Home – ClinicalTrials.gov. (2005). Web. 18 Aug 2009. . Kastrup, J. Stem cell therapy for vasculogenesis in patients with severe myocardial ischemia.

MSC Immunoregulation Home – ClinicalTrials.gov. (2005). Web. 18 Aug 2009. . 23. Lee, O. K., T. K. Kuo, W. M. Chen, K. D. Lee, S. L. Hsieh, T. H. Chen. Isolation of multipotent mesenchymal stem cells from umbilical cord blood. Blood 103 (2004): 1669–675.

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24. Bianco, P., S. A. Kuznetsov, M. Riminucci, L. W. Fisher, A. M. Spiegel, P. G. Robey. Reproduction of human fibrous dysplasia of bone in immunocompromised mice by transplanted mosaics of normal and Gs a-mutated skeletal progenitor cells. The Journal of Clinical Investigation 101.8 (1998): 1737–744.

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Chapter 17 In Vitro Analyses of the Immunosuppressive Properties of Neural Stem/Progenitor Cells Using Anti-CD3/CD28-Activated T Cells Virginie Bonnamain, Isabelle Neveu, and Philippe Naveilhan Abstract Neural stem/progenitor cells (NSPCs) are multi-potent cells defined by their ability to self-renew and differentiate into cells of glial and neuronal lineage. Because of these properties, NSPCs have been proposed as therapeutic tools to replace lost neurons. Recent observations in animal models of immunerelated diseases indicate that NSPCs display immunomodulatory properties that might be a great interest for cell therapy. In particular, transplantation of NSPCs might be very useful as local immunosuppressive agent to promote the long-term survival of neuronal xenotransplant in the brain. To study this possibility, we have analysed the impact of NSPCs on anti-CD3/CD28-activated T cells. In vitro analyses clearly show that porcine, rat, and mouse NSPCs inhibit the proliferation of activated T cells. This result raises new perspectives concerning the use of NSPCs in cell therapy. Key words: Neural progenitor, Neural stem cell, T cell, Immunosuppression, Culture, Rat, Pig, Mouse

1. Introduction Neural stem/progenitor cells (NSPCs) are multipotent cells present in embryonic and foetal germinal zones (1). In the adult brain, NSPCs persist in particular areas such as the subventricular zone or the hippocampus. Because of their self-renewal capacity and their multi-lineage differentiation, NSPCs have received much attention (2–4). Indeed, NSPCs isolated from the fetal or adult mammalian central nervous system can be easily expanded in vitro as free-floating clusters upon treatment with EGF, basic fibroblast growth factor (bFGF) (5, 6). The expanded NSPCs display a high level of plasticity, giving rise to the three major neural Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_17, © Springer Science+Business Media, LLC 2011

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lineages – oligodendrocytes, astrocytes, and neurons – both in vitro and following transplantation in vivo (6–9). NSPCs are therefore considered as a very interesting source of transplantable cells in case of neurodegenerative disorders or stroke. Besides their utility to replace lost neural cells, NSPCs appear as an excellent tool in cell therapy because of their immunoregulatory properties. These unexpected properties have been recently brought to light by transplanting NSPCs in animal models of immune-related diseases. For instance, systemic or intraventricular transplantation of NSPCs in rodent model of multiple sclerosis attenuates the primary demyelinating process and reduces the acute axonal injury by decreasing brain inflammation (10–12). The mechanism by which transplanted NSPCs protect rat and mouse from EAE is not clear yet. Einstein et al., suggested that NSPCs act by inhibiting the proliferation and the activation of T lymphocytes in the lymph nodes (10) while Pluchino et al., thought that NSPCs exert neuroprotective effects by inducing programmed cell death of blood-borne, CNS-infiltrating pro-inflammatory Th1 cells (13). Whatever is the exact mechanism, both works substantiate the immunosuppressive property of NSPCs, and boost up further research to decipher the cross talk between NSPCs and T cells. Indeed, the ability of NSPC to dampen T cell immune response has many potential applications, including the use of NSPCs as local immunosuppressors in case of neuronal xenotransplantation in the brain (14, 15). As a first step to analyse the immunoregulatory impact of NSPCs on T cells in vitro, we have developed primary cultures of NSPCs and established co-culture of NSPCs with freshly isolated T cells. Our results indicate that mice, rat, and porcine NSPCs have immunosuppressive effect on spleenderived T cells.

2. Materials 2.1. Cell Culture

1. Sprague -Dawley rat embryos at 15 days of embryonic life (E15) and C57 Black mice embryos at E13 (Centre d’Elevage Janvier, Le Genest-Saint-Isle, France). 2. Domestic Large White porcine embryos at 28 days after artificial insemination (G28). 3. Hanks’ Balanced Salt Solution (HBSS) without phenol red (Sigma-Aldrich) supplemented with 100-U/ml penicillin and 0.1-mg/ml streptomycin (Gibco). 4. Basal medium: Dulbecco’s Modified Eagle’s Medium (DMEM)/ Ham’s F12 1:1 (Gibco) supplemented with 33-mM D-glucose, 5-mM HEPES (pH 7.2), 100-U/ml penicillin, and 0.1-mg/ml streptomycin, 2-mM L-glutamine.

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5. Complete medium: basal medium supplemented with 10% heat-inactivated fetal calf serum (FCS, Lonza, Basel, Switzerland). 6. Defined medium: Basal medium supplemented with N2supplement (Gibco). 7. RPMI medium: RPMI 1640 (Gibco) supplemented with 5-mM HEPES, 100-U/ml penicillin and 0.1-mg/ml streptomycin 2-mM L-glutamine, 1-mM sodium pyruvate (Gibco), 1-mM 2-mercaptoethanol, 1% non essential amino acids (Gibco), and 5% heat-inactivated FCS. 8. Phosphate buffered saline (PBS): Prepare 10× stock with 1.37-M NaCl, 27-mM KCl, 100-mM Na2HPO4, and 18-mM KH2PO4 (adjust to pH 7.4 with HCl if necessary). Prepare working solution by dilution of one part with nine parts of water and autoclave before storage at room temperature. 9. Trypsin TPCK-treated from bovine pancreas (Sigma-Aldrich) is dissolved in PBS at 25 mg/ml and deoxyribonuclease I from bovine pancreas (DNase I, Sigma-Aldrich) is dissolved in HBSS at 10 mg/ml. Both are stored in aliquots at −20°C and then used for tissue dissociation as required. 10. Human bFGF (Peprotech EC, London) is dissolved at 25 mg/ml in PBS supplemented with 4% bovine serum albumine (BSA). bFGF is stored in aliquots at −20°C and then added to cell culture dishes as required. 11. Sterile 70-mm filter (BD Biosciences). 12. Red cell lysis solution: NH4Cl 150 mM, KHCO3 10 mM, Na2–EDTA 0.1 mM, diluted in PBS (pH = 7.2). 13. Dynabeads® goat anti-mouse IgG (Invitrogen Dynal AS, Oslo, Norway). 14. The following mouse anti-rat monoclonal antibodies were used for cell depletion and T cell stimulation. OX42 (CD11b/c) and 3.2.3 (NKR-P1A) were obtained from the European Collection of Cell Culture (ECCC). HIS24 (CD45R) and G4.18 (CD3) were obtained from BD Biosciences Pharmingen (San Diego, CA, USA). The JJ319 mouse hybridoma producing anti-rat CD28 antibody was kindly provided by T. Hüning (University of Würzburg, Germany) and a stock solution at 2 mg/ml was prepared in sterile PBS. 15. Resuspension solution for T cell: 2% SVF and 0.5-mM EDTA in PBS. 16. Cell culture dishes (100 × 20 mm), Microtest™ tissue culture plate 96 wells, Multi-well culture plate (12 wells) are from BD Falcon™, Le Pont De Claix, France.

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2.2. Immunocytofluorescence

1. Microscope cover glasses, 20 mm diameter (Marienfeld GmbH and Co. KG, Lauda-Königshofen, Germany). 2. Poly-L-ornithine hydrobromide (PORN, Sigma-Aldrich) is dissolved at 500 mg/ml in tissue-culture water, stored in aliquots at −20°C, and then used for coverslip coating, as required. 3. PBT: Permeabilising/blocking solution : 1× PBS, 4% BSA, and 0.1% Triton X-100. 4. PBT-NGS: Permeabilising/blocking solution (1× PBS, 4% BSA, 0.1% Triton X-100, and 10% normal goat serum). 5. Primary monoclonal anti-rat antibodies: anti-Nestin (rat 401; Developmental Studies Hybridoma Bank, Iowa City, IA), anti-glial fibrillary acidic protein (GFAP, BD Pharmingen), anti-TUJ-1 (Sigma-Aldrich), and RIP (DSHB). 6. Secondary antibodies: fluorescein isothiocyanate (FITC)conjugated anti-mouse IgG (Jackson Immunoresearch, Cambridgeshire, UK). 7. Paraformaldehyde 16% solution (Electron Microscopy Sciences, Hatfield, PA, USA) is diluted at 4% in 1× PBS and stored at 4°C. 8. Sodium azide 0.1% in PBS. 9. Dabco anti-fading medium (Sigma-Aldrich). 10. Axioskop 2 plus microscope (Zeiss, Le Pecq, France). Micrographs are acquired with a 20× and 63× objectives using a digital camera (AxioCam HRC, Zeiss) driven by AxioVision Release 4.2 software.

2.3. T Cell Proliferation Assay

1. (3H) Thymidine (5 mCi) is stored at 4°C and added to the cells as required for the 12 last hours of the culture. 2. Cells are harvested on fibreglass using a harvester (Tomtec). 3. Radioactivity was measured by the standard scintillation technique using a Top Count NXT cell counter.

3. Methods 3.1. Primary Cultures of Rat, Mouse, and Porcine NSPCs (16)

1. Rat embryos at E15, mouse embryos at E13, and porcine embryos at G28 were collected by hysterectomy and sacrificed in the institute’s accredited slaughterhouse in accordance with the institutional guidelines of the Institut National de la Santé et de la Recherche Médicale (INSERM).

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2. Dissection includes isolation of the whole brain, cutting from the mesencephalon to the frontal cortex, above the eyes. The meninges are removed and the brain tissues are collected in a 15-ml Falcon tube containing 2 ml of ice-cold HBSS (see Note 1). 3. Continue the protocol under tissue culture laminar flow hood and use strict sterile technique. Put the brain tissues freed of meninges in a culture dish. Using a scalpel blade, mince tissue for ~30 s. 4. Using 1,000 ml plastic tips together with a Pipetman P1000, transfer all the minced embryonic tissues in a total volume of 5 ml of basal medium into a 50-ml Falcon tube. 5. Add 0.5 mg/ml of trypsin and incubate for 15 min in a 37°C water bath. 6. Return the tube to the hood and add 10 ml of complete medium (containing 10% FCS) to inhibit the enzymatic process. Leave the tube at room temperature for 5 min. 7. Add 0.1 mg/ml of DNase I and incubate the tube for 10 min in a 37°C water bath. 8. Dissociate the tissue digest mechanically with a 5-ml pipette, avoiding air bubbles. 9. Let the suspension settle for 5 min and transfer 10 ml of the cell suspension to a clean, labelled tube, leaving 5 ml behind. To the latter, triturate again ten times with a P1000. Let the suspension settle for 5 min. Transfer all but 200 ml from this tube to the labelled tube, thus pooling the cells from both trituration steps. 10. Pellet the cells by centrifugation at 50 × g for 10 min at RT. 11. Remove virtually all the supernatantand gently resuspend the pellet in complete medium, so as to bring the total volume of the resulting cell suspension to 1 ml. 12. Place the cells in 100 × 20 mm culture dishes in order to have the quantity of cells corresponding to five brains per dish and incubate in 10 ml of fresh complete medium. 13. Incubate at 37°C, 5% CO2 in a humidified incubator for 12 h. 3.2. Splitting of the Rat, Mouse, and Porcine NSPCs

1. After 12 h in complete medium, transfer culture dishes to the hood. Using a 10-ml pipette, aspire half of the medium and rinse two times to remove all the cells. 2. Collect the medium containing the floating cells into a 50-ml Falcon tube and pellet the cells by centrifugation at 50 × g for 10 min at RT.

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3. Suck off the supernatant and gently resuspend the cells in fresh defined medium. 4. Plate the cells in 100 × 20 mm culture dishes in order to have the quantity of cells corresponding to one brain per dish (1:5 split) and incubate in 10 ml of fresh defined medium. 5. Add bFGF at 25 ng/ml. 6. Incubate at 37°C, 5% CO2 in a humidified incubator. 7. Add bFGF every 2 days to stimulate the proliferation of NSPC as spherical clusters. These primary neurospheres should be ready for subculture 5 days after initial plating. 8. At day 5, bring culture dishes to the hood. Collect the medium containing the neurospheres and transfer it to a 50-ml Falcon tube, pulling the cells from each culture dish. 9. Pellet the cells by centrifugation at 50 × g for 10 min at RT and resuspend the neurospheres in 1 ml of fresh defined medium. Dissociate the neurospheres mechanically by gently pipetting up and down in order to get a homogenous cell suspension. 10. Plate the cells in new 100 × 20 mm culture dishes (1:2 split). Add bFGF as required and incubate in 10 ml of fresh defined medium. 11. Incubate at 37°C, 5% CO2 in a humidified incubator for another 5 days to allow the formation of secondary neurospheres (Fig. 1). Renew the addition of bFGF at day 7. 3.3. Differentiation of Rat NSPCs

1. In a tissue culture hood, place sterile glass coverslips in a 12-well plate (see Note 2). 2. Incubate coverslips in 50-mg/ml poly-L-ornithine (PORN) in sterile water for 2 h at 37°C in a tissue culture incubator. 3. Rinse PORN-coated coverslips with PBS and plate undissociated or mechanically dissociated rat neurospheres onto PORN-coated coverslips at a density of 200 neurospheres/ well or 2 × 105 cells/cm2, respectively, in 1 ml of complete medium. 4. Incubate the plate in a humidified incubator at 37°C, 5% CO2 and allow cells to adhere to the coverslips for 12 h (a minimum of 2 h is usually required). 5. After 12 h, replace the complete medium with 1 ml of defined medium. 6. Incubate the plate in a humidified incubator at 37°C, 5% CO2 and allow cells to differentiate. After 7 days, individual neurospheres should have dispersed in such a manner so as to appear as a flattened monolayer of cells (Fig. 1).

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Fig. 1. Culture of rat neural stem/progenitor cells (NSPCs). (a). Proliferation (a, b). NSPCs plated on uncoated dishes were grown as floating neurospheres for 10 days in defined medium supplemented with basic fibroblast growth factor (bFGF). Optical microscopy pictures were acquired with a ×4 (a) or ×10 (b) objectives. Differentiation (c, d). NSPCs plated onto polyornithine-coated glass coverslips, were allowed to differentiate for 7 days in defined medium free of growth factor. Optical micrographs were acquired after 4 days (c) or 7 days (d) of differentiation using a ×10 objective. Analyses of NSPC multipotency (b). After 10 days of proliferation, whole neurospheres (a, c, e, g) or mechanically dissociated neurospheres (b, d, f, h) were placed in differentiation conditions for 7 days to analyse the fate of rat NSPCs. Immunocytofluorescence was performed using primary antibodies directed against Nestin (a, b), glial fibrillary acidic protein (GFAP) (c, d), Tuj-1 (e, f), and RIP (g, h), and immunostaining was revealed with a FITC-conjugated anti-mouse IgG (a–h). Optical micrographs were acquired with ×10 (a, c, e, g) or ×63 (b, d, f, h) objectives using a digital camera (Zeiss).

7. Proceed to fix the cells by adding 4% paraformaldehyde in PBS. After 15 min at room temperature, wash the cells three times with PBS, and add 1 ml of PBS containing 0.1% Sodium Azide. Store the plate at 4°C until the immunostaining is performed (see Note 3).

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3.4. Characterisation of Rat NSPCs by Immunocytofluorescence (Fig. 1)

1. Wash the cells once with PBS to eliminate the traces of sodium azide. 2. Incubate the cells in PBT-NGS for 1 h at RT (see Note 4). 3. Dilute the primary antibodies in PBT (Nestin: 1/1,000; GFAP: 1/800; TUJ-1: 1/1,000; RIP: 1/3,000). Place 300 ml of diluted primary antibody per well and incubate at 4°C overnight. 4. Wash the cells three times with PBS, 5 min each time. 5. Dilute the secondary antibody in PBT (FITC-conjugated goat anti-mouse IgG: 1/250). Place 300 ml of diluted secondary antibody per well and incubate 2 h at RT in the dark. 6. Wash the cells three times with PBS, 5 min each time. 7. Mount the coverslips on a slide using Dabco antifading medium to minimise photobleaching (10 ml/20 mm coverslip). 8. Analyse the slides by fluorescence microscopy. 9. Slides are stored at −20°C.

3.5. Analyses of the Impact of NSPCs on T Cell Proliferation (Fig. 2)

1. Preparation of the assay plates. Prepare a 5-mg/ml solution of anti-CD3 in sterile PBS for the coating of assay plates. Calculate the number of wells required for each experimental condition and consider triplicate samples for each condition. Dispense 30 ml of the anti-CD3 antibody solution to each well of a flatbottom 96-well assay plate. Control wells are prepared by adding 30 ml of sterile PBS. Incubate the plate 2 h at 37°C. 2. Isolation and purification of T cells. T cells are obtained from 6- to 8-week-old female Sprague-Dawley rats. After sacrifice, the animals are splenectomised. The spleens are then placed in sterile cold PBS before isolation and purification of rat T cells (17).

Fig. 2. Modulation of T cell proliferation by NSPCs. Purified rat T cells (105 cells/well) were stimulated with anti-CD3/CD28 antibodies in the absence (0/1) or the presence of 2.5 × 104 (1/4) or 105 (1/1) irradiated (30 Gy) mouse (a), rat (b) or pig (c) NSPC per well. T cell proliferation was determined by analysing (3H)thymidine incorporation at day 3. Shown are means ± SEM of triplicate wells from 6 (a), 17 (b), and 7 (c) independent experiments. Statistical analysis: Kruskal–Wallis test followed by Dunn’s multiple comparison test. *p < 0.05; **p < 0.001; ***p < 0.0001 vs. the control (stimulated T cells in the absence of NSPCs).

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3. Under a sterile cell culture hood, pour the spleen and the medium in a cell culture dish. Cut the spleen in several pieces using a scalpel and transfer them into a sterile 70 mm filter placed in a dish. 4. Using the plunger of a 5-ml syringe, grind the spleen until a fine suspension is obtained. 5. Transfer the spleen-cell suspension to a 50-ml Falcon tube and complete to 50 ml volume with sterile PBS. Centrifuge at 900 × g for 10 min at 4°C. 6. Eliminate the supernatant and resuspend the pellet with 8 ml of red cell lysis solution. Incubate for 8 min at 4°C. 7. Add 42 ml of PBS and centrifuge at 900 × g for 10 min at 4°C. Resuspend the pellet with 10 ml of PBS/SVF (2%)/ EDTA (0.5 mM). Count the cells. 8. Incubate the cells for 15 min at 4°C with specific mAbs (clones 3.2.3 (0.7 mg/ml), HIS24 (1 mg/ml), and OX42 (0.5 mg/ml)) to deplete NK cells, B cells, and monocytes, respectively. Centrifuge the cells at 900 × g for 10 min at 4°C. 9. Resuspend the pellet with 2.5 ml of PBS/SVF/EDTA and incubate the cells with 500 ml magnetic anti-mouse IgG-coated Dynabeads for 20 min at 4°C on a wheel. 10. Preparation of irradiated NSPC. During this time, transfer the NSPCs to 50-ml Falcon tubes and submit them to g-irradiation (30 Gy). Centrifuge NSPCs at 50 × g for 10 min at 4°C and gently resuspend the pellet with 1 ml of defined medium using a P1000 to dissociate the neurospheres mechanically. Count the cells and adjust the concentration to 2 × 106 cells/ml. Perform 1:4 dilution to obtain another set of irradiated NSPCs at the concentration of 5 × 105 NSPCs/ml. 11. Preparation of T cell suspension. Collect purified T cells from splenocytes using a magnet and centrifuge the cells at 900 × g for 10 min at 4°C. Resuspend the pellet in 1 ml RPMI medium. Count the cells and adjust the concentration to 2 × 106 T cells/ml. 12. Preparation of the assay plates. Remove the anti-CD3 antibody solution of the flat-bottom 96-well assay plate with a multichannel pipettor. Rinse each well three times with 150 ml of sterile PBS. Discard PBS. 13. Co-cultures of T cells with increasing number of NSPCs. In each well, add 50 ml of T cell suspension at 2 × 106 cells/ml. Then, perform a rising scale by adding no NSPCs (ratio NSCPCs/ Tcells: 0/1), 50 ml of NSPCs at 5 × 105 cells/ml (ratio NSPCs/T cells: 1/4), or 50 ml of NSPC at 2 × 106 (ratio NSPC/T cells: 1/1). Complete to a final volume of 200 ml with a mix of defined medium and RPMI medium (vol/vol:1/1).

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14. Stimulation of T cells. Add anti-CD28 antibody (final concentration: 5 mg/ml) to the wells. 15. Incubate the plates for 72 h at 37°C with 5% CO2 in a humidified incubator. 16. Analyses of T cell proliferation. Add 25 ml of (3H)thymidine to each well (0.625 mCi/well) and incubate the plates for 12 h at 37°C, 5% CO2 in a humidified incubator. Harvest the cells on fibreglass filters using a harvester, and measure radioactivity by standard scintillation technique.

4. Notes 1. The dissection should be performed as quickly as possible (within 2 h), as the tissues become soft and sticky over time. 2. German glass coverslips are preferable for the culture of primary neural stem cells. To obtain sterile coverslips, dip the coverslips in a culture dish containing a solution of 70% ethanol for 5 min. Dry coverslips at RT for 30 min in a sterile cell culture hood. 3. When removing the solutions from the wells, be careful that suction does not dry out the cells. 4. As the antibodies are directed against intracellular antigens, the cell membranes are permeabilised with a detergent (Triton X-100). Non-specific binding of the antibodies is prevented by treating the cells with blocking agents such as BSA and NGS.

Acknowledgements The authors are very grateful to Dr. I. Anegon and Dr. Vanhove for their helpful advices. We also gratefully acknowledge Dr. P. Brachet and Pr. J.-P. Soulillou for their support and encouragement. We also express special thanks to “Etablissement Français du Sang” (EFS, Nantes) that kindly irradiated the NSPCs. The Nestin monoclonal antibody was developed by Susan Hockfield and obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by the University of Iowa, Department of Biological Sciences, Iowa City, IA 52242. The work was supported by the “Association Française contre les Myopathies” (AFM), the “Fédération des Groupements de Parkinsoniens”, and Progreffe. V.Bonnamain was supported by a fellowship from Ministère de l’Enseignement Supérieur et de la Recherche.

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References 1. Alvarez-Buylla, A., and Temple, S. (1998) Stem cells in the developing and adult nervous system. J. Neurobiol. 36, 105–110. 2. Svendsen, C. N., and Smith, A. G. (1999) New prospects for human stem-cell therapy in the nervous system. Trends Neurosci. 22, 357–364. 3. Martinez-Serrano, A., Rubio, F. J., Navarro, B., Bueno, C., and Villa, A. (2001) Human neural stem and progenitor cells: in vitro and in vivo properties, and potential for gene therapy and cell replacement in the CNS. Curr. Gene Ther. 1, 279–299. 4. Bjorklund, A., and Lindvall, O. (2000) Cell replacement therapies for central nervous system disorders. Nat. Neurosci. 3, 537–544. 5. Reynolds, B. A., Tetzlaff, W., and Weiss, S. (1992) A multipotent EGF-responsive striatal embryonic progenitor cell produces neurons and astrocytes. J. Neurosci. 12, 4565–4574. 6. Reynolds, B. A., and Weiss, S. (1992) Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255, 1707–1710. 7. Harrower, T. P., Tyers, P., Hooks, Y., and Barker, R. A. (2006) Long-term survival and integration of porcine expanded neural precursor cell grafts in a rat model of Parkinson’s disease. Exp. Neurol. 197, 56–69. 8. Smith, P. M., and Blakemore, W. F. (2000) Porcine neural progenitors require commitment to the oligodendrocyte lineage prior to transplantation in order to achieve significant remyelination of demyelinated lesions in the adult CNS. Eur. J. Neurosci. 12, 2414–2424. 9. Karbanova, J., Mokry, J., and Kotingova, L. (2004) Neural stem cells transplanted into intact brains as neurospheres form solid grafts composed of neurons, astrocytes and oligodendrocyte precursors. Biomed. Pap. Med. Fac. Univ. Palacky Olomouc Czech Repub. 148, 217–220. 10. Einstein, O., Fainstein, N., Vaknin, I., Mizrachi-Kol, R., Reihartz, E., Grigoriadis, N., Lavon, I., Baniyash, M., Lassmann, H., and Ben-Hur, T. (2007) Neural precursors attenuate autoimmune encephalomyelitis by peripheral immunosuppression. Ann. Neurol. 61, 209–218.

11. Einstein, O., Grigoriadis, N., Mizrachi-Kol, R., Reinhartz, E., Polyzoidou, E., Lavon, I., Milonas, I., Karussis, D., Abramsky, O., and Ben-Hur, T. (2006) Transplanted neural precursor cells reduce brain inflammation to attenuate chronic experimental autoimmune encephalomyelitis. Exp. Neurol. 198, 275–284. 12. Pluchino, S., Quattrini, A., Brambilla, E., Gritti, A., Salani, G., Dina, G., Galli, R., Del Carro, U., Amadio, S., Bergami, A., Furlan, R., Comi, G., Vescovi, A. L., and Martino, G. (2003) Injection of adult neurospheres induces recovery in a chronic model of multiple sclerosis. Nature 422, 688–694. 13. Pluchino, S., Zanotti, L., Rossi, B., Brambilla, E., Ottoboni, L., Salani, G., Martinello, M., Cattalini, A., Bergami, A., Furlan, R., Comi, G., Constantin, G., and Martino, G. (2005) Neurosphere-derived multipotent precursors promote neuroprotection by an immunomodulatory mechanism. Nature 436, 266–271. 14. Remy, S., Canova, C., Daguin-Nerriere, V., Martin, C., Melchior, B., Neveu, I., Charreau, B., Soulillou, J. P., and Brachet, P. (2001) Different mechanisms mediate the rejection of porcine neurons and endothelial cells transplanted into the rat brain. Xenotransplantation 8, 136–148. 15. Michel, D. C., Nerriere-Daguin, V., Josien, R., Brachet, P., Naveilhan, P., and Neveu, I. (2006) Dendritic cell recruitment following xenografting of pig fetal mesencephalic cells into the rat brain. Exp. Neurol. 202, 76–84. 16. Sergent-Tanguy, S., Veziers, J., Bonnamain, V., Boudin, H., Neveu, I., and Naveilhan, P. (2006) Cell surface antigens on rat neural progenitors and characterization of the CD3 (+)/CD3 (−) cell populations. Differentiation 74, 530–541. 17. Dugast, A. S., Haudebourg, T., Coulon, F., Heslan, M., Haspot, F., Poirier, N., Vuillefroy de Silly, R., Usal, C., Smit, H., Martinet, B., Thebault, P., Renaudin, K., and Vanhove, B. (2008) Myeloid-derived suppressor cells accumulate in kidney allograft tolerance and specifically suppress effector T cell expansion. J. Immunol. 180, 7898–7906.

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Part II Regulatory Molecules

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Chapter 18 Immunoregulatory Properties of Heme Oxygenase-1 Philippe Blancou, Virginie Tardif, Thomas Simon, Séverine Rémy, Leandro Carreño, Alexis Kalergis, and Ignacio Anegon Abstract Heme oxygenase-1 (HO-1) is one of the three isoforms of the heme oxygenase enzyme that catabolyzes the degradation of heme into biliverdin with the production of free iron and CO. HO-1 is induced by its substrate and by other stimuli, including agents involved in oxidative stress and proinflammatory cytokines as well as several anti-inflammatory stimuli. A growing body of evidence points toward the capacity of this molecule to inhibit immune reactions and the pivotal role of HO-1 in inflammatory diseases. We will first review the physiological role of HO-1 as determined by the analysis of HO-1-deficient individuals. This will be followed by an examination of the effect of HO-1 within immunopathological contexts such as immune disorders (autoimmunity and allergy) or infections. A section will be devoted to the use of an HO-1 inducer as an immunosuppressive molecule in transplantation. Finally, we will review the molecular basis of HO-1 actions on different immune cells. Key words: Heme oxygenase, Carbon monoxide, Inflammation, Autoimmunity, Allergy, Infection, Transplantation, Dendritic cell, Lymphocyte, Macrophage

1. HO-1 Has Immunosuppressive Effects Throughout the Immune System 1.1. HO-1 Is a Natural Immunosuppressive Molecule

Table 1 collects most of the references of this section.

The first important step toward our comprehension of HO-1 function was made more than a decade ago when K.D. Poss and S. Tonegawa generated the first hmox1 (HO-1 coding gene) knock-out (KO) mice (2), shortly followed by M. Lee’s group (3). Even though hmox1−/− mice were difficult to breed due to a high rate of prenatal mortality, some of them survived. However, these survivors subsequently developed anemia due to iron

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_18, © Springer Science+Business Media, LLC 2011

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Inhibition of MHC II expression by APC within the CNS Inhibition of Th and CD8 T-cell accumulation, proliferation, and effector function within the CNS Persistent activation of antigen-presenting cells Enhanced infiltration of Th17 cells in CNS Nonregressing myelin-specific T-cell reactivity Regulation of IFN-b production

HO-1- deficient mice CoPP administration CO inhalation

Conditional ablation of HO-1 expression in myeloid cells (HO-1M-KO mice)

EAE

Allergic asthma (mouse model)

Allergy

Inhibition of DC immunogenicity Loss of autoreactive CD8+ T-cell function

HO-1 inducer (CoPP) CO release molecule 2 (CORM2)

DC-induced autoimmune diabetes model

Increase in percentage and suppressive function of CD4+CD25(high) Tregs, FoxP3 mRNA expression, and IL-10 level in sera Increase in percentage and suppressive function of CD4+CD25(high) Tregs Increase in FoxP3, IL-10, and membrane bound TGF-b 1 expression in sera, splenic CD4+CD25+ Tregs cell, and lung

HO-1 inducer (Hemin) HO-1 inhibitor (SnPP)

HO-1 inducer (Hemin) HO-1 inhibitor (SnPP)

Increase in antiapoptotic protein level in pancreas Increase b-cell survival in pancreatic islets Decrease in CD11c+ DC infiltration in pancreas

CoPP/SnPP administration

Curative treatment

Downregulation of MHC II+ DC population Downregulation of type 1 T helper cell mediated response

Mechanism

AAV-HO-1 injection CO inhalation

Treatment

Preventive treatment

NOD mice

Type 1 diabetes

Autoimmunity

Pathology/Model

Table 1 Effects of HO-1 in immunopathology and transplantation

(32)

(29)

(22)

(35)

(43)

(37, 105)

(36)

References

248 Blancou et al.

No inhibition of parasitemia Preservation of liver function by antioxidant effect of HO-1 that prevents hepatocytes from undergoing TNF-mediated apoptosis

HO-1- deficient mice AdHO-1 injection CO inhalation

Noncerebral severe malaria Plasmodium chabaudi chabaudi

In the presence of poly I/C: enhanced bacterial clearance and improved survival of HO-1M-KO mice caused by reduced type I IFN-mediated apoptosis of macrophages NO-induced HO-1 upregulation during Salmonella infection via 8-nitro-cGMP formation Impaired host defense against Salmonella by HO-1 inhibition (higher bacterial survival and apoptotic cell level)

Conditional ablation of HO-1 expression in myeloid cells (HO-1M-KO mice)

HO-1 SiRNA HO-1 inhibitor (PEG-ZnPP)

Listeria monocytogenes

Salmonella enterica serovar Typhimurium

(continued)

(17)

(22)

(21)

No inhibition of parasitemia Prevention of blood–brain barrier disruption, brain microvasculature congestion, and activated CD8+ T cell-mediated neuroinflammation by avoiding hemoglobin oxidation and free heme release via CO binding to hemoglobin

HO-1- deficient mice a CoPP, administration a CO, inhalation

Experimental cerebral malaria (ECM) Plasmodium berghei

Bacterial infections

(26)

Inhibition of inflammatory cytokine level (promotion of the liver stage infection of plasmodium)

HO-1- deficient mice siRNA targeting HO-1 AdHO-1 injection a CO, inhalation

Liver infection Plasmodium berghei and Plasmodium yoelii

Malaria (24, 25)

(30)

Suppression of type 1 and type 2 T-cell dependent inflammation Dominant effect of HO-1 overexpression on antigen-presenting cells

HO-1 inducer (CoPP) HO-1 inhibitor (SnPP)

Skin inflammation Mice model of induced contact hypersensitivity Mice model of induced late phase reaction

Infections

(31)

Attenuation of skin lesions Increase in serum IL-18 level Low serum IgE level

HO-1 inducer (Hemin) HO-1 inhibitor (SnPP)

Atopic dermatitis (DS-Nh mouse)

References

Mechanism

Treatment

Pathology/Model

Immunoregulatory Properties of Heme Oxygenase-1 249

Warm I/R (mouse model)

Liver

I/R

No effect on intrahepatic TLR-4 expression Downregulation of IP-10 and proinflammatory cytokines Inhibition of Phospho-STAT1 and CXCL10 Upregulation of intrahepatic HO-1 gene expression Inhibition of neutrophil infiltration Inhibition of local and systemic expression of TNF-a in TLR-4-deficient mice Inhibition of HO-1 in TLR-4-deficient mice restore I/R damage

TLR-4-deficient mice SnPP administration

Impaired IFN-b production and IRF-3 phosphorylation after macrophage SeV infection

CoPP administration

Conditional ablation of HO-1 expression in myeloid cells (HO-1M-KO mice)

Paramyxoviruses (SeV)

Transplantation

HO-1 inducer (CoPP) AdHO-1 transfer

Hepatitis B virus

Reduction of hepatic HBV replication and HBV viremia by interfering with capsid formation

(80)

(74, 75)

(22)

(23)

(15, 16, 25)

Upregulation of HO-1 in macrophages and mice during MTB infection Induction of MTB dormancy regulation by CO (but not biliverdin or iron)

HO-1- deficient mice CO gas treatment HO-1 inhibitor (SnPP)

Mycobacterium tuberculosis (MTB)

Virus infections

(13)

Cortical nonheme iron increase due to the induction of HO-1 Prevention of iron-induced oxidative damage by ferritin upregulation independently of HO-1 activity

HO-1 inhibitor (SnPP)

References

Experimental pneumococcal meningitis

Mechanism

Treatment

Pathology/Model

Table 1 (Continued)

250 Blancou et al.

CO + Biliverdin (dual treatment)

CO inhalation (recipient) CO-satured preservation solution Biliverdin administration

CO-satured preservation solution

Heart

Intestine

Recombinant AdHO-1 (i.g., i.v., or i.m. injection)

HO-1 transgenic mice (as donor or recipient) ZnPP administration

Mouse cardiac model

CO-satured preservation solution

Rat cardiac model

Acute rejection

Lung (cold I/R)

Improved intestinal barrier function Increased intragraft cyclic guanosine monophosphate (cGMP) levels Protective effect of CO reversed by soluble guanylyl cyclase inhibition

CoPP (donor)

Pancreas

Biliverdin administration (donor recipient)

Increased intestinal graft blood flow Inhibition of inflammatory mediator upregulation Improved intestinal injuries and transit

CO inhalation (recipient)

Kidney

(58)

Inhibition of vasculitis and inflammatory cell infiltrate Inhibition of CD4(+) lymphocyte infiltration and CD25 expression (HO-1 Tg as recipient)

(continued)

(57)

(106)

(47–49)

(53)

(64)

(50, 51)

(52)

References

i.v. injection: decreased the number of graft-infiltrating leukocytes, cytokine mRNA accumulation, and apoptosis in transplanted hearts No general immunosuppression

Improved gas exchange Reduction of intragraft inflammation (reduced inflammatory mediators and cellular infiltration)

Decreased neutrophil infiltration Improved intestinal circular muscle contractility

Inhibition of I/R injury induced inflammatory mediators

Inhibition of proinflammatory cytokine expression Increase in anti-inflammatory cytokine expression

Inhibition of inflammatory mediator expression Reduced macrophage infiltration Reduced tubular epithelial cell apoptosis Upregulation of vascular endothelial growth factor (VEGF) via activation of its upstream hypoxia-inducible factor (HIF)-1

Inhibition of I/R injury induced STAT1 and STAT3 expression Downregulation of MEK/ERK1/2 signaling pathway Inhibition of early proinflammatory and stress-response gene expression

CO inhalation (recipient)

Cold I/R (rat model)

Mechanism

Treatment

Pathology/Model

Immunoregulatory Properties of Heme Oxygenase-1 251

Inhibition HO-1 (SnPP)

Inhibition HO-1 (SnPP)

Induced by recipient adult MSC

Breaking of tolerance

Breaking of tolerance (in vitro prevention of inhibition of T cell proliferation)

Inhibition of apoptotic islets Inhibition of lymphocytic infiltration in engrafted islets

Blockade of islet TLR4 expression

CO inhalation (donor) CO exposure of pancreatic islet

Ad-HO-1 islet transfection

(67)

Promotion of migration and de novo generation of recipient-derived FoxP3+ Treg (combined treatment)

CoPP administration: donor CO treatment: donor or recipient or both, isolated islets Bilirubin administration: recipient

(62)

(61)

(39)

(38)

(66)

Suppression of intragraft macrophage infiltration (donor treatment)

Bilirubin administration: donor or recipient or islets

(65)

(55)

(54, 56)

References

Decreased intragraft mRNA expression of proinflammatory genes Increased intragraft mRNA expression of antiapoptotic genes Suppression of intragraft macrophage infiltration (donor treatment)

Induced by recipient iBMDC

Rat tolerance model

Rat model

Mice model

Inhibition of allograft T-cell infiltrationNo difference in activated T cells in draining lymph nodes

Inhibition of leukocyte and VSMC infiltration Inhibition of costimulation molecules, adhesion molecules, and cytokines

Mechanism

CoPP administration: donor or recipient or both CO treatment: donor or recipient or both, isolated islets

Hemin donor i.p. administration

Mouse aorta model

Pancreatic islet

Recombinant AdHO-1 (donor lumen aortas), a methylene chloride orally administrated

Treatment

Rat aorta model

Chronic rejection

Pathology/Model

Table 1 (Continued)

252 Blancou et al.

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accumulation in the liver and kidney as a result of the role of HO-1 in recycling iron from these organs. Importantly, this anemia was associated with a chronic inflammatory state characterized by an increased secretion of immunoglobulin and cytokines after LPS- or CD3/28-stimulation of splenocytes, with a concomitant deviation of the Th response into a Th1-type cytokine profile (4). Of note, no increase in HO-2 protein was observed in the spleen of these animals as a compensatory mechanism (4, 5). HO-1 KO mice are also more susceptible to polymicrobial sepsis due to lower bacterial clearance mediated by HO-1-derived CO phagocytosis activity of Gram-positive bacteria (6). Two years after the description of HO-1 KO mice, Yachie et al. described the first and unique case to date of human HO-1-deficiency (7). The clinical picture of human and murine HO-1 deficiency is strikingly similar from an immunological perspective, with all the symptoms of chronic inflammation (lymph node swelling and leukocytosis), suggesting that HO-1 activity modulates the inflammatory response in a similar fashion in both species. The question arises as to how the absence of HO-1 in humans and mice leads to chronic inflammation. Different hypotheses can be put forward. For example, a high level of monocyte chemoattractant protein-1 (MCP-1 or CCL2) in HO-1 KO mice compared to wild-type mice is the hallmark of an inflammatory phenotype (8, 9), suggesting a defect in the control of tissue inflammation by myeloid cells. While Otterbein et al. showed that CO stimulates the synthesis of the anti-inflammatory cytokine IL-10 by macrophages (10), Lee et al. demonstrated that IL-10 induces HO-1 expression, potentially leading to a self-amplification of the anti-inflammatory effect (11). 1.2. HO-1 Is Most of the Time an Ally but Can Sometimes Be an Enemy When Fighting Infections

The role of HO-1 in viral or bacterial infection is complex due to the multifunctional role of this protein. HO-1 can be induced by a wide variety of bacteria such as Rickettsia rickettsii (12), Streptococcus pneumoniae (13), Enterohemorrhagic Escherichia coli (14), Mycobacterium tuberculosis (15, 16), Salmonella enterica (17) as well as viruses such as Kaposi sarcoma-associated herpes virus (KSHV) (18), and Hepatitis C virus (HCV) (19, 20). This induction is probably related to the increase in oxidative stress and inflammation leading to Hmox-1 gene transcription. HO-1 is not only passively induced, its expression protects mice against infectious diseases such as S. enterica (17), Plasmodium chabaudi (21), Listeria monocytogenes (22), Hepatitis B virus (HBV) (23), and Pneumococcal meningitis (13) infections. The molecular basis of such protection can be linked to direct in vitro inhibition of HBV replication (23) or S. enterica multiplication (17). In certain other infections, the infection is associated with HO-1 upregulation to prevent iron-induced oxidative damage such as in the Plasmodium species (21) or in Pneumococcal meningitis (13) infections.

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By contrast, in certain infections, HO-1 was found to facilitate pathogen growth, i.e., in the Plasmodium species (24) and Mycobacterium tuberculosis (25). In the latter case, it was shown that CO resulting from HO-1 activity is a strong stimulator of dormancy regulation in bacteria (15, 16). Interestingly, the role of HO-1 in plasmodium infection is twofold in that it reduces neutrophils and macrophages infiltration at the beginning of the infection (24) and subsequently protects against free heme release that triggers cerebral malaria pathogenesis (26). 1.3. HO-1 Can Regulate Immune Disorders

The observation of HO-1 upregulation in different rodent models of allergy (27, 28) together with the anti-inflammatory properties of this molecule has led to the hypothesis that HO-1 could play a protective role in this disease. HO-1 upregulation produced a significant protective effect against airway inflammation in a mouse model of allergic asthma (29). HO-1 inducers also strongly inhibit other allergies such as skin allergy, both in the mouse (30, 31) and in humans (31). The precise underlying mechanisms for HO-1based protection against allergic inflammation have not yet been completely understood. At least part of the protective effect of HO-1 in asthma may depend upon the prevention of free heme from participating in prooxidant reactions, and in regulating vascular tone, cell growth, or apoptosis. This effect is associated with the expansion and enhanced suppressive function of CD4+CD25+ Treg populations (29, 32). However, in the rat, HO-1 acts directly on the immune response by suppressing IgE-induced mast cell degranulation (33) and cytokine synthesis (34). Other important immune disorders are autoimmune diseases. Ablation of the HO-1 gene has never been associated with an increased incidence of spontaneous autoimmune disease in humans or mice so far. This is corroborated in the context of diabetes by the fact that there is no association between the Hmox-1 locus and type 1 diabetes (T1D) susceptibility locus in the NOD mouse model or in the human (personal communication). However, HO-1 induction reduces experimental autoimmune encephalomyelitis (EAE) symptoms whereas HO-1 KO mice develop an exacerbated form of the disease (35). In the former model, infiltration of the CNS by T-lymphocytes is reduced and the lymphocytes themselves display a nonaggressive Th phenotype. The latter model has recently been completed by G. Kolias’ group who show that the exacerbated form of the disease is a consequence of the absence of HO-1 in macrophage/granulocytes lineage (22). In addition, Chora et al. demonstrated that CD11c+ cells from HO-1-treated animals have a lower antigen-presenting capacity associated with MHC class II downregulation. Similarly, HO-1 induction or CO treatment lowers the incidence of diabetes (36) and can even cure recently diabetic NOD mice (36, 37). This protective effect can be linked to the general cytoprotective,

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antiapoptotic, vascular effect of HO-1 on the pancreas or probably more specifically to TLR4 downregulation on beta cells (see transplantation section (38)). However, HO-1 also seems to act directly on the immune system as adoptive transfer experiments have demonstrated that splenocytes isolated from HO-1-induced or CO-treated mice are less diabetogenic in the NOD mouse model (36). This protection is associated with lower T-cell infiltration in the pancreas following HO-1 induction in spontaneous diabetes in mice (37) or in rat islet allotransplantation (39). We have recently generated data in both models of spontaneous or induced diabetes in mice showing that CO-treated DC impair autoreactive CD8 T-cell infiltration in the pancreas (submitted manuscript). The cellular basis of such effects on autoimmune and allergic diseases remains elusive. A generalized function of HO-1 may include a reduction in inflammatory cell rolling, adhesion, and migration from the vascular compartment, possibly by downregulating the function and expression of adhesion molecules on the vessel wall in both humans (40) and mice (41). However, certain hypotheses have been forged as to the direct role of HO-1 on immune cells. A possibility worth contemplating is that in vivo HO-1 induction leads to impaired antigen presentation by DC (35), which in a tolerogenic environment (42) causes loss of function of autoreactive CD8 T cells (43) or the induction of Treg expansion/gain of function (32). 1.4. HO-1 Is a Promising Immunosuppressive Molecule in Transplantation

Depending on the model, HO-1 improves graft survival via graft cytoprotective and local anti-inflammatory effects as well as systemic immunosuppressive effects. The local protective role of HO-1 within the transplant was initially demonstrated using HO-1-deficient mouse heart grafts that were rapidly rejected after transplantation, in contrast to HO-1-nondeficient mouse hearts that survived indefinitely (44). Several years later, the ability of HO-1 to suppress the rejection of mouse-to-rat cardiac transplants was shown to be dependent on the generation of CO (45). On the other hand, the systemic protective role of HO-1 has also been reported in a mouse cardiac transplantation model in which tolerance was induced by the administration of anti-CD40L antibody and donor-specific transfusion (DST). In fact, in the same treatment conditions, the long-term survival in HO-1-deficient animals was abrogated (46). The local and systemic cytoprotective effects of HO-1 against intestinal ischemia/reperfusion (I/R) injury have also been reported, notably after the treatment of the recipient with CO (47), ex vivo administration of CO to the graft (48), or biliverdin administration to the recipient, donor, and graft (49). All of these strategies result in significantly improved animal survival compared with the controls. The same results were obtained in different

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rodent I/R models when recipients were treated with CO (50–52). It is important to note that in some models, a combination of CO and biliverdin treatment provides enhanced protection against I/R injury because of the distinctive action of these two products, i.e., CO improves graft blood flow and biliverdin decreases lipid peroxidation (53). The local graft protective effect of HO-1 has additionally been studied in the rodent model of aorta chronic rejection (54, 55). Donor aortas previously transduced with a recombinant adenovirus coding for human HO-1 cDNA (AdHO-1) (54) or pretreated with hemin (an HO-1 inducer) (55) showed a significant decrease in intimal thickening, due to both a significant reduction in leukocyte infiltration (notably T-cell infiltration) (55), and vascular smooth muscle cells (VSMC) (54) in the intima, compared to untreated aortas or aortas transduced with the control adenovirus. Moreover, CO delivery brought about a more pronounced inhibition of intimal VSMC infiltration compared to AdHO-1 treatment (54). Interestingly, aortas treated with AdHO-1 showed a more considerable reduction in the number of leukocytes as well as in the expression of costimulation molecules, adhesion molecules, and cytokines than those treated with CO (56). In a rat model of acute cardiac allograft rejection, intragraft (i.g.) cardiac injection of AdHO-1 as well as intramuscular (i.m.) or intravenous (i.v.) injection, prolonged allograft survival with more efficiency after AdHO-1 i.v. injection, suggesting not only a local graft protective effect but also a systemic immunosuppressive effect for HO-1 (57). Improved cardiac allograft survival was also observed after systemic HO-1 overexpression by using HO-1 transgenic mice as recipients (for systemic HO-1 expression) or donors (for local graft HO-1 expression), but survival was longer for systemic HO-1 expression vs. local graft expression (58). The results from these last two manuscripts strongly suggest that HO-1 acts more prominently in reducing the immune response, rather than protecting graft cells, as supported by the inhibition of allogeneic responses in MLRs using splenocytes from recipients treated with AdHO-1 administrated intravenously (57). When only purified T cells from splenocytes or mesenteric lymph node cells were used as stimulators, no inhibition of MLR was detected, suggesting a decreased proliferation dependent on the presence of non-T cells such as APC. In addition, the implication of DCs in the HO-1- or CO-induced immunologic mechanism has been reported in rat kidney transplantation (59). In fact, kidney donor pretreatment with an HO-1 inducer or CO led to a significantly reduced mRNA expression of MHC II, CD80, CD86, IP-10, and CCL19 as well as of all three inducible proteasome subunits, i.e., LMP2, LMP7, and MECL1, in the recipient spleen compared to untreated control spleens (59). In contrast, intragraft mRNA expression of all these molecules was unaffected by HO-1 or CO

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pretreatment of the donor, suggesting that HO-1 or CO targets the antigenicity and migration of donor DCs. Using DCs or others cells, such as mesenchymal stem cells (MSCs) or embryonic stem cells (ESCs), as therapeutic tools is now commonly advocated, especially in the field of transplantation when the aim is to induce tolerance. Nevertheless, cell therapy requires a sound understanding of the molecular pathways involved in the regulation of the immune response by these different cells and the biology of the in vitro-generated cells. Again, HO-1 has been shown to play a role in the active inhibition of immune responses by these cells. In fact, in a rat heart transplantation model, injection of syngeneic immature DCs generated from bone marrow progenitors (corresponding to adherent DCs) one day prior to transplantation was shown to prolong allograft survival (60). This prolongation of allograft survival induced by tolerogenic DCs was totally abrogated by the administration of an inhibitor of HO-1 activity (61). In the same rat model, the prolongation of cardiac allograft survival was also reproduced by two injections of adult rat MSCs 7 days and 1 day before transplantation (62). As described above for DCs, the inhibition of HO-1 activity, but also the inhibition of iNOS activity, totally reversed the protective effect of adult rat MSCs and induced graft rejection (62). In contrast, in the MLR assay using human cells, only the inhibition of HO-1 but not the inhibition of iNOS completely abolished the suppressive effect of human adult MSCs on allogeneic proliferation, suggesting a key role for HO-1 in the human MSC-mediated immunosuppression (62). In fact, the suppressive effect of human embryonic stem cells (hESCs) observed on PBMC and T-cell proliferation was also reversed by the inhibition of HO-1 activity, providing a hypothetical mechanism for protecting hESCs in vivo (63). In pancreas transplantation, HO-1 induction in the donor led to improved rat allograft survival and unimpaired endocrine and exocrine function compared to controls (64). Likewise, HO-1 induction, bilirubin administration, or CO treatment of the donor, recipient, or both led to long-term survival of islet grafts transplanted in a partially MHC-incompatible strain combination, from DBA/2 (H-2d) to B6AF1 (H-2b,k/d) mice (65, 66). Moreover, recipients of long-term surviving islet grafts showed antigenspecific tolerance to a second islet graft from the first-donor strain, whereas all third party islet grafts were rejected (65). The immunologic mechanism implicated in the CO-treated group involved a reduction in macrophage infiltration concomitant to a significant decrease in the mRNA of MCP-1, a chemokine secreted by b cells resulting in the migration of macrophages. By using a mouse islet allograft model but in a stronger immunogenic strain combination with a complete MHC incompatibility (Balb/C (H-2d) to C57BL/6 (H-2b)), a more recent publication showed that the best

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results in terms of long-term islet allograft survival were obtained with combined treatment, i.e., donors treated with an HO-1 inducer as well as with CO inhalation or donor islets cultured in a CO chamber and recipients exposed to CO, in addition to bilirubin administration. HO-1 has been shown to act on the immune response by promoting the migration and the de novo generation of recipient-derived Foxp3+ Treg cells in mice treated with the combined protocol (67). In addition, the CD4+CD25+ T cells have been shown to be essential for maintai ning the efficiency of the combined protocol. In fact, no prolongation of islet allograft survival can be observed after depletion of CD4+CD25+ T cells in recipients even when the combined protocol is performed. In rats, prolonged islet allograft survival associated with lower intragraft lymphocyte infiltration has also been reported after islet transduction with AdHO-1 (39). On the other hand, a combination of HO-1 with other immunosuppressive techniques such as the immobilization of polyethylene glycol (PEG) molecules on islets, which is known to attenuate the immune response, has been shown to reduce immunosuppressive therapy (68). TLR4 expression by pancreatic b cells has also been implicated in determining the outcome of islet allografts. When b cells are isolated from the pancreas, TLR4 is upregulated on the b-cell membrane in response to the stress caused by isolation process. In fact, CO exposure of the donor reduced TLR4 expression on b cells (38). The use of TLR4-deficient islets for transplantation induced a long-term allograft survival compared to wild-type islets, and reduced inflammation as seen by a reduced intragraft expression of TNF-a, iNOS, MCP-1, and Fas. Thus, CO exposure of pancreatic islets possibly induced long-term allograft survival by blocking TLR4 expression (38). However, suppression of the donor inflammatory response by knocking out TLR4 was not sufficient to prevent islets from rapidly rejecting if HO-1 activity was blocked in the recipient. Thus, HO-1 activity is still probably required for an effective anti-inflammatory response in the recipient. TLR4 is also known to play an important role in the development of an inflammatory response after renal (69), heart (70, 71), and liver (72) I/R injury or in the pathogenesis of chronic cardiac allograft rejection (73), at least partly through CXCL-10 production. Thus, reduced TLR4 expression could also explain the protective role of CO in I/R injury. HO-1 activity or CO treatment can interfere with TLR4 signaling by downregulating STAT-1 activation via the type 1 IFN pathway, which in turn decreases CXCL-10 production (74, 75). In macrophages, CO has been shown to reduce TLR4 signaling through inhibition of lipid raft formation (76). In addition, human TLR4 genes can be encoded by two loss-of-function SNPs (Asp299Gly and Thr399Ile), characterized by lower binding to the LPS receptor

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(77) and identical TLR4 genes and protein expression (78). These TLR4 mutations in human kidney transplants were associated with improved immediate renal graft function, but not with the incidence of acute rejection (79). Mutated TLR4 was linked both to a significant intragraft increase in HO-1 gene expression and to a significant decrease in that of TNF-a and MCP-1 (79). This suggests a cross talk between the HO-1 and TLR system that has been previously reported in a murine model of hepatic I/R, in which TLR4-deficent mice showed reduced I/R damage compared to wild-type mice (80). HO-1 gene polymorphism has also been shown to improve allograft outcome. In fact, HO-1 expression is modulated by the presence of (GT)n polymorphism and an SNP (A-413T) in the promoter. The HO-1 promoter activity increases with the short (GT)n allele or the A-allele. Renal allograft function (81, 82) and survival improved and the likelihood of developing chronic allograft nephropathy (CAN) was reduced (83) when transplanted kidneys were carriers of the short (GT)n allele. However, these results contradict those of another study showing no impact of the short (GT)n allele on acute rejection or CAN incidence (84). This divergence in results could be explained by the difference in sample sizes between the two studies (type 1 error) (85). The impact of the (GT)n polymorphism has also been studied for other organ transplants. No association of the (GT)n allele was found with the development of cardiac allograft vasculopathy (86) or with outcome after liver transplantation (87). However, the presence of the A-allele in the promoter of HO-1 in the liver donor correlated with a significantly better 1-year graft survival as well as with less frequent graft loss due to primary dysfunction compared with the TT-allele genotype (87). Interestingly, the two most frequent haplotypes encountered are the A-allele combined with a long (GT)n allele and the T-allele combined with a short (GT)n, which is counterproductive because the A-allele and a short (GT)n allele are associated with higher HO-1 promoter activity. However, in terms of clinical outcome and hepatic HO-1 mRNA level, the A(-413) SNP is found to be dominant over the (GT)n polymorphism (87). These findings thus provide another explanation for the divergence in results described above, in which only the presence of (GT)n allele was studied.

2. Possible Mechanism of Action of HO-1 in Immune Cells 2.1. HO-1 Exerts Limited Direct Effect on T-Lymphocytes

Table 2 collects most of the references of this section. T-cell functions can be modified either by endogenous expression of HO-1 or by extracellular HO-1-derived products.

HO-1 induction (+LPS)

Rat

Reduction of mRNA expression of CD80, CD86, and all three inducible proteasome subunits, i.e., LMP2, LMP7, and MECL1

(42, 30, 43)

Inhibition of IRF-3 phosphorylation Reduction of ROS level Inhibition of phenotypic maturation (CD80, CD86…) Inhibition of proinflammatory cytokines (IL-12, IL-6, TNF-a, and IFN-b) secretion Increase or maintenance of IL-10 production Impairment of stimulation capacities

HO-1 induction or CO treatment (+LPS)

Impairment of genes induction encoding RANTES, IP-10, and MCP-1 Reduction of IRF-3 activation Impairment of IFN-b production

HO-1 inactivation (+ LPS or poly IC)

Human/ rat/mouse

Reduction of NAD(P)H oxidase activity and ROS level Inhibition of TLR4 translocation to lipid raft Suppression of TLR4 and TRIF/MyD88 interactions Inhibition of transcriptional factors (NF-KB and IRF-3) activation (not with poly IC) Reduction of IFN-b gene expression (not with poly IC activation) Reduction of IP-10 and RANTES production (not with poly IC activation)

CO treatment (+LPS)

Dendritic cells

(22)

Impairment of gp91phox and p22phox expression Reduction of NAD(P)H oxidase activity and ROS level

HO-1 induction

(59)

(76)

(107)

(10, 11)

Reduction of proinflammatory cytokines (TNF-a, IL-1b, and MIP-1a) production Increase in the production of anti-inflammatory cytokine IL-10

HO-1 induction or CO treatment (+LPS)

Mouse

Macrophages

References

Effects

Treatment

Species

Cell type

Table 2 List of immune cells affected by HO-1 activity

260 Blancou et al.

Mast cells

Rat

Mouse

Human

Enhancement of FoxP3 expression Increase in IL-10 and TGF-b expression Increase in the immunosuppressive activity

HO-1 induction

Reduction of IgE-induced degranulation Downregulation of mast cell-dependent leukocyte adhesion

No modification of the immunosuppressive activity

HO-1 KO mouse

HO-1 induction

No correlation for FoxP3 and HO-1 expression in CD4+ T cells

(33)

(32)

(5)

(92)

(90)

No modification in FoxP3 expression Increase in Tregs proliferation No modification of the immunosuppressive activity

HO-1 induction

HO-1 KO mouse

(89)

Induction of HO-1 expression Loss of the immunosuppressive activity

FoxP3 gene transfection HO-1 inhibition

(96)

Suppression of T-lymphocyte proliferation Inhibition of caspase activity

(88)

(88)

Inhibition of CD8+ or CD4+ T-cells proliferation

HO-1 overexpression

(95)

(104)

(5)

References

Inhibition of ERK phosphorylation (CD4+) Inhibition of IL-2 production (CD4+) Inhibition of cell cycle progression (CD4+)

Abolishment of suppression of T-cell activation and transplantation tolerance

Impairment of Treg suppressive function

Effects

Anti-CD3/CD28 activation

CO treatment

Mouse

CD4+CD25+ Treg

HO-1 induction or CO treatment

Human

T-lymphocytes (other than CD4+ CD25+ Treg)

HO-1 inhibition

HO-1 inhibition

Mouse

Mouse

Treatment

Species

Myeloid-derived suppressor cells (MDSC)

Cell type

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Pae et al. showed that in contrast to human CD4CD25− T cells, human CD4CD25+ T cells (Treg) constitutively express HO-1 and that HO-1 expression increased in both cell types upon T-cell stimulation (88). However, even though a subsequent article from the same group showed that inhibition of endogenous HO-1 activity abrogated the suppressive function of Tregs on effector cells (89), these data have been recently revisited (90, 91), prompting the conclusion that HO-1 is not directly responsible for Treg impairment. One of the most convincing pieces of data has been obtained from HO-1 KO mutants where in vitro and in vivo regulatory T-cell function are normal (92). By contrast, DCs lacking HO-1 abrogated the immunosuppressive effect of Treg (5). Some authors have, however, raised the possibility that an HO-1-derived product, released from cells other than T cells, may play an immunosuppressive role by inhibiting T-cell activation or functions. As instance, HO-derived bilirubin down regulates costimulatory molecules on CD4 T cells, (93) and biliverdin interferes with T-cell signaling and suppresses Th1 function (94). Likewise, unspecific T-lymphocyte proliferation is strongly inhibited by CO delivered in gaseous form when compared with controls (95, 96). Altogether these data suggest that systemic HO-1 activation could lead to generalized T-cell dysfunction. 2.2. The Role of HO-1 in Macrophages Is Only Accountable for a Part of HO-1 Immunosuppressive Capabilities

In vitro studies have shown both reduced proinflammatory cytokine (TNF-a, IL-1b, MIP-1b, and IL-6) release and increased IL-10 expression in LPS stimulated in macrophages following HO-1 overexpression and/or CO exposure (10, 11, 97). Both treatment also lead to reduction of NAD(P)H oxidase activity and reactive-oxygen species level (52) (84). CO mediates these anti-inflammatory effects through a pathway involving the mitogen-activated protein kinases (MAPK p38) (10) and caveolin-1 (cav-1). The molecular mechanism of HO-1/CO inhibition of LPS-activated macrophage has recently been elucidated. In order to be fully active, upon LPS encounter, TLR4 has to be recruited to lipid rafts (i.e., plasma membranes of cells containing specialized microdomains). However, exogenous CO inhibits the trafficking of TLR4 toward the lipid raft after LPS stimulation in macrophages (76) by increasing the interaction of cav-1 (the major structural component of caveolae) with TLR4, which in turn redirects TLR4 toward caveolae and prevents its association with downstream transducing molecules (98). HO-1 also acts on transcriptional activity by binding to transcription factors such as the interferon regulatory factor-3 (IRF-3) leading to protein phosphorylation and nuclear migration resulting in abnormal function of TLR3/4 transduction pathways in macrophages (22).

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However, the in vivo relevance of macrophage loss of function still requires demonstration since LPS-stimulated peritoneal macrophages from HO-1 KO mice do not show global functional defects (99). A recent report using conditional HO-1 ablation in cells of the granulocyte/macrophage lineage has also shown that these cells are affected within TLR3 and partially TLR4 signaling pathway leading to very specific physiopathological defects (22). However, these mice do not recapitulate the HO-1 KO phenotype, originally described by Poss and Tonegawa suggesting that the inhibition of inflammation by naturally produced HO-1 is not mediated by the macrophage/granulocyte lineage. Interestingly, only a minor depletion of HO-1 expression in DC is achieved by this approach (100). 2.3. The Role of HO-1 in Dendritic Cells Could Be the Missing Link to Explain HO-1 Immunosuppressive Capabilities

The first evidence that HO-1 can influence DC function came from studies in humans and rats (42) followed by studies in mice (43). In these species, HO-1 overexpression by a chemical agent or gene transfer or CO treatment caused a decreased ratio of IL-12/IL-10 secretion and impaired stimulation capacities through inhibition of IRF-3 pathway (43). The use of a chemical agent to induce HO-1 is a matter of some debate since some of the effects produced by pharmacologic inducers can be seen in HO-1 KO mice (101). However, the same inhibition of DCs has also been observed by HO-1-end products such as bilirubin (93), HO-1 gene transfer, or CO (43). The physiological relevance of DCs immunomodulation by HO-1 through CO or bilirubin has been revealed in two induced models of autoimmune disease, namely EAE (35, 93) and diabetes (36, 43) where HO-1, bilirubin, or CO-treated DC can impair autoreactive T-cell function. Various authors have proposed at least four possible roles of HO-1/ CO-modified DC. The first is the inhibition of DCs presenting function resulting in the inhibition of helper T-cell and/or CD8 T-cell accumulation in organs (35, 36). The second possibility is the role of HO-1 on DCs in supporting the suppressive activity of Treg cells on effector T cells (5). This hypothesis is supported by the inhibition of heminic enzyme activities by HO-1-derived CO, as it is the case for NADPH oxydase (1) or indoleamine 2,3-dioxygenase (102). Treatment of DCs with HO-1 or with CO but not the other end products or heme depletion inhibited ROS, production of proinflammatory molecules such as IL-12, IL-6, TNF-a, and IFN type 1 while preserving IL-10 production (42, 43) and thus represent a favorable milieu for the emergence of Tregs. A third possibility is the inhibition of CD8 T-lymphocyte autoreactive function by HO-1/CO-modified DCs as demonstrated in an induced model of T1D (43). The last possibility recently emerged from a paper that reported on a natural CD8 regulatory T cell specific for HO-1 peptide (HO212) in humans (103). HO-1-specific CD8 Tregs isolated from PBL are very

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efficient at inhibiting lymphocyte functions and are found at a higher frequency in cancer patients as compared to healthy persons. However, the in vivo relevance of these inhibitory cells still has to be demonstrated. 2.4. Other Effect of HO-1 on Immune Cells

A recent paper showed that the inhibition of HO-1 in myeloidderived suppressor cells completely abolishes their suppressive function on T cells (104). The suppression of mast cell functions such as IgE secretion (33) or cytokine production (34) has also been recently demonstrated both in vitro and in vivo. Finally, immunomodulatory properties of MSCs on T-cell proliferation have been shown to act through HO-1 activity alone in humans and through both HO-1 and inducible-NO synthase (iNOS) activity in mice (62).

Acknowledgments This work was supported by funding from, La Région Pays de la Loire through the “Chaire d’excellence program” for AK and the IMBIO program, l’Agence de la Biomédecine, Ministère de la Recherche, Fondation CENTAURE, Fondation Progreffe, an ECOS France-Chile grant and Millennium Nucleus on Immunology and Immunotherapy from Chile (P04/030-F). LC is a CONICYT fellow. References 1. Ryter S W, Alam J, and Choi A M (2006) Heme oxygenase-1/carbon monoxide: from basic science to therapeutic applications. Physiol Rev 86: 583–650. 2. Poss KD and Tonegawa S (1997) Heme oxygenase 1 is required for mammalian iron reutilization. Proc Natl Acad Sci U S A 94: 10919–10924. 3. Yet S F, Perrella M A, Layne M D, et al (1999) Hypoxia induces severe right ventricular dilatation and infarction in heme oxygenase-1 null mice. J Clin Invest 103: R23–R29. 4. Kapturczak M H, Wasserfall C, Brusko T, et al (2004) Heme oxygenase-1 modulates early inflammatory responses: evidence from the heme oxygenase-1-deficient mouse. Am J Pathol 165: 1045–1053. 5. George J F, Braun A, Brusko T M, et al (2008) Suppression by CD4+CD25+ regulatory T cells is dependent on expression of heme oxygenase-1 in antigen-presenting cells. Am J Pathol 173: 154–160.

6. Chung S W, Liu X, Macias A A, et al (2008) Heme oxygenase-1-derived carbon monoxide enhances the host defense response to microbial sepsis in mice. J Clin Invest 118: 239–247. 7. Yachie A, Niida Y, Wada T, et al (1999) Oxidative stress causes enhanced endothelial cell injury in human heme oxygenase-1 deficiency. J Clin Invest 103: 129–135. 8. Nath K A, Vercellotti G M, Grande J P, et al (2001) Heme protein-induced chronic renal inflammation: suppressive effect of induced heme oxygenase-1. Kidney Int 59: 106–117. 9. Pittock S T, Norby S M, Grande J P, et al (2005) MCP-1 is up-regulated in unstressed and stressed HO-1 knockout mice: pathophysiologic correlates. Kidney Int 68: 611–622. 10. Otterbein L E, Bach F H, Alam J, et al (2000) Carbon monoxide has anti-inflammatory effects involving the mitogen-activated protein kinase pathway. Nat Med 6: 422–428.

Immunoregulatory Properties of Heme Oxygenase-1 11. Lee T S and Chau L Y (2002) Heme oxygenase-1 mediates the anti-inflammatory effect of interleukin-10 in mice. Nat Med 8: 240–246. 12. Rydkina E, Sahni A, Baggs R B, et al (2006) Infection of human endothelial cells with spotted fever group rickettsiae stimulates cyclooxygenase 2 expression and release of vasoactive prostaglandins. Infect Immun 74: 5067–5074. 13. Ren H, Leib S L, Ferriero D M, et al (2007) Induction of haem oxygenase-1 causes cortical non-haem iron increase in experimental pneumococcal meningitis: evidence that concomitant ferritin up-regulation prevents ironinduced oxidative damage. J Neurochem 100: 532–544. 14. Vareille M, Rannou F, Thelier N, et al (2008) Heme oxygenase-1 is a critical regulator of nitric oxide production in enterohemorrhagic Escherichia coli-infected human enterocytes. J Immunol 180: 5720–5726. 15. Shiloh M U, Manzanillo P, and Cox J S (2008) Mycobacterium tuberculosis senses host-derived carbon monoxide during macrophage infection. Cell Host Microbe 3: 323–330. 16. Kumar A, Deshane J S, Crossman D K, et al (2008) Heme oxygenase-1-derived carbon monoxide induces the Mycobacterium tuberculosis dormancy regulon. J Biol Chem 283: 18032–18039. 17. Zaki M H, Fujii S, Okamoto T, et al (2009) Cytoprotective function of heme oxygenase 1 induced by a nitrated cyclic nucleotide formed during murine salmonellosis. J Immunol 182: 3746–3756. 18. McAllister S C, Hansen S G, Ruhl R A, et al (2004) Kaposi sarcoma-associated herpesvirus (KSHV) induces heme oxygenase-1 expression and activity in KSHV-infected endothelial cells. Blood 103: 3465–3473. 19. Abdalla M Y, Ahmad I M, Spitz D R, et al (2005) Hepatitis C virus-core and non structural proteins lead to different effects on cellular antioxidant defenses. J Med Virol 76: 489–497. 20. Ghaziani T, Shan Y, Lambrecht R W, et al (2006) HCV proteins increase expression of heme oxygenase-1 (HO-1) and decrease expression of Bach1 in human hepatoma cells. J Hepatol 45: 5–12. 21. Seixas E, Gozzelino R, Chora A, et al (2009) Heme oxygenase-1 affords protection against noncerebral forms of severe malaria. Proc Natl Acad Sci U S A 106: 15837–15842. 22. Tzima S, Victoratos P, Kranidioti K, et al (2009) Myeloid heme oxygenase-1 regulates

23.

24.

25. 26.

27.

28.

29.

30.

31.

32.

33.

34.

265

innate immunity and autoimmunity by modulating IFN-beta production. J Exp Med 206: 1167–1179. Protzer U, Seyfried S, Quasdorff M, Sass G, et al (2007) Antiviral activity and hepatoprotection by heme oxygenase-1 in hepatitis B virus infection. Gastroenterology 133: 1156–1165. Epiphanio S, Mikolajczak S A, Goncalves L A, et al (2008) Heme oxygenase-1 is an antiinflammatory host factor that promotes murine plasmodium liver infection. Cell Host Microbe 3: 331–338. Sinnis P and Ernst J D (2008) CO-opting the host HO-1 pathway in tuberculosis and malaria. Cell Host Microbe 3: 277–279. Pamplona A, Ferreira A, Balla J, et al (2007) Heme oxygenase-1 and carbon monoxide suppress the pathogenesis of experimental cerebral malaria. Nat Med 13: 703–710. Willis D, Moore A R, and Willoughby D A (2000) Heme oxygenase isoform expression in cellular and antibody-mediated models of acute inflammation in the rat. J Pathol 190: 627–634. Kitada O, Kodama T, Kuribayashi K, et al (2001) Heme oxygenase-1 (HO-1) protein induction in a mouse model of asthma. Clin Exp Allergy 31: 1470–1477. Xia Z W, Zhong W W, Xu L Q, et al (2006) Hemeoxygenase-1-mediatedCD4+CD25high regulatory T cells suppress allergic airway inflammation. J Immunol 177: 5936–5945. Listopad J, Asadullah K, Sievers C, et al (2007) Heme oxygenase-1 inhibits T celldependent skin inflammation and differentiation and function of antigen-presenting cells. Exp Dermatol 16: 661–670. Kirino M, Kirino Y, Takeno M, et al (2008) Heme oxygenase 1 attenuates the development of atopic dermatitis-like lesions in mice: implications for human disease. J Allergy Clin Immunol 122: 290–297, 297, e291–298. Xia Z W, Xu L Q, Zhong W W, et al (2007) Heme oxygenase-1 attenuates ovalbumininduced airway inflammation by upregulation of foxp3 T-regulatory cells, interleukin-10, and membrane-bound transforming growth factor-1. Am J Pathol 171: 1904–1914. Takamiya R, Murakami M, Kajimura M, et al (2002) Stabilization of mast cells by heme oxygenase-1: an anti-inflammatory role. Am J Physiol Heart Circ Physiol 283: H861–H870. Yasui Y, Nakamura M, Onda T, et al (2007) Heme oxygenase-1 inhibits cytokine

266

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

Blancou et al. production by activated mast cells. Biochem Biophys Res Commun 354: 485–490. Chora A A, Fontoura P, Cunha A, et al (2007) Heme oxygenase-1 and carbon monoxide suppress autoimmune neuroinflammation. J Clin Invest 117: 438–447. Hu C M, Lin H H, Chiang M T, et al (2007) Systemic expression of heme oxygenase-1 ameliorates type 1 diabetes in NOD mice. Diabetes 56: 1240–1247. Li M, Peterson S, Husney D, Inaba M, et al (2007) Interdiction of the diabetic state in NOD mice by sustained induction of heme oxygenase: possible role of carbon monoxide and bilirubin. Antioxid Redox Signal 9: 855–863. Goldberg A, Parolini M, Chin B Y, et al (2007) Toll-like receptor 4 suppression leads to islet allograft survival. FASEB J 21: 2840–2848. Li Y, Li G, Dong W, Chen J, et al (2006) Transplantation of rat islets transduced with human heme oxygenase-1 gene using adenovirus vector. Pancreas 33: 280–286. Wagener F A, da Silva J L, Farley T, et al (1999) Differential effects of heme oxygenase isoforms on heme mediation of endothelial intracellular adhesion molecule 1 expression. J Pharmacol Exp Ther 291: 416–423. Wagener F A, Eggert A, Boerman O C, et al (2001) Heme is a potent inducer of inflammation in mice and is counteracted by heme oxygenase. Blood 98: 1802–1811. Chauveau C, Remy S, Royer P, et al (2005) Heme oxygenase-1 expression inhibits dendritic cell maturation and pro-inflammatory function but conserves IL-10 expression. Blood 106: 1694–1702. Remy S, Blancou P, Tesson L, et al (2009) Carbon monoxide inhibits TLR-induced dendritic cell immunogenicity. J Immunol 182: 1877–1884. Soares M P, Lin Y, Anrather J, et al (1998) Expression of heme oxygenase-1 can determine cardiac xenograft survival. Nat Med 4: 1073–1077. Sato K, Balla J, Otterbein L, Smith R, et al (2001) Carbon monoxide generated by heme oxygenase-1 suppresses the rejection of mouse-to-rat cardiac transplants. J Immunol 166: 4185–4194. Yamashita K, Ollinger R, McDaid J, et al (2006) Heme oxygenase-1 is essential for and promotes tolerance to transplanted organs. FASEB J 20: 776–778. Nakao A, Kimizuka K, Stolz D B, et al (2003) Protective effect of carbon monoxide

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

58.

inhalation for cold-preserved small intestinal grafts. Surgery 134: 285–292. Nakao A, Toyokawa H, Tsung A, et al (2006) Ex vivo application of carbon monoxide in University of Wisconsin solution to prevent intestinal cold ischemia/reperfusion injury. Am J Transplant 6: 2243–2255. Nakao A, Otterbein L E, Overhaus M, et al (2004) Biliverdin protects the functional integrity of a transplanted syngeneic small bowel. Gastroenterology 127: 595–606. Neto J S, Nakao A, Kimizuka K, et al (2004) Protection of transplant-induced renal ischemia-reperfusion injury with carbon monoxide. Am J Physiol Renal Physiol 287: F979–F989. Faleo G, Neto J S, Kohmoto J, et al (2008) Carbon monoxide ameliorates renal cold ischemia-reperfusion injury with an upregulation of vascular endothelial growth factor by activation of hypoxia-inducible factor. Transplantation 85: 1833–1840. Kaizu T, Ikeda A, Nakao A, et al (2008) Protection of transplant-induced hepatic ischemia/reperfusion injury with carbon monoxide via MEK/ERK1/2 pathway downregulation. Am J Physiol Gastrointest Liver Physiol 294: G236–G244. Nakao A, Neto J S, Kanno S, et al (2005) Protection against ischemia/reperfusion injury in cardiac and renal transplantation with carbon monoxide, biliverdin and both. Am J Transplant 5: 282–291. Bouche D, Chauveau C, Roussel J C, et al (2002) Inhibition of graft arteriosclerosis development in rat aortas following heme oxygenase-1 gene transfer. Transpl Immunol 9: 235–238. Clarke H M, Shrivastava S, Motterlini R, et al (2009) Donor HO-1 expression inhibits intimal hyperplasia in unmanipulated graft recipients: a potential role for CD8+ T-cell modulation by carbon monoxide. Transplantation 88: 653–661. Chauveau C, Bouchet D, Roussel J, et al (2002) Gene transfer of heme oxygenase-1 and carbon monoxide delivery inhibit chronic rejection. Am J Transplant 2: 581–592. Braudeau C, Bouchet D, Tesson L, et al (2004) Induction of long-term cardiac allograft survival by heme oxygenase-1 gene transfer. Gene Ther 11: 701–710. Araujo J A, Meng L, Tward A D, et al (2003) Systemic rather than local heme oxygenase-1 overexpression improves cardiac allograft outcomes in a new transgenic mouse. J Immunol 171: 1572–1580.

Immunoregulatory Properties of Heme Oxygenase-1 59. Kotsch K, Martins P N, Klemz R, et al (2007) Heme oxygenase-1 ameliorates ischemia/ reperfusion injury by targeting dendritic cell maturation and migration. Antioxid Redox Signal 9: 2049–2063. 60. Peche H, Trinite B, Martinet B, et al (2005) Prolongation of heart allograft survival by immature dendritic cells generated from recipient type bone marrow progenitors. Am J Transplant 5: 255–267. 61. Moreau A, Hill M, Thebault P, et al (2009) Tolerogenic dendritic cells actively inhibit T cells through heme oxygenase-1 in rodents and in nonhuman primates. FASEB J 23: 3070–3077. 62. Chabannes D, Hill M, Merieau E, et al (2007) A role for heme oxygenase-1 in the immunosuppressive effect of adult rat and human mesenchymal stem cells. Blood 110: 3691–3694. 63. Trigona W L, Porter C M, HorvathArcidiacono J A, et al (2007) Could hemeoxygenase-1 have a role in modulating the recipient immune response to embryonic stem cells? Antioxid Redox Signal 9: 751–756. 64. Becker T, Zu Vilsendorf A M, Terbish T, et al (2007) Induction of heme oxygenase-1 improves the survival of pancreas grafts by prevention of pancreatitis after transplantation. Transplantation 84: 1644–1655. 65. Wang H, Lee S S, Gao W, et al (2005) Donor treatment with carbon monoxide can yield islet allograft survival and tolerance. Diabetes 54: 1400–1406. 66. Wang H, Lee S S, Dell’Agnello C, et al (2006) Bilirubin can induce tolerance to islet allografts. Endocrinology 147: 762–768. 67. Lee S S, Gao W, Mazzola S, et al (2007) Heme oxygenase-1, carbon monoxide, and bilirubin induce tolerance in recipients toward islet allografts by modulating T regulatory cells. FASEB J 21: 3450–3457. 68. Lee D Y, Lee S, Nam J H, et al (2006) Minimization of immunosuppressive therapy after islet transplantation: combined action of heme oxygenase-1 and PEGylation to islet. Am J Transplant 6: 1820–1828. 69. Pulskens W P, Teske G J, Butter L M, et al (2008) Toll-like receptor-4 coordinates the innate immune response of the kidney to renal ischemia/reperfusion injury. PLoS One 3: e3596. 70. Kaczorowski D J, Nakao A, Vallabhaneni R, et al (2009) Mechanisms of toll-like receptor 4 (TLR4)-mediated inflammation after cold ischemia/reperfusion in the heart. Transplantation 87: 1455–1463.

267

71. Kaczorowski D J, Nakao A, Mollen K P, et al (2007) Toll-like receptor 4 mediates the early inflammatory response after cold ischemia/reperfusion. Transplantation 84: 1279–1287. 72. Zhai Y, Shen X D, O’Connell R, et al (2004) Cutting edge: TLR4 activation mediates liver ischemia/reperfusion inflammatory response via IFN regulatory factor 3-dependent MyD88-independent pathway. J Immunol 173: 7115–7119. 73. Methe H, Zimmer E, Grimm C, et al (2004) Evidence for a role of toll-like receptor 4 in development of chronic allograft rejection after cardiac transplantation. Transplantation 78: 1324–1331. 74. Tsuchihashi S, Zhai Y, Fondevila C, et al (2005) HO-1 upregulation suppresses type 1 IFN pathway in hepatic ischemia/reperfusion injury. Transplant Proc 37: 1677–1678. 75. Tsuchihashi S, Zhai Y, Bo Q, et al (2007) Heme oxygenase-1 mediated cytoprotection against liver ischemia and reperfusion injury: inhibition of type-1 interferon signaling. Transplantation 83: 1628–1634. 76. Nakahira K, Kim H P, Geng X H, et al (2006) Carbon monoxide differentially inhibits TLR signaling pathways by regulating ROSinduced trafficking of TLRs to lipid rafts. J Exp Med 203: 2377–2389. 77. Arbour N C, Lorenz E, Schutte B C, et al (2000) TLR4 mutations are associated with endotoxin hyporesponsiveness in humans. Nat Genet 25: 187–191. 78. Rallabhandi P, Bell J, Boukhvalova M S, et al (2006) Analysis of TLR4 polymorphic variants: new insights into TLR4/MD-2/CD14 stoichiometry, structure, and signaling. J Immunol 177: 322–332. 79. Kruger B, Krick S, Dhillon N, et al (2009) Donor toll-like receptor 4 contributes to ischemia and reperfusion injury following human kidney transplantation. Proc Natl Acad Sci U S A 106: 3390–3395. 80. Shen X D, Ke B, Zhai Y, et al (2005) Tolllike receptor and heme oxygenase-1 signaling in hepatic ischemia/reperfusion injury. Am J Transplant 5: 1793–1800. 81. Exner M, Bohmig G A, Schillinger M, et al (2004) Donor heme oxygenase-1 genotype is associated with renal allograft function. Transplantation 77: 538–542. 82. Ozaki K S, Marques G M, Nogueira E, et al (2008) Improved renal function after kidney transplantation is associated with heme oxygenase-1 polymorphism. Clin Transplant 22: 609–616.

268

Blancou et al.

83. Baan C, Peeters A, Lemos F, et al (2004) Fundamental role for HO-1 in the selfprotection of renal allografts. Am J Transplant 4: 811–818. 84. Courtney A E, McNamee P T, Middleton D, et al (2007) Association of functional heme oxygenase-1 gene promoter polymorphism with renal transplantation outcomes. Am J Transplant 7: 908–913. 85. Hribova P, Reinke P, Petrasek J, et al (2008) Heme oxygenase-1 polymorphisms and renal transplantation outcomes: balancing at the detection limit of allelic association studies. Am J Transplant 8: 1077–1078; author reply 1079. 86. Ullrich R, Exner M, Schillinger M, et al (2005) Microsatellite polymorphism in the heme oxygenase-1 gene promoter and cardiac allograft vasculopathy. J Heart Lung Transplant 24: 1600–1605. 87. Buis C I, van der Steege G, Visser D S, et al (2008) Heme oxygenase-1 genotype of the donor is associated with graft survival after liver transplantation. Am J Transplant 8: 377–385. 88. Pae H O, Oh G S, Choi B M, et al (2003) Differential expressions of heme oxygenase-1 gene in CD25− and CD25+ subsets of human CD4+ T cells. Biochem Biophys Res Commun 306: 701–705. 89. Choi B M, Pae H O, Jeong Y R, et al (2005) Critical role of heme oxygenase-1 in Foxp3mediated immune suppression. Biochem Biophys Res Commun 327: 1066–1071. 90. Biburger M, Theiner G, Schadle M, et al (2010) Pivotal advance: heme oxygenase 1 expression by human CD4+ T cells is not sufficient for their development of immunoregulatory capacity. J Leukoc Biol 87: 193–202. 91. Blancou P and Anegon I (2010) Heme oxygenase-1 and dendritic cells: what else? J Leuckoc Biol 87: 185–187. 92. Zelenay S, Chora A, Soares M P, et al (2007) Heme oxygenase-1 is not required for mouse regulatory T cell development and function. Int Immunol 19: 11–18. 93. Liu Y, Li P, Lu J, Xiong W, et al (2008) Bilirubin possesses powerful immunomodulatory activity and suppresses experimental autoimmune encephalomyelitis. J Immunol 181: 1887–1897. 94. Yamashita K, McDaid J, Ollinger R, et al (2004) Biliverdin, a natural product of heme catabolism, induces tolerance to cardiac allografts. FASEB J 18: 765–767. 95. Pae H O, Oh G S, Choi B M, et al (2004) Carbon monoxide produced by heme oxygenase-1 suppresses T cell proliferation via inhi-

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

107.

bition of IL-2 production. J Immunol 172: 4744–4751. Song R, Mahidhara R S, Zhou Z, et al (2004) Carbon monoxide inhibits T lymphocyte proliferation via caspase-dependent pathway. J Immunol 172: 1220–1226. Morse D, Pischke S E, Zhou Z, et al (2003) Suppression of inflammatory cytokine production by carbon monoxide involves the JNK pathway and AP-1. J Biol Chem 278: 36993–36998. Wang X M, Kim H P, Nakahira K, et al (2009) The heme oxygenase-1/carbon monoxide pathway suppresses TLR4 signaling by regulating the interaction of TLR4 with caveolin-1. J Immunol 182: 3809–3818. Poss KD and Tonegawa S (1997) Reduced stress defense in heme oxygenase 1-deficient cells. Proc Natl Acad Sci U S A 94: 10925–10930. Clausen B E, Burkhardt C, Reith W, et al (1999) Conditional gene targeting in macrophages and granulocytes using LysMcre mice. Transgenic Res 8: 265–277. Mashreghi M F, Klemz R, Knosalla I S, et al (2008) Inhibition of dendritic cell maturation and function is independent of heme oxygenase 1 but requires the activation of STAT3. J Immunol 180: 7919–7930. Hill M, Pereira V, Chauveau C, et al (2005) Heme oxygenase-1 inhibits rat and human breast cancer cell proliferation: mutual cross inhibition with indoleamine 2,3-dioxygenase. FASEB J 19: 1957–1968. Andersen M H, Sorensen R B, Brimnes M K, et al (2009) Identification of heme oxygenase1-specific regulatory CD8+ T cells in cancer patients. J Clin Invest 119: 2245–2256. De Wilde V, Van Rompaey N, Hill M, et al (2009) Endotoxin-induced myeloid-derived suppressor cells inhibit alloimmune responses via heme oxygenase-1. Am J Transplant 9: 2034–2047. Li M, Peterson S, Husney D, et al (2007) Long-lasting expression of HO-1 delays progression of type I diabetes in NOD mice. Cell Cycle 6: 567–571. Kohmoto J, Nakao A, Sugimoto R, et al (2008) Carbon monoxide-saturated preservation solution protects lung grafts from ischemia-reperfusion injury. J Thorac Cardiovasc Surg 136: 1067–1075. Taille C, El-Benna J, Lanone S, et al (2005) Mitochondrial respiratory chain and NAD(P)H oxidase are targets for the antiproliferative effect of carbon monoxide in human airway smooth muscle. J Biol Chem 280: 25350–25360.

Chapter 19 Indoleamine 2,3-Dioxygenase and Regulatory Function: Tryptophan Starvation and Beyond Ciriana Orabona and Ursula Grohmann Abstract Indoleamine 2,3-dioxygenase (IDO) is an ancestral enzyme that, initially confined to the regulation of tryptophan availability in local tissue microenvironments, is now considered to play a wider role that extends to homeostasis and plasticity of the immune system. Thus, IDO biology has many implications for many aspects of immunopathology, including viral infections, neoplasia, autoimmunity, and chronic inflammation. Its immunoregulatory effects are mainly mediated by dendritic cells (DCs) and involve not only tryptophan deprivation but also production of kynurenines that act on IDO− DCs – thus rendering an otherwise stimulatory DC capable of regulatory effects – as well as on T cells. As a result, IDO+ DCs mediate multiple effects on T lymphocytes, including inhibition of proliferation, apoptosis, and differentiation toward a regulatory phenotype. Key words: IDO, Tryptophan catabolism, Amino acid starvation, Kynurenines, Dendritic cells, Regulatory T cells

1. Introduction Over 40 years ago, Hayaishi and coworkers isolated a tryptophancatabolizing enzyme from small rabbit intestine (1), which, initially designated as D-tryptophan pyrrolase, is now known as indoleamine 2,3-dioxygenase (IDO; EC 1.13.11.52). Although soon recognized as a crucial molecule in the defense against infections and tumors (2), a very fruitful area of study was opened no sooner than 30 years later by Munn and Mellor (3), who provided the first evidence that IDO is involved in the maintenance of immune tolerance at the maternal–fetal interface. As a result, the number of IDO-related publications has increased almost tenfold since the early 90s, reaching the figure of almost five hundreds in 2008. Conserved through the last 600 million years Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_19, © Springer Science+Business Media, LLC 2011

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of evolution, IDO is presently recognized as an authentic regulator of the immune system, with implications for many aspects of immunopathology.

2. Biochemistry of IDO IDO is a heme-containing monomeric and cytoplasmic enzyme of 42–45 kDa that catalyzes the first and the rate-limiting step of the kynurenine pathway, which produces a series of metabolites collectively known as kynurenines (Kyns; Fig. 1). The definition of the crystal structure of human IDO has revealed a folding into a catalytic large C-terminal domain (containing the heme), a noncatalytic small N-terminal domain, and a long loop connecting the two domains (4). Apart from covering the top of the heme-binding site (4), the role of the noncatalytic small domain is still unknown. IDO transforms L-tryptophan (L-TRP), the least

TRYPTOPHAN STARVATION

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IDO N-Formylkynurenine KYNA Kynurenine formamidase Kynurenine aminotransferase

KYN Kynureninase Kynurenine 3-monooxygenase

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AA 3-HK Kynureninase

3-HAA

3-Hydroxyanthranilate 3.4-dioxygenase

QUIN

Fig. 1. Schematic representation of tryptophan catabolism along the kynurenine pathway. Indoleamine-2,3-dioxygenase (IDO) induces both tryptophan starvation and kynurenine production. L-TRP L-tryptophan; KYN L-kynurenine; 3-HK 3-hydroxykynurenine; KYNA kynurenic acid; AA anthranilic acid; 3-HAA 3-hydroxyanthranilic acid; QUIN quinolinic acid.

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available essential amino acid (EAA), into N-formylkynurenine, a product that is rapidly converted by formamidase to L-kynurenine (KYN), which in turn can either enter the bloodstream or be further metabolized to downstream Kyns. IDO is not selective for L-TRP and in fact other indoleamine derivatives, i.e., D-TRP, tryptamine, 5-hydroxytryptophan, and serotonin, can act as substrates for the enzyme. Expression of the IDO protein does not always correlate with enzyme activity, which requires incorporation of the heme prosthetic group and the presence of superoxide anion in addition to still undefined posttranslational modifications (5). No longer considered as a cosubstrate by most authors, superoxide is now regarded as a cofactor, a substrate precursor or, more recently, an activating agent for IDO, as it controls the shift between the inactive and active states of the enzyme (6, 7). 2.2. Measurements of IDO Activity

Since IDO is the first and rate-limiting enzyme in the pathway (Fig. 1), measurements of its activity are mainly based on the determination of KYN, the first stable metabolite. Two methods are mainly used, namely high pressure liquid chromatography (HPLC) and spectrophotometric analysis. In the latter, KYN can be reacted with Ehrlich’s reagent (p-dimethylaminobenzaldehyde in a strong acid medium), producing a yellow compound that absorbs at 490 nm. This method is simple, fast, and easily automated. However, the reagent can also react with other molecules containing aldehyde or ketone groups. In particular, the hydroxyl group of 3-hydroxyanthranilic acid (3-HAA) and 3-hydroxykynurenine (3-HK) can tautomerize to ketone in acidic medium, thereby becoming reactive to the reagent (8). HPLC is more specific and is thus widely used. This method requires a C18 column, a mobile phase with a variable composition but constant pH (3, 4) to ensure KYN polarity, and an UV detector at a wavelength of 360 nm. By HPLC, L-TRP can be measured in the same analysis, an advantage that is particularly useful in the determination of IDO activity in biological fluids – blood sera and urine – in which variability could be extremely high. In order to increase the signalto-noise ratio, samples can be subjected to deproteinization by means of trichloroacetic acid or methanol, before HPLC analysis. In addition, some authors incubate samples at 50°C for 30 min to hydrolyze completely the intermediate N-formylkynurenine to KYN. KYN determination in cell culture supernatants implies the incubation of cells with an excess of L-TRP for 2–8 h and the addition of appropriate stimuli (see below). More sensitive than HPLC, however, though less affordable, mass spectrometry can be used in positive ionization mode, which is performed as a single step or after HPLC. This approach is particularly indicated for downstream Kyns, i.e., 3-HAA and quinolinic acid (QUIN), which are normally produced at lower levels than KYN.

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2.3. Regulation of IDO Expression

IDO is coded by the IDO1 gene (Ido1, previously Indo, in the mouse) that contains ten exons and spans 15 kb in chromosome 8, 8p12-p11 (8 A2 in the mouse). Although basally expressed at low levels in several tissues (i.e., lung, gut, epididymis, brain, and thymus), high-rate transcription of IDO1 is stringently controlled and confined to a limited range of cell types (5). Highest levels of expression can be found in professional antigen-presenting cells (APCs) such as dendritic cells (DCs), mainly in response to type I (IFN-a and IFN-b) and, to a greater extent, type II (IFN-g) IFNs. The IFN-dependent transcription of IDO1 involves an IFN-gactivated site (GAS) and two IFN-stimulated response elements (ISRE) that respond to signal transducer and activator of transcription 1 (STAT1) and IFN-regulatory factor-1 (IRF-1), respectively. A novel mode of IDO induction, involving phosphatidylinositol 3-kinase (PI3K)/Akt and noncanonical NF-kB, has been described in DCs in response to TGF-b (9), a suppressive cytokine that can be produced by and can act on both DCs and T cells. Since IDO in turn produces soluble and immunoregulatory Kyns, the TGF-b/IDO axis may represent one mechanism underlying the occurrence of infectious tolerance (10), i.e., a tolerant state that can be spread from one cell population to another. Soluble inducers of IDO also include hormones such as human chorionic gonadotropin (11) and estrogens (12), supporting the hypothesis that remission from autoimmunity observed during pregnancy could be related to L-TRP catabolism, and the arachidonate metabolite prostaglandin E2, which may mediate early events during induction of immune unresponsiveness in cancer (13). In addition, ligands of Toll-like receptor 9, including microbial CpG-rich oligodeoxynucleotides and endogenous thymosin a1, can also induce Ido1 (14). Besides soluble molecules, IDO expression is controlled by membrane-anchored coreceptors involved in the cross talk between DCs and T cells. Upon engagement of B7 molecules (CD80 and CD86) on DCs by the inhibitory receptor CTLA-4 expressed on activated or regulatory T (Treg) cells, IDO is induced in DCs in an IFN-g-mediated fashion (15, 16). However, IDO expression also increases when B7 coreceptors are bound by the stimulatory receptor CD28, providing that the gene coding for suppressor of cytokine signaling 3 (SOCS3) is silenced in DCs (17, 18). In the subset of plasmacytoid DCs (pDCs), Ido1 transcription can be activated upon engagement of glucocorticoid-induced TNF-related ligand (GITR-L) by its coreceptor GITR (19), also highly expressed by Treg cells. GITR/GITRL-mediated induction of IDO can be triggered by glucocorticoids, is mediated by IFN-a, and requires the activation of the noncanonical pathway of NF-kB. Ido1 negative modulators include Bin1, a BAR-adapterencoding gene, whose downregulation in several tumors clearly associates with IDO upregulation (20), and Tyrobp, coding for

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the transmembrane adapter DAP12 that represses Ido1 transcription by an unknown mechanism (21, 22). Once expressed, IDO protein can be still subjected to negative regulation. In fact, recent evidence indicates that IDO undergoes regulatory proteolysis in DCs programmed to direct immunity. Two putative immunoreceptor tyrosine-based inhibitory (ITIM) motifs present in IDO protein sequence can be bound by SOCS3, which, induced in DCs by proinflammatory IL-6, drives proteasomal degradation of the enzyme in an ubiquitin-dependent fashion (23). 2.4. The Family of IDO-Related Genes

3. Mechanisms of IDO-Mediated Immune Suppression

In mammalians, two additional enzymes besides IDO catalyze the rate-limiting step along the kynurenine pathway. These are tryptophan 2,3-dioxygenase (TDO), coded by TDO2 in humans and Tdo2 in mice, and indoleamine 2,3-dioxygenase-2 (IDO2, previously INDOL1), coded by IDO2 in humans and Ido2 in mice. While IDO and IDO2 proteins are structurally quite similar (43%) (24), TDO displays only 10% homology with IDO, contains the large catalytic but not the small domain, and has a homotetrameric structure (25). Furthermore, TDO is mostly confined to the liver, where it metabolizes exogenous L-TRP, and is not inducible by proinflammatory signals. IDO2 is a paralogue of IDO1 and is considered to be a protoIDO1 gene, since it shows higher similarity to IDOs that are expressed in low vertebrates (24). Compared with IDO, IDO2 is characterized by a very low substrate affinity and enzyme activity, suggesting that L-TRP might not be the true substrate (24). In addition, while the L isomer of 1-methyltryptophan (1MT), the gold standard in IDO inhibition, is more selective for IDO blockade (26), the D isomer appears to inhibit only IDO2 activity (27). Therefore, if we also consider the peculiar expression pattern of IDO2 – kidney, liver, and reproductive system, with DCs still being discussed in terms of enzyme activity (26, 27) – it appears that IDO represents the most important regulator of immune responses among L-TRP-catabolizing enzymes. Tools such as Ido1−/− mice (5), which retain Ido2 expression (24), could be pivotal in ascertaining the true biological function of IDO2. The fact that these mice do not show any grossly aberrant immunological phenotype may indeed suggest that IDO2 and/or other redundant mechanisms (see below) may compensate for IDO activity.

IDO activity contributes not only to maternal tolerance in pregnancy, but also controls allograft rejection (15) and protects against autoimmunity (28, 29), inflammatory pathology – colitis (30),

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rheumatoid arthritis (31), and granulomatous diseases (7, 32) – and allergy (19, 33). In addition, IDO contributes to host’s immune unresponsiveness to neoplasia (20, 34–36) and in viral infection (37). The wide spectrum of physiopathologic conditions in which IDO is at work suggests that multiple mechanisms are used by this effector system to downregulate T cell and inflammatory responses. 3.1. Starvation of Tryptophan, an Essential Amino Acid

Metabolism-targeting drugs are emerging as a new class of immunomodulators. While fatty acid metabolism generates long-lived immune memory cells (38), starvation of EAAs in APCs seems to be mandatory in the control of T cell differentiation (39). During shortage of EAAs, a starvation response program protective for self-structures is initiated in which the expression of most proteins is blocked with the exception of a selected set of genes (39). This integrated stress response involves the general control nonderepressible-2 kinase (GCN2) that, after sensing uncharged tRNAs, inactivates the eukaryotic translation initiation factor 2a (eIF2a) and shuts down the expression of several proteins, including immunostimulatory IL-6 (40). The activation of the GCN2 pathway can lead to profound anergy in T cells (41) and can block the conversion of Treg into Th17-like cells (40). However, recent evidence indicates that, in addition to GCN2, the mammalian target of rapamycin (mTOR), a downstream effector of the PI3Kmediated pathway, also plays a crucial role in the EAA starvation program. In this case, a reduction in EAA inhibits mTOR signaling in T cells – similarly to rapamycin – and induces de novo expression of FOXP3 (the cell lineage factor of Treg cells) in synergism with TGF-b (42). Although the enzymatic catabolism of other EAAs, i.e., L-arginine and L-histidine, can also inhibit mTOR, L-TRP catabolism mediated by IDO may have additional specialized immunomodulatory properties (42). In fact, IDO not only activates GCN2 (41) and inhibits mTOR (42), but also generates a variety of Kyns (Fig. 2) that are collectively endowed with an array of immunoregulatory effects.

3.2. Effects of Immunoregulatory Kynurenines

In the kynurenine pathway (Fig. 1), IDO activity produces KYN, which can be transformed by downstream enzymes – also inducible by IFN-g – into 3-HK, 3-HAA, and QUIN. Kynurenic acid (KYNA), a neuroprotective molecule detectable in the brain, is not apparently produced by DCs (Fallarino et al. unpublished observations). Kyns produced by DCs can exert a series of effects on cells of the immune system. In vitro, 3-HAA and QUIN induce selective apoptosis of murine thymocytes and Th1 but not Th2 cells; KYN, 3-HK, and 3-HAA inhibit the proliferation of T lymphocytes, whereas KYN also blocks NK cell proliferation (43). 3-HAA has been shown to induce T cell apoptosis via inhibition of PDK1 kinase and canonical NF-kB, an effect involved in the

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INFECTIOUS TOLERANCE eIF2a

PI3K/mTOR

? GCN2

TRYPTOPHAN STARVATION

IDO

KYNURENINE PRODUCTION

Fig. 2. Schematic diagram of IDO contribution to infectious tolerance. IDO activity in dendritic cells (DC) leads to tryptophan starvation that, in T cells, inhibits PI3K/mTOR signaling and, in combination with kynurenine production, activates GCN2 and thus inhibits eIF2a. Kynurenines may also exert direct effects on T cells via still unknown targets. The combination of multiple mechanisms activated by IDO may be involved in the spread and maintenance of infectious tolerance.

suppression of experimental asthma (44). However, each L-TRP catabolite could act directly on target cells or be further transformed into active molecule/s, if DCs are present in the microenvironment. As matter of fact, IDO− DCs can be rendered tolerogenic by the addition of exogenous KYN and IFN-g (45). The effect implies the capacity of DCs to take up extracellular KYN by means of the large neutral amino acid transporter – the same carrier for L-TRP entrance into the cell – and to further transform it by enzymes downstream IDO. Thus, these data demonstrate that L-TRP consumption may not always be an absolute requirement for the kynurenine pathway to exert suppressive properties and that few IDO-competent DCs may extend their immunosuppressive potential to other bystander IDO-noncompetent cells via L-TRP catabolism. In vivo, administration of 3-HAA or tranilast, an orally active synthetic drug structurally similar to L-TRP catabolites, skews the T cell response from Th1 to Th2 in mice with experimental autoimmune encephalomyelitis (EAE) (29). In an experimental model of chronic granulomatous disease (CGD), coadministration of KYN and recombinant IFN-g reverses hyperinflammation and enables the emergence of Treg cells, but only if the L-TRP catabolite can be further transformed in vivo (7). Therefore, although the molecular targets of Kyns in T cells are still unknown (Fig. 2), these molecules, either singly or in combination with other Kyns, clearly represent important regulator of the immune system.

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3.3. Multiple Mechanisms May Underlie Long-Lived and Potent IDO Immunosuppression

4. IDO and Therapeutic Applications 4.1. Neoplasia and Viral Infections: How to Dampen IDO Overexpression

IDO can deplete an EAA in local tissue microenvironments, but can also generate immunoregulatory Kyns. Since neither mechanism alone can explain the diversity (3, 7, 28, 31), durability (15), and antigen specificity (35) of the IDO-mediated suppressive effect, a fine combination of multiple processes may be necessary for long-lived and efficient immunosuppression. Notably, both L-TRP starvation and presence of all Kyns at physiologic concentrations are absolutely required for the downregulation of z chain in CD8+ T cells and for the GCN2-dependent conversion of naïve CD4+CD25− T cells into CD4+CD25+FOXP3+ Treg cells, capable of suppressing the occurrence of fulminant autoimmune diabetes in mice (46). For their suppressive action, these cells require CTLA-4 and IL-10 in vitro and IDO activity in vivo, thus suggesting that IDO may represent the functional bridge between DCs and Treg cells in the spread and maintenance of infectious tolerance (10, 47). Although as yet uninvestigated, we cannot presently exclude that the in vivo efficacy of Kyns, particularly when administered in conditions of IDO deficiency (7), may still require the activation of the starvation program by deprivation of EAA other than TRP or of alternative nutrients.

A bulk of data points to IDO as one of the main causes of immune unresponsiveness in neoplasia and viral infection (20, 34–37). In tumor-bearing hosts, IDO can be highly expressed not only by DCs within tumor-infiltrating lymph nodes, but also by neoplastic cells themselves. IDO expression in DCs has been correlated with suppression of T cell responses and a poor prognosis (5). In tumor cells, downregulation of Bin1 is associated with Ido1 upregulation. Thus, blockade of IDO expression and/or activity may represent a new effective strategy in cancer (20, 27, 34). Although I-1MT is a better inhibitor of IDO in vitro than the D isomer (26), in tumor-bearing animals the latter seems to be more effective (27), suggesting that in neoplasia, IDO2 may also contribute to immunosuppression. These data have led to direct testing of D-1MT in clinical trials (NCT00567931). However, since the exact mode of action of D-1MT is still unknown, perplexities have arisen (48). More recently, a novel approach for dampening IDO expression in a more specific fashion has been proposed. Skin administration of small interfering RNA specific for Ido1 significantly downregulates IDO expression in DCs, an effect that prolongs survival in tumor-bearing mice (49). In principle, overexpression of IDO could not only be controlled by direct targeting of the enzyme transcript/activity but also by blocking the main signaling pathways inducing L-TRP catabolism. However, IDO staining in macrophages and/or tumor cells shows no

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definite pattern of variation in melanoma patients undergoing therapy with tremelimumab, a monoclonal antibody capable to block CTLA-4 (50). In contrast, effective antiretroviral therapy coadministered with the antiCTLA-4 ipilimumab decreases the expression of IDO in lymph nodes (51). Thus, although not extensively investigated yet, indirect targeting of IDO could also be exploited in specific pathological conditions. 4.2. Autoimmunity and Chronic Inflammation: How to Overcome IDO Deficiency

IDO deficiency has been revealed in some experimental disease models, including experimental autoimmune diabetes (28) and CGD (7). In autoimmunity, the deficiency is related to a signaling defect, mostly in pancreatic DCs and consisting of an imbalanced canonical vs. noncanonical NF-kB activation (52, 53). In CGD, IDO is normally expressed but is not functionally active, because the IDO cofactor superoxide anion is completely lacking (7). Although experiments in our laboratory are still ongoing in autoimmune diabetes, complete cure of lethal pulmonary aspergillosis in CGD mice can be achieved by replacement therapy with KYN and recombinant IFN-g (7). IDO deficiency may not represent the only condition requiring IDO-based therapies. In EAE, in which IDO deficiency does not seem to be evident, administration of 3-HAA reverses paralysis (29). In addition, up-regulation of IDO activity via glucocorticoids protects mice in a model of allergic airway inflammation, in which IDO is basally active (19). A completely new approach in immunosuppressive therapy could be represented by the use of immunoproteasome inhibitors (54) that, already demonstrated to be effective in experimental arthritis, may act via blockade of IDO-regulatory proteolysis (23).

5. Conclusions Although our appreciation of both the complexity and potential for therapeutic intervention of the IDO pathway has expanded enormously in recent years, key unanswered questions still remain. These mainly relate to the precise mechanisms underlying the long-lived, potent immunosuppressive IDO effects, particularly those mediated by Kyns alone, and the relationship between IDO and IDO2 in neoplasia. Clarifying these aspects could ultimately offer considerable promise in facilitating our understanding of the general mechanisms of tolerance.

Acknowledgments The Authors would like to thank Gianluca Andrielli for digital art.

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References 1. Yamamoto, S., and Hayaishi, O. (1967) Tryptophan pyrrolase of rabbit intestine. D- and L-tryptophan-cleaving enzyme or enzymes. J. Biol. Chem. 242, 5260–5266. 2. Taylor, M. W., and Feng, G. S. (1991) Relationship between interferon-gamma, indoleamine 2,3-dioxygenase, and tryptophan catabolism. FASEB J. 5, 2516–2522. 3. Munn, D. H., Zhou, M., Attwood, J. T., et al. (1998) Prevention of allogeneic fetal rejection by tryptophan catabolism. Science 281, 1191–1193. 4. Sugimoto, H., Oda, S., Otsuki, T., et al. (2006) Crystal structure of human indoleamine 2,3-dioxygenase: catalytic mechanism of O2 incorporation by a heme-containing dioxygenase. Proc. Natl. Acad. Sci. U.S.A. 103, 2611–2616. 5. Mellor, A. L., and Munn, D. H. (2004) IDO expression by dendritic cells: tolerance and tryptophan catabolism. Nat. Rev. Immunol. 4, 762–774. 6. Macchiarulo, A., Nuti, R., Bellocchi, D., et al. (2007) Molecular docking and spatial coarse graining simulations as tools to investigate substrate recognition, enhancer binding and conformational transitions in indoleamine2,3-dioxygenase (IDO). Biochim. Biophys. Acta 1774, 1058–1068. 7. Romani, L., Fallarino, F., De Luca, A., et al. (2008) Defective tryptophan catabolism underlies inflammation in mouse chronic granulomatous disease. Nature 451, 211–215. 8. Alegre, E., Lopez, A. S., and Gonzalez, A. (2005) Tryptophan metabolites interfere with the Ehrlich reaction used for the measurement of kynurenine. Anal. Biochem. 339, 188–189. 9. Belladonna, M. L., Volpi, C., Bianchi, R., et al. (2008) Cutting edge: autocrine TGF-b sustains default tolerogenesis by IDOcompetent dendritic cells. J. Immunol. 181, 5194–5198. 10. Belladonna, M. L., Orabona, C., Grohmann, U., and Puccetti, P. (2009) TGF-b and kynurenines as the key to infectious tolerance. Trends Mol. Med. 15, 41–49. 11. Ueno, A., Cho, S., Cheng, L., et al. (2007) Transient upregulation of indoleamine 2,3-dioxygenase in dendritic cells by human chorionic gonadotropin downregulates autoimmune diabetes. Diabetes 56, 1686–1693. 12. Zhu, W. H., Lu, C. Z., Huang, Y. M., et al. (2007) A putative mechanism on remission of multiple sclerosis during pregnancy: estrogen-

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

induced indoleamine 2,3-dioxygenase by dendritic cells. Mult. Scler. 13, 33–40. von Bergwelt-Baildon, M. S., Popov, A., Saric, T., et al. (2006) CD25 and indoleamine 2,3-dioxygenase are up-regulated by prostaglandin E2 and expressed by tumor-associated dendritic cells in vivo: additional mechanisms of T-cell inhibition. Blood 108, 228–237. Fallarino, F., and Puccetti, P. (2006) Toll-like receptor 9-mediated induction of the immunosuppressive pathway of tryptophan catabolism. Eur. J. Immunol. 36, 8–11. Grohmann, U., Orabona, C., Fallarino, F., et al. (2002) CTLA-4-Ig regulates tryptophan catabolism in vivo. Nat. Immunol. 3, 1097–1101. Fallarino, F., Grohmann, U., Hwang, K. W., et al. (2003) Modulation of tryptophan catabolism by regulatory T cells. Nat. Immunol. 4, 1206–1212. Orabona, C., Grohmann, U., Belladonna, M. L., et al. (2004) CD28 induces immunostimulatory signals in dendritic cells via CD80 and CD86. Nat. Immunol. 5, 1134–1142. Orabona, C., Belladonna, M. L., Vacca, C., et al. (2005) Cutting edge: silencing suppressor of cytokine signaling 3 expression in dendritic cells turns CD28-Ig from immune adjuvant to suppressant. J. Immunol. 174, 6582–6586. Grohmann, U., Volpi, C., Fallarino, F., et al. (2007) Reverse signaling through GITR ligand enables dexamethasone to activate IDO in allergy. Nat. Med. 13, 579–586. Muller, A. J., DuHadaway, J. B., Donover, P. S., et al. (2005) Inhibition of indoleamine 2,3-dioxygenase, an immunoregulatory target of the cancer suppression gene Bin1, potentiates cancer chemotherapy. Nat. Med. 11, 312–319. Orabona, C., Tomasello, E., Fallarino, F., et al. (2005) Enhanced tryptophan catabolism in the absence of the molecular adapter DAP12. Eur. J. Immunol. 35, 3111–3118. Orabona, C., Puccetti, P., Vacca, C., et al. (2006) Toward the identification of a tolerogenic signature in IDO-competent dendritic cells. Blood 107, 2846–2854. Orabona, C., Pallotta, M. T., Volpi, C., et al. (2008) SOCS3 drives proteasomal degradation of indoleamine 2,3-dioxygenase (IDO) and antagonizes IDO-dependent tolerogenesis. Proc. Natl. Acad. Sci. U.S.A. 105, 20828–20833. Ball, H. J., Yuasa, H. J., Austin, C. J., et al. (2009) Indoleamine 2,3-dioxygenase-2; a

IDO and Immune Regulation

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

new enzyme in the kynurenine pathway. Int. J. Biochem. Cell Biol. 41, 467–471. Zhang, Y., Kang, S. A., Mukherjee, T., et al. (2007) Crystal structure and mechanism of tryptophan 2,3-dioxygenase, a heme enzyme involved in tryptophan catabolism and in quinolinate biosynthesis. Biochemistry 46, 145–155. Löb, S., Königsrainer, A., Schafer, R., et al. (2008) Levo- but not dextro-1-methyl tryptophan abrogates the IDO activity of human dendritic cells. Blood 111, 2152–2154. Metz, R., Duhadaway, J. B., Kamasani, U., et al. (2007) Novel tryptophan catabolic enzyme IDO2 is the preferred biochemical target of the antitumor indoleamine 2,3-dioxygenase inhibitory compound D-1-methyl-tryptophan. Cancer Res. 67, 7082–7087. Grohmann, U., Fallarino, F., Bianchi, R., et al. (2003) A defect in tryptophan catabolism impairs tolerance in nonobese diabetic mice. J. Exp. Med. 198, 153–160. Platten, M., Ho, P. P., Youssef, S., et al. (2005) Treatment of autoimmune neuroinflammation with a synthetic tryptophan metabolite. Science 310, 850–855. Gurtner, G. J., Newberry, R. D., Schloemann, S. R., et al. (2003) Inhibition of indoleamine 2,3-dioxygenase augments trinitrobenzene sulfonic acid colitis in mice. Gastroenterology 125, 1762–1773. Seo, S. K., Choi, J. H., Kim, Y. H., et al. (2004) 4-1BB-mediated immunotherapy of rheumatoid arthritis. Nat. Med. 10, 1088–1094. Popov, A., Abdullah, Z., Wickenhauser, C., et al. (2006) Indoleamine 2,3-dioxygenaseexpressing dendritic cells form suppurative granulomas following Listeria monocytogenes infection. J. Clin. Invest. 116, 3160–3170. Hayashi, T., Beck, L., Rossetto, C., et al. (2004) Inhibition of experimental asthma by indoleamine 2,3-dioxygenase. J. Clin. Invest. 114, 270–279. Uyttenhove, C., Pilotte, L., Theate, I., et al. (2003) Evidence for a tumoral immune resistance mechanism based on tryptophan degradation by indoleamine 2,3-dioxygenase. Nat. Med. 9, 1269–1274. Munn, D. H., Sharma, M. D., Hou, D., et al. (2004) Expression of indoleamine 2,3-dioxygenase by plasmacytoid dendritic cells in tumor-draining lymph nodes. J. Clin. Invest. 114, 280–290. Curti, A., Trabanelli, S., Salvestrini, V., et al. (2009) The role of indoleamine 2,3-dioxygenase in the induction of immune tolerance: focus on hematology. Blood 113, 2394–2401.

279

37. Boasso, A., Herbeuval, J. P., Hardy, A. W., et al. (2007) HIV inhibits CD4+ T-cell proliferation by inducing indoleamine 2,3-dioxygenase in plasmacytoid dendritic cells. Blood 109, 3351–3359. 38. Prlic, M., and Bevan, M. J. (2009) Immunology: a metabolic switch to memory. Nature 460, 41–42. 39. Blander, J. M., and Amsen, D. (2009) Immunology. Amino acid addiction. Science 324, 1282–1283. 40. Sharma, M. D., Hou, D. Y., Liu, Y., et al. (2009) Indoleamine 2,3-dioxygenase controls conversion of Foxp3+ Tregs to TH17-like cells in tumor-draining lymph nodes. Blood 113, 6102–6111. 41. Munn, D. H., Sharma, M. D., Baban, B., et al. (2005) GCN2 kinase in T cells mediates proliferative arrest and anergy induction in response to indoleamine 2,3-dioxygenase. Immunity 22, 633–642. 42. Cobbold, S. P., Adams, E., Farquhar, C. A., et al. (2009) Infectious tolerance via the consumption of essential amino acids and mTOR signaling. Proc. Natl. Acad. Sci. U.S.A. 106, 12055–12060. 43. Grohmann, U., Fallarino, F., and Puccetti, P. (2003) Tolerance, DCs and tryptophan: much ado about IDO. Trends Immunol. 24, 242–248. 44. Hayashi, T., Mo, J. H., Gong, X., et al. (2007) 3-Hydroxyanthranilic acid inhibits PDK1 activation and suppresses experimental asthma by inducing T cell apoptosis. Proc. Natl. Acad. Sci. U.S.A. 104, 18619–18624. 45. Belladonna, M. L., Grohmann, U., Guidetti, P., et al. (2006) Kynurenine pathway enzymes in dendritic cells initiate tolerogenesis in the absence of functional IDO. J. Immunol. 177, 130–137. 46. Fallarino, F., Grohmann, U., You, S., et al. (2006) The combined effects of tryptophan starvation and tryptophan catabolites downregulate T cell receptor z-chain and induce a regulatory phenotype in naive T cells. J. Immunol. 176, 6752–6761. 47. Puccetti, P., and Grohmann, U. (2007) IDO and regulatory T cells: a role for reverse signalling and non-canonical NF-kB activation. Nat. Rev. Immunol. 7, 817–823. 48. Löb, S., Königsrainer, A., Rammensee, H. G., et al. (2009) Inhibitors of indoleamine-2,3dioxygenase for cancer therapy: can we see the wood for the trees? Nat. Rev. Cancer 9, 445–452. 49. Yen, M. C., Lin, C. C., Chen, Y. L., et al. (2009) A novel cancer therapy by skin delivery of indoleamine 2,3-dioxygenase. Clin. Cancer Res. 15, 641–649.

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Orabona and Grohmann

50. Ménard, C., Ghiringhelli, F., Roux, S., et al. (2008) Ctla-4 blockade confers lymphocyte resistance to regulatory T-cells in advanced melanoma: surrogate marker of efficacy of tremelimumab? Clin. Cancer Res. 14, 5242–5249. 51. Hryniewicz, A., Boasso, A., Edghill-Smith, Y., et al. (2006) CTLA-4 blockade decreases TGF-b, IDO, and viral RNA expression in tissues of SIVmac251-infected macaques. Blood 108, 3834–3842. 52. Fallarino, F., Bianchi, R., Orabona, C., et al. (2004) CTLA-4-Ig activates forkhead

transcription factors and protects dendritic cells from oxidative stress in nonobese diabetic mice. J. Exp. Med. 200, 1051–1062. 53. Fallarino, F., Volpi, C., Zelante, T., et al. (2009) IDO mediates TLR9-driven protection from experimental autoimmune diabetes. J. Immunol. 183, 6303–6312. 54. Muchamuel, T., Basler, M., Aujay, M. A.,et al. (2009) A selective inhibitor of the immunoproteasome subunit LMP7 blocks cytokine production and attenuates progression of experimental arthritis. Nat. Med. 15, 781–787.

Chapter 20 Regulatory T Cell Enrichment by IFN-g Conditioning Gang Feng, Kathryn J. Wood, and Andrew Bushell Abstract IFN-g was originally characterized as a proinflammatory cytokine with T helper type 1 inducing activity, but it is now clear that it also has important immunoregulatory functions. Regulatory T cells play an important role in models of autoimmunity, GVHD, and transplantation, and offer potential as a cellular therapy. In rodent models, in vivo-generated CD25+CD4+ T cells can prevent allograft rejection, but therapeutic exploitation of Treg will more likely depend on protocols that allow the generation or selection of Treg ex vivo. The experiments described in this chapter will show that alloantigen-reactive Treg can be generated/ expanded ex vivo using IFN-g, a cytokine more usually associated with allograft rejection. Although IFN-g production has hitherto been generally regarded as nonpermissive for allograft survival, we believe this paradoxical “good–bad” role for IFN-g may reflect an important physiological negative feedback loop. Key words: IFN-g, Regulatory T cell, Transplantation

1. Introduction The role of IFN-g in cellular immunity is somewhat paradoxical in that although it is usually considered to be a proinflammatory effector cytokine, increasing evidence suggests that it plays a nonredundant immunoregulatory role (1–8). Although the classical view of IFN-g is that it favors Th1 cell development (9, 10), IFN-g also has regulatory functions. For example, IFN-g can inhibit the proliferation of IL-4-producing Th2 cells (11) and suppress the development of Th17 effector cells, now known to play an important role in many autoimmune models (12, 13). In addition, IFN-g also plays an important role in maintenance of T cell homeostasis by inducing apoptosis-dependent activationinduced cell death to limit T cell expansion following antigen encounter (14–19). In the context of adaptive regulation, we have recently shown that IFN-g is produced rapidly and transiently by Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_20, © Springer Science+Business Media, LLC 2011

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alloantigen-reactive Treg following reactivation and this is required for its functional activity in vivo (20). IFN-g can induce indoleamine 2,3-dioxygenase (IDO) in several cell types and that this enzyme has been shown to play an important role in limiting T cell responses in vivo (21–23). A paradoxical role for IFN-g is also seen in organ transplantation (24–26). Evidence of IFN-g induction is readily found in rejecting allografts (27, 28). However, IFN-g appears not to be essential for acute cellular rejection as both IFN-g deficient and wild type mice reject cardiac allografts with similar kinetics (29, 30). In fact, IFN-g may be required for successful engraftment (24, 31, 32). In this chapter, we describe a novel technique for shaping T cell responses ex vivo that could have cellular therapeutic potential in transplantation and autoimmunity.

2. Materials 2.1. Mice

CBA.Ca (CBA, H2k), C57BL/10 (B10, H2b), and CBArecombination-activating gene 1 knockout (CBA-Rag−/−, H2k; kindly provided by Dr. D. Kioussis, Division of Molecular Immunology, National Institute for Medical Research, Mill Hill, London, U K) mice were obtained from and housed in the Biomedical Services Unit, John Radcliffe Hospital (Oxford, UK). Sex-matched mice between 6 and 12 weeks of age at the time of first experimental procedure were used in all experiments (Table 1). All housing and procedures were in accordance with the Animals (Scientific Procedure) Act 1986.

Table 1 Mouse strains used Strain

Abbreviation

H2 haplotype

CBA/Ca

CBA

k

CBA-RAG1−/−

CBA-RAG−/−

k

C57BL/10

B10

b

BALB

d

BALBRAG−/−

d

C57BL/6

B6

b

C57BL/6 RAG1−/−

B6-RAG−/−

b

BALB/c BALB/c RAG2 common g chain KO

,

−/−

Notes

RAG1 deficient

RAG2 deficient, common g chain KO

RAG1 deficient

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283

Table 2 Antibodies used in vitro and in vivo Specificity

Clone

Species

Isotype

Source

Ref

CD4

YTS3.1.2

Rat

IgG2b

HW

(33)

CD8

YTS169.4.2

Rat

IgG2b

HW

(33)

MHC class II

M5/114.15.2

Rat

IgG2b, k

ATCC

(34)

Mac-1

M1/70

Rat

IgG2b, k

ATCC

(35)

CD45R/B220

RA3-6B2

Rat

IgG2a, k

ATCC

(36)

HW: Professor Herman Waldmann, Sir William Dunn School of Pathology, Oxford, UK; ATCC: American Type Culture Collection

2.2. Antibodies

Monoclonal antibodies (mAbs) used for cell purification and in vivo administration are listed in Table 2. The hybridomas YTS 177 and YTA 3.1 (kind gifts from Professor Herman Waldmann, Sir William Dunn School of Pathology, Oxford) were grown in roller cultures, and antibodies were purified by ammonium sulfate precipitation followed by DEAE Sephacel ion exchange chromatography. Other antibodies were purified on protein G columns by fast protein liquid chromatography (FPLC). mAbs used for flow cytometry were purchased from BD Pharmingen (San Diego, CA, USA) and are listed in Table 3. Foxp3 mAb (FJK-16s) and fixation/permeabilization solution were purchased from eBiosciences (CA, USA).

2.3. Other Reagents

All cell culture procedures were performed using complete culture medium consisting of RPMI 1640 containing 10% FCS, 2 mM l-glutamine, 0.5 mM 2-mercaptoethanol (Sigma), and 100 U/ml each of penicillin and streptomycin (Sigma). DynaBeads (Invitrogen) and MicroBeads (Miltenyi) were used for bead-based positive and negative selection. Recombinant mouse and human cytokines were purchased from PeproTech (London, UK). FACS buffer consists of PBS (Oxoid Ltd., Basingstoke, UK, containing 2.6 mM KH2PO4, 26 mM Na2HPO4, 145 mM NaCl, final pH 7.2) supplemented with 2% FCS (PAA Laboratories GmBH, Linz, Austria) and 0.02% sodium azide (Sigma). FACS buffer containing 1% v/v formaldehyde (Sigma, USA) is used for fixation (FACS Fix). FACS data were acquired on a FACSort flow cytometer (Becton Dickinson, San Jose, CA, USA) and analyzed using the CellQuest software package (Becton Dickinson). Domitor (Pfizer Pharmaceutical Ltd.), Ketaset (Fort Dodge Animal Health Ltd.), Antisedan (Pfizer Pharmaceutical Ltd.), Metacam (Boehringer Ingelheim, UK), and Vetergesic (Alstoe Ltd, UK) were used in surgical procedures. Flexible Collodion

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Table 3 Antibodies used for flow cytometry Specificity

Clone

Species

Isotype

CD3e

145-2C11

Armenian hamster

IgG1, k

CD4

RM4-5

Rat

IgG2a, k

CD11c

HL3

Armenian hamster

IgG1, l2

CD16/32

2.4G2

Rat

IgG2b, k

CD25

PC61

Rat

IgG1, l

CD40

3/23

Rat

IgG2a, k

CD62L

MEL-14

Rat

IgG2a, k

CD80

16-10A1

Armenian hamster

IgG2, k

CD86

GL1

Rat

IgG2a, k

CTLA4

UC10-4F10-11

Armenian hamster

IgG1, k

Foxp3

FJK16s

Rat

IgG2a, k

Rat IgG1

R3-34

Rat

IgG1, k

Rat IgG2a

R35-95

Rat

IgG2a, k

Hamster, IgG1

A19-3

Armenian hamster

IgG1

BP (William Ransom & Son, Herts, UK) and povidone-iodine nonadherent dressing (Inadine, Johnson & Johnson Medical, Ascot, UK) were used in skin transplantation.

3. Methods 3.1. Flow Cytometric Technique 3.1.1. Cell Surface Markers Staining

3.1.2. Transcription Factor Foxp3 Staining

1. 50 ml of cells at a concentration of 1 × 107/ml were incubated with 50 ml of FACS buffer containing the appropriate mAbs at 4°C for 20 min. 2. Following this incubation, the cells were washed in FACS buffer and resuspended for additional staining or fixed in 250 ml of FACS buffer containing 1% v/v formaldehyde (FACS Fix) and stored at 4°C until acquisition. 1. 50 ml of cells (5 × 105) were added to each tube and mixed with 0.5 ml of freshly prepared fixation/permeabilization working solution. 2. Cells were fixed and permeabilized at 4°C for 2 h in the dark. 3. Then, cells were washed twice with 1 ml 1× permeabilization buffer, and purified antiCD16/32 in 1× permeabilization

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buffer was added to block Fc receptors, in a 100 ml volume, at 4°C for 15 min. 4. Without washing, anti-mouse/rat Foxp3 antibody, diluted in 1× permeabilization buffer, was directly added and incubated at 4°C for 30 min in the dark. 5. Cells were washed twice with 1 ml 1× permeabilization buffer, then resuspended in 250 ml FACS buffer and acquired immediately. 3.2. Cell Purification 3.2.1. Single Cell Suspension from Spleen

1. Spleens were harvested and a single cell suspension was prepared by passing the tissue through sterile 70 mm cell strainers and washing in PBS containing 2% FCS. 2. Erythrocytes were removed by hypotonic lysis.

3.2.2. CD25+CD4+ and CD25−CD4+ T Cell Purification

1. CD4+ T cells were enriched by negative selection using DynaBeads. Rat anti-mouse mAbs used for CD4+ T cell negative selection were YTS 169, RA3.6B2 (B220), TIB120, and M1/70 (Table 2). 2. CD4+-enriched cells were then resuspended at 108/ml with MACS buffer and incubated with 50 ml anti-CD25-PE antibody for 15 min on ice. 3. Cells were then washed, resuspended at 108/ml with MACS buffer, and incubated with 50 ml anti-PE MicroBeads for 15 min at 4°C, and then washed, resuspended, and applied onto the rinsed column. 4. CD25−CD4+ T cells were collected from the negative selected part and CD25+CD4+ from the positive selected part. On reanalysis, all populations were more than 95% pure.

3.3. Cell CultureRelated Procedures 3.3.1. Generation of Bone Marrow-Derived Dendritic Cells

Bone marrow (BM) derived DCs were generated from B10 donors using a modification of published methods (37, 38). 1. Mouse femurs removed of muscles were placed in a 60-mm dish with RPMI 1640; both ends of the bone were cut and the marrow was flushed out using 2 ml RPMI 1640; RBCs were lysed by hypotonic shock. 2. B cells, T cells, and MHC class II positive cells were depleted using specific antibodies (RA3.6B2, YTS 3.1, YTS 169, and TIB120) followed by negative selection using antirat magnetic Dynabeads. 3. 1 × 106 enriched DC precursor cells were placed in 24-well plates in 1 ml of medium supplement with 2 ng/ml each of recombinant mouse GM-CSF and recombinant human TGF-b1 (see Note 1); 75% of the medium was changed every 48 h. 4. At day 6, DCs were harvested, washed, and counted prior to use.

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3.3.2. Ex Vivo Polarization and Conditioning Protocol

1. 5 × 105 Purified naive CBA CD4+ T cells were cocultured with 5 × 104 B10 BM DCs (g-irradiated, 3,000 rad) in the presence of 5 ng/ml exogenous recombinant mouse IFN-g (IFN-g conditioning) (see Note 2). 2. On day 7, half of the medium was exchanged with fresh media containing the same concentration of recombinant IFN-g and the same number of DCs. 3. After two rounds of stimulation, cells were harvested and then either characterized by phenotypic assay, or used in an adoptive transfer model for functional evaluation (see Note 4).

3.4. Surgical Procedure 3.4.1. Anesthesia

1. General anesthesia was induced using a combination of Domitor and Ketaset. Domitor was diluted with sterile saline to a working solution containing 100 mg/ml medetomidine hydrochloride, and Ketaset was diluted to give a working solution of 7.6 mg/ml ketamine hydrochloride. Mice were injected subcutaneously with 0.15 ml of each of the Domitor and Ketaset working solution before any procedure. 2. Analgesia: Vetergesic was diluted with sterile saline to provide a working solution containing 10 mg/ml buprenorphine hydrochloride. Metacam was diluted with sterile saline to a working solution containing 250 mg/ml meloxicam. Mice were injected subcutaneously with 0.15 ml of each of the Vetergesic and Metacam working solution before any procedure. 3. Antisedan, sedation-reversal agent for Domitor, was diluted with sterile saline to a working solution containing 100 mg/ml atipamezole hydrochloride. Each anesthetic procedure was followed by injection of 0.30 ml Antisedan working solution.

3.4.2. Adoptive Transfer and Skin Grafting

1. CBA-Rag−/− mice were reconstituted intravenously with 1 × 105 CD25−CD4+ cells from naive CBA with or without 2 × 105 in vitro-conditioned cells (Fig. 1). 2. The following day full-thickness B10 tail skin allografts were transplanted onto graft beds prepared on the flanks of the reconstituted mice. Briefly, donor tail skin was cut into individual grafts approximately 12 mm in length. Under general anesthesia, a graft bed was prepared on the flank of the recipient. The graft was placed on the bed and secured by applying Flexible Collodion BP. A povidone-iodine nonadherent dressing was placed over the graft and covered with adhesive tape wrapped around the animal. The dressing was removed 7 days later. 3. Grafts were inspected regularly and graft rejection was defined as complete necrosis of the donor skin. Graft survival between groups was compared using the Kaplan–Meier log-rank test with SPSS software.

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287

B10 H-2b (donor ) cytokines

Harvest

CBA H-2k (recipients)

conditioned cells

CD25negative effector cells 24 hours

T deficient CBA

donor-specific skin graft

Fig. 1. In vitro cytokine conditioning and adoptive transfer system. Purified naive CBA CD4 T cells were cocultured with B10 splenic APCs under neutral conditions (no added cytokines or antibodies), or in the presence of IFN-g (IFN-g conditioning). After 2–3 rounds of stimulations, cells were harvested. Cells were then characterized by phenotypic assay, or used in an adoptive transfer model for functional evaluation.

3.5. IFN-g Conditioning Results in Up-Regulation of Foxp3 Expression

1. Foxp3 is a transcription factor closely associated with regulatory T cells (39–41). In order to ask whether the effect of IFN-g conditioning could lead to an increase in Foxp3 expression in single cells level, CBA CD4+ T cells were stimulated with B10 BM DCs in the presence or absence of IFN-g (0.5, 5, and 50 ng/ml). As shown in Fig. 2a, coculture of purified naive CBA CD4+ T cells with GM-CSF/TGF-b B10 bone marrow (BM)-derived DCs in the presence of exogenous IFN-g resulted in a dose-dependent up-regulation of Foxp3 expression. Optimal up-regulation was seen at 5 ng/ml IFN-g. CBA CD4+ T cells were also stimulated with B10 BM DCs differentiated with GM-CSF, only in the presence of 5 ng/ml IFN-g, and in this situation, no significant up-regulation of Foxp3 was observed (not shown) (see Note 1). As shown in Fig. 2b, up-regulation of Foxp3 was also associated with increased expression of CD25 and CD62L. Taken together, these data indicate that CD4+ T cells stimulated with allogeneic BM DCs up-regulate Foxp3 expression. 2. To determine whether induced Foxp3 expression results in regulatory function in vivo, CD4+ responder cells conditioned with 5 ng/ml IFN-g as shown in Fig. 1 were transferred (2 × 105) into CBA-Rag−/− mice together with 1 × 105 CD25−CD4+ syngeneic cells as an effector population. As shown in Fig. 2c,

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mice reconstituted with CD25−CD4+ cells only rejected B.10 skin allografts acutely (MST = 25). In contrast, mice that received CD25− cells plus CD4+ cells conditioned in the presence of IFN-g accepted their grafts long term (MST > 100) (Fig. 2c). Thus, IFN-g can be used ex vivo to shape the alloreactive immune response ex vivo and lead to a population enriched for regulatory T cells. 3.6. Generation of IFN-g Conditioned Treg in Different Strain Combinations

Work from this laboratory has previously shown that the generation of Treg in vivo using an anti-CD4 antibody plus DST protocol is influenced significantly by the recipient mouse strain used. Given the fact that the genetic background can affect Th1/Th2 cell differentiation (42), it was possible that this might also have an impact on Treg generation. In order to determine whether IFN-g conditioning established using CBA T cells driven by B10 APC is relevant in other situations, two different responder/ stimulator combinations were compared in parallel with CBA cells driven by B10 BM DC as a positive control (Fig. 3).

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Fig. 2. (continued) (b) Purified naive CBA CD4 T cells were cocultured with B10 GM-CSF/TGF-b-differentiated bone marrow dendritic cells (GT-DC) in the presence of exogenous IFN-g (5 ng/ml), restimulated under the same conditions on day 7, harvested on day 14, and stained for intracellular Foxp3, CD25, and CD62L. Dot plots are gated on T cells and numbers indicate the frequency of cells in each quadrant. Data are representative of two independent experiments. (c) CBA-RAG−/− mice were reconstituted with syngenic CD25−CD4+ T cells, with or without cotransfer of 2 × 105 IFN-gconditioned T cells, and 1 day later these mice received allogeneic skin grafts.

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a 5ng/ml IF N -g

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Fig. 3. Titration of IFN-g in various stimulator–responder combinations: B10 to CBA, BALB/c to B6, and B10 to BALB/c. (a) CBA CD4 + T cells were conditioned with GM-CSF/TGF-b-differentiated B10 BM DCs and IFN-g (0.5–50 ng/ml). (b) B6 CD4 + T cells were conditioned with GM-CSF/TGF-b-differentiated BALB BM DCs and IFN-g (0.5–50 ng/ml). (c) BALB CD4+ T cells were conditioned with GM-CSF/TGF-b-differentiated B10 BM DCs and IFN-g (0.5–50 ng/ml). After two rounds of stimulation, harvested conditioned cells were stained for Foxp3 expression.

1. B6 CD4+ T cells were driven with GM-CSF/TGF-bdifferentiated BM DCs from BALB/c mice, and BALB/c CD4+ T cells were driven by GM-CSF/TGF-b-differentiated BM DCs from B10 mice. IFN-g was used at three different concentrations on the basis that the response to IFN-g might

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vary between the three responder populations. Cells were harvested on day 14 and analyzed for Foxp3 expression by FACS. As shown in Fig. 3, the percentage of cells positive for Foxp3 after culture in the absence of IFN-g was 9, 1, and 5% in CBA, B6, and BALB/c T cells, respectively. In the presence of 5 ng/ml IFN-g (the “standard” concentration used throughout this chapter), both CBA cells and B6 cells showed a significant Foxp3 up-regulation (34 and 27% respectively). Significantly, however, the presence of IFN-g at this concentration had only a modest effect on Foxp3 expression in BALB/c cells (10%) and was unaffected by addition of this cytokine at a tenfold higher concentration (8%). Therefore, in terms of Foxp3 expression, BALB/c cells were relatively resistant to the effect of IFN-g in cultures shown to drive the generation of Treg in CBA cells, stimulated by the same BM DC population (see Note 3). 2. In order to determine whether Foxp3 expression does indeed reflect the regulatory property of these conditioned cells, B6 or BALB/c CD4+ responder cells conditioned with 5 ng/ml IFN-g as shown in Fig. 3 were transferred (2 × 105) into syngeneic immunodeficient recipients, together with 1 × 105 CD25−CD4+ syngeneic cells as an effector population. Mice reconstituted with CD25−CD4+ cells served as rejection controls. B6 cells driven under the same conditions were capable of regulating rejection of donor-specific allografts, although two animals rejected their grafts albeit with much delayed kinetics (Fig. 4a). Importantly, however, cells derived from BALB/c mice were unable to prevent acute graft rejection (Fig. 4b). Given the fact that the IFN-g conditioning protocol led to a significant up-regulation of Foxp3 in CD4+ T cells from CBA and B6 mice but not from BALB/c mice (Fig. 3), these data demonstrate a clear functional correlation between Foxp3 expression and acquisition of regulatory function in this system.

4. Notes 1. TGF-b is essential for successful conditioning of BM DC. DCs play an important role in the development of naturally occurring Treg in the thymus, and immature DCs have been shown to be favorable for the generation of IL-10-producing Tr1 cells ex vivo (43). The BM DCs used in the IFN-gconditioning protocol described in this chapter are conditioned with TGF-b and GM-CSF and although exogenous TGF-b is not added to the T cell-DC cultures, TGF-b is

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Fig. 4. In vivo function of IFN-g-conditioned cells in the B10 to BALB/c and BALB/c to B6 donor/recipient strain combinations. Immunodeficient recipient mice were reconstituted with syngeneic CD25−CD4+ T cells, with or without cotransfer of 2 × 105 IFN-gconditioned T cells, and 1 day later these mice received allogeneic skin grafts.

essential for the correct maturation of the BM DC population in that BM DCs conditioned with GM-CSF in the absence of TGF-b are unable to drive the up-regulation of Foxp3 in responding T cells (43). TGF-b has been shown to be capable of blocking DC maturation (38) and as shown in Fig. 5, GM-CSF plus TGF-b-differentiated BM DCs express a lower level of the costimulatory molecules CD40, CD80, and CD86 compared with DCs differentiated in the absence of TGF-b, and thus have a relatively immature phenotype. In addition, as shown in Fig. 5b, BM DCs generated in the presence of GM-CSF plus TGF-b produce less IL-6, a proinflammatory cytokine that is known to be incompatible with Treg generation (44). Therefore, a suboptimal APC maturation status (45) together with favorable cytokine environment (44) appears to be essential for Treg generation in this system.

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Fig. 5. (a). Bone marrow-derived dendritic cells generated in the presence of GM-CSF plus TGF-b express lower levels of costimulatory molecules. B10 bone marrow cells were collected after precursor enrichment and cultured in the presence of 2 ng/ml GM-CSF with (GT-DC) or without (G-DC) 2 ng/ml TGF-b. Cells were harvested on day 6, and CD86, CD80, and CD40 expression was examined on CD11c-gated cells. Overlay of G-DC and GT-DC staining: filled curve represents GT-DC, solid curve represents G-DC, and dotted curve represents isotype control. Data are representative of three independent experiments. (b). Bone marrow-derived dendritic cells generated in the presence of GM-CSF plus TGF-b produce less IL-6. Bone marrow cells were collected after precursor enrichment and cultured in the presence of 2 ng/ml GM-CSF with (GT-DC) or without (G-DC) 2 ng/ml TGF-b, harvested at day 6, and stimulated in the absence of T cells with recombinant CD40L for 24 h to mimic reverse stimulation by CD40–CD40L interactions. RNA was extracted and Taqman analysis performed. Results are normalized to a housekeeping gene GAPDH, and data are means with SEM (error bars) of two independent experiments (upper panel ). The supernatant was also collected for ELISA analysis and results are normalized to 4 × 105 cells in 2 ml medium (lower panel ).

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2. Dual role of 2-mercaptoethanol in the generation of Treg. 2-Mercaptoethanol (2-ME) is a derivative of ethanol, which contains a thiol (–SH) group and acts as a biological antioxidant by releasing thiol. As a reducing agent, 2-ME is widely used to protect proteins from inactivation, and adding 2-mercaptoethanol to ex vivo mouse lymphocyte cultures has been a standard practice for many years where its function is to improve cell survival and growth by promoting cystine uptake (46). Cystine is an essential amino acid normally added in the culture medium and contributes to the maintenance of a certain level of intracellular glutathione necessary for cell survival and proliferation (47). 2-ME may also increase cell viability by scavenging oxygen-free radicals (48) and maintaining the integrity of cytokine signaling transduction (49). 2-ME is also reported to promote nonspecific polyclonal T cell activation and proliferation (50, 51). We have shown that nitric oxide (NO) plays an import role in IFN-g conditioning and up-regulation of Foxp3 expression (52). Since 2-ME may partially antagonize the function of NO, a thiol oxidant, it may attenuate the effect of NO in the up-regulation of Foxp3 expression. In the ex vivo Treg generation protocol described in this chapter, 2-ME was used in all tissue culture media at a concentration of 0.5 mM. This concentration was based on historical experience in this laboratory, but it was considered possible that because of the connection between NO and Foxp3 expression, 0.5 mM 2-ME may be far from the optimal concentration. In order to test this, purified naive CBA CD4+ T cells were cocultured with GM-CSF/TGF-b-generated B10 BM DCs in the presence of 5 ng/ml IFN-g plus various concentrations of 2-ME (0.005–5 mM). Cells were harvested on day 14, and intracellular Foxp3 expression was analyzed by FACS. As shown in Fig. 6, the proportions of Foxp3+ cells were 14, 28, 39, 29, and 23% when the 2-ME concentrations were 0, 0.005, 0.05, 0.5, and 5 mM, respectively. Thus, in terms of Foxp3 expression, the optimal concentration of 2-ME was tenfold lower than that routinely used in the ex vivo-conditioning protocol. 3. Genetic background can determine the results of IFN-g conditioning. The genetic background of the cell donor can affect in vitro Th1/Th2 cell differentiation (42), immune responses against infectious disease (53), and graft outcome (54). CD4+ T cells from CBA and C57BL/6 mice respond similarly to IFN-g conditioning in terms of up-regulation of Foxp3 expression, with about 30% of cells becoming positive for Foxp3 when IFN-g was added at 5 ng/ml or 50 ng/ml (Fig. 3).

Regulatory T Cell Enrichment by IFN-g Conditioning No 2-ME 14% M1

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Fig. 6. The effect of 2-mercaptoethanol on Foxp3 expression. CBA CD4+ cells were cultured with GM-CSF/TGF-b-differentiated B10 BM DCs in the presence of different concentrations of 2-mercaptoethanol. All cultures contained IFN-g (5 ng/ml). Cells were harvested at day 14 for Foxp3 staining. Populations were gated on CD4+ cells. 2-ME: 2-Mercaptoethanol (mM); currently used at 0.5 mM in the standard protocol.

However, CD4+ T cells from BALB/c mice respond poorly to IFN-g conditioning with a maximum of 10% Foxp3+ cells at 5 ng/ml. This is not due to insufficient IFN-g because when IFN-g concentration was increased to 50 ng/ml, the Foxp3+ proportion was 8%. Importantly, as shown in Fig. 4, the proportion of Foxp3+ cells correlated with the graft outcome. After being reconstituted with ex vivo IFN-gconditioned cells and CD25−CD4+ effector cells, all BALB/ c-Rag−/− recipients rejected their grafts in 28 days post transplantation, in contrast to CBA-Rag−/− and B6-Rag−/−

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recipients which accepted their grafts long term. If the potential of this type of approach for Treg generation is to be translated into a potential clinical protocol, it is important to know what makes BALB/c CD4+ T cells poor responders to IFN-g-induced Foxp3 expression. It has been shown that BALB/c mice produce a burst of IL-4 in their draining lymph nodes within 24 h of Leishmania infection (55, 56) and this burst of endogenous IL-4 derived from naive BALB/c CD4+ T cells was sufficient to cause the in vitro differentiation of CD4+ T cells into Th2 cells (57). The effect of the Th2-polarizing cytokine IL-4 in inducing Th2 development can be dominant over Th1-polarizing cytokines, and if the IL-4 concentration reaches a certain threshold at the initiation of an immune response, Th2 cells will differentiate with priority and dominate the immune response (10). Recombinant IL-4 (6 ng/ml) added to IFN-g conditioning was sufficient to abolish the induction of Foxp3 expression in CBA CD4+ T cells (58), suggesting that one of the possibilities to explain the failure to induce Foxp3+ expression in BALB/c CD4+ T cells may be due to a critical change of the cytokine environment required for a successful IFN-g conditioning, and endogenous IL-4 may account for this change. Therefore, one modification to the standard IFN-g-conditioning protocol would be adding an anti-IL-4 antibody. This has yet to be formally tested. 4. Using yield and Annexin V staining as a gage of quality control of IFN-g conditioning. We have found that the presence of exogenous IFN-g significantly reduces the overall number of cells recovered by two- to fivefold in successful conditioning and that profound proliferation is usually incompatible with the development of a dominant Foxp3+ population (data not shown). In fact, a reduced cell yield (10–20% of input cell number) can be a simple initial indicator of successful conditioning. Given that IFN-g contributes to T-cell homeostasis in vivo through the inhibition of proliferation and/or by increasing activation-induced cell death (AICD) (14–16, 19), a simple explanation for the increased proportion of Foxp3+ cells in the IFN-g conditioning protocol might be a preferential elimination of Foxp3− cells. To examine this in more detail, CBA CD4 T cells were driven by GM-CSF/TGF-b B10 BM DC, harvested, and stained for annexin V as a marker of necrotic or apoptotic cell death, washed, fixed, and stained for Foxp3, thus allowing the analysis of apoptosis within both Foxp3+ and Foxp3− populations. As shown in Fig. 7, the presence of IFN-g enhances cell death within the Foxp3− population and importantly, when cells were analyzed separately on the

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Foxp3 P90%) from these cultures (7, 8, 9, 15) (see Note 4). 3.2. Isolation of Splenic T Cells

1. Spleens were removed from 6 to 8 weeks-old OT-I mice (7, 8, 9, 15). 2. The spleens were homogenized by using the grinding zones of two microscope slides in 10 ml of RPMI 1640 medium on a petri dish. The homogenate was transferred into a 15 ml tube and let cells to sediment. The supernatant was transferred into a new tube and centrifuged it for 10 min at 300 × g at 4°C (7, 8, 9, 15). 3. The cellular pellet was resuspended in 10 ml of ACK buffer and incubated for 2 min at room temperature and then it was centrifuged for 10 min at 300 × g at 4°C (7, 8, 9, 15). 4. The pellet was retrieved and washed twice with sterile cold PBS (7, 8, 9, 15). 5. Cells were obtained from the pellet and resuspended in 500 ml of MACS buffer. CD8+ T-cell purification was performed by using MACS CD8+ negative selection isolation kit (7, 8, 9, 15). 6. CD8+ T-cell purity was analyzed by staining the purified cell fraction with fluorochrome-conjugated monoclonal antibody specific for CD8 (7, 8, 9, 15).

3.3. Induction of Tolerogenic DCs with Andrographolide and Rosiglitazone

1. To induce tolerogenic DCs with andrographolide or rosiglitazone, fresh solutions of these drugs were prepared immediately before each experiment (0.5 mM, see Subheading 2.2). It is recommendable to prepare a fresh drug working solution for each experiment given that drugs can lose pharmacological activity (7, 9). 2. The induction of tolerogenic DCs for further assays (see below) was achieved by treating DCs at day 5 of culture with 10 mM of andrographolide or 10 mM of rosiglitazone for 24 h (7, 9). 3. In order to verify the tolerogenic phenotype of either andrographolide- or rosiglitazone-treated DCs, the drugs were washed out (washing twice with sterile PBS) after the treatment, and maturation was induced with 1 mg/ml LPS (Sigma) for 36 h. After LPS-induced maturation, DC phenotype was analyzed by staining with fluorochrome-conjugated antibodies specific for CD11c, I-Ab, H-2Kb, CD80, CD86, and CD40. The analysis for the expression of each marker was performed by flow cytometry. Drug-treated DCs showed significantly lower levels of co-stimulatory molecules as compared to the untreated

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DCs. These data suggest that treatment with these drugs imprints an immature-tolerogenic phenotype to DCs (7, 9). 4. To be sure that the tolerogenic DCs generated by this method have a reduced capacity to activate antigen-specific T cells, andrographolide- or rosiglitazone-treated DCs were pulsed for 16 h with the OVA-derived peptide SIINFEKL at a final concentration equal to 10 ng/ml. DCs pulse with antigen was performed directly on 24-well plates. After SIINFEKL pulse, DCs were harvested and washed twice with sterile PBS. Then, DCs were co-cultured with OT-I T cells (2.5 × 104 DCs and 5 × 104 OT-I T cells on 96-well plates in supplemented RPMI 1640 medium). After 20 h of DC-T cell co-culture, IL-2 was measured on the supernatants by cytokine ELISA using specific antibodies (see Subheading 2.2, step 6). ELISA was performed using Streptavidin-HRP and revealing with TMB. The reaction was stopped with 2 M H2SO4 after 10 min. The IL-2 concentration was analyzed by reading absorbance at 450 nm on an ELISA plate reader (7, 9). 3.4. Induction of EAE in Mice

1. EAE was induced in 6- to 8-week-old female C57BL/6 mice. Mice were injected subcutaneously with 50 mg of MOG35-55 peptide emulsified in Complete Freund’s adjuvant supplemented with heat-inactivated Mycobacterium tuberculosis H37. To promote brain–blood barrier permeability, mice were injected intraperitoneally with 500 ng of pertussis toxin at the time of sensitization and 48 h after the injection with MOG35-55 peptide (7, 9, 16). 2. The mice were monitored on a daily basis for clinical signs of EAE-caused damage. Usually between 15 and 18 days after sensitization, mice develop clear signs of EAE. To score the disease severity, the following criteria can be used (7, 9, 16): 0 when mice show no signs of disease 1 when mice have flaccid tails 2 when mice show abnormal gait or hind limb weakness 3 when mice show complete hind limb paralysis 4 when mice show paralyzed fore and hind limbs 5 when mice are moribund or dead 3. To prevent unnecessary animal suffering, those mice that were severely affected by EAE were euthanized by cervical dislocation with the supervision of a veterinarian. 4. Clinical scores were recorded on a daily basis and a curve for the progression of disease over time was generated with these data for each mouse (7, 9, 16).

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1. DCs were treated for 24 h with either andrographolide or rosiglitazone (according to Subheading 3.3) and then pulsed with the MOG35-55 peptide at a concentration equal to 5 mg/ml in 24-well plates (106 DCs/well) during an extra incubation time equal to 24 h (7, 9). 2. After antigen pulse, a cell suspension consisting of 2 × 106/ml of tolerogenic DCs treated with either andrographolide or rosiglitazone and with MOG35-55 peptide was prepared in sterile PBS (7, 9). 3. 106 DCs/mice of the tolerogenic DC suspension was intravenously injected into 6- to 8-week-old female C57BL/6 mice 1 and 2 weeks before EAE induction (7, 9). 4. Two weeks after the first tolerogenic DCs injection, EAE was induced and clinical symptoms of disease were monitored as described in Subheading 3.4 (7, 9). 5. The analyses of the clinical score for each group of mice showed that those animals that received tolerogenic DCs did not develop EAE as did the control mice (a group that received only PBS) (Fig. 2) (7, 9).

3.6. Direct Treatment with Andrographolide or Rosiglitazone to Reduce EAE in Mice

1. In order to promote tolerance via endogenous DCs, 6- to 8-week-old female C57BL/6 mice were intraperitoneally injected with a daily dose of 4 mg/kg of andrographolide or rosiglitazone in 100 ml of PBS. After 1 week of treatment with either drug, EAE was induced in these mice according to the method described in Subheading 3.4. The drug treatment continued through all the experiment period. The drugs were well tolerated by the mice and no signs of toxicity were observed at the indicated concentrations (7, 9). 2. Clinical scores for each mouse were evaluated on a daily basis as stated in Subheading 3.4. Our data showed that mice treated with either andrographolide or rosiglitazone did not develop measurable signs of EAE, as compared to control animals that received PBS (7, 9).

3.7. Treatment with Andrographolide or Rosiglitazone to Prevent SLE in Mice

1. To analyze the effect of andrographolide or rosiglitazone on SLE, we used the FcgRIIb−/− mice, which spontaneously develop a SLE-like disorder. It is recommendable to use female mice due to their higher susceptibility to develop SLE (8). 2. As an attempt to prevent SLE development, one group of FcgRIIb−/− mice were treated intraperitoneally with 4 mg/kg of andrographolide or rosiglitazone in 100 ml of PBS, twice a week. Treatment started at the age of 6 weeks and extended until 7 months old. At the same time, a control group of mice was treated with PBS (8).

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Fig. 2. Myelin-specific tolerance induced by DCs treated with NF-kB inhibitors. (a) DCs derived from C57BL/6 mice are treated with either andrografolide or roziglitazone to inhibit NF-kB function and pulsed with myelin antigens, as represented in the diagram. (b) C57BL /6 mice received two i.v. injections of either PBS, immature DCs presenting myelin antigens or andrografolide- or roziglitazone-treated DCs presenting myelin antigens. DC injections are given 1 and 2 weeks before experimental autoimmune encephalomyelitis (EAE) induction, as represented in the diagram. Clinical EAE symptoms were monitored on a daily basis, showing that mice receiving DCs treated with either andrografolide or roziglitazone were significantly protected from EAE and showed large amounts of regulatory T cells.

3. SLE progression was monitored on a weekly basis for both groups of mice, starting at the age of 2 months. Blood and urine samples were collected from control and treated mice for the assessment of anti-nuclear antibodies (ANAs) and proteinuria, respectively (8). 4. For ANAs determination, serum was obtained from blood samples. The blood was centrifuged for 10 min at 250 × g at 4°C and the supernatant was collected. Then, serum dilutions were added to 12-well plates containing HEp-2 cells. These cells were incubated for 30 min at room temperature. After this time, FITC-conjugated anti-mouse IgG antibody was

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added to the cells. The presence of IgG in HEp-2 cells was analyzed by fluorescence microscopy, which indicated the presence of autoantibodies (8). 5. The protein level in the urine was measured by using Combur Test sticks for urinalysis. This test uses a scale of 0–3, where 0/trace = negative, 1 = 30 mg/dl, 2 = 100 mg/dl, and 3 = 500 mg/dl. Scores above 2 are considered to present severe glomerulonephritis, according to the manufacturer instructions (8). 6. To determine kidney damage caused by the inflammation induced by SLE, histological analyses were performed. To observe IC deposition, kidneys were embedded in the histology tissue freezing medium Tissue-tek OCT compound, and then snap frozen. Ten-micrometer sections were obtained by cutting with cryostate and then they were fixed with ice-cold acetone and stained with FITC-conjugated anti-mouse IgG antibody. Analysis of IgG deposition was performed by fluorescence microscopy (8). 7. Histological data indicated that the treatment with andrographolide or rosiglitazone prevented SLE development. In contrast, control mice developed SLE (8). 3.8. Determination of Activating/ Inhibiting Fcg Receptor Ratio on Splenic DCs

1. Spleens were removed from C57BL/6 mice or SLE-prone mice. 2. Spleens were homogenized by using the grinding zones of two microscope slides in 10 ml of RPMI 1640 medium on a petri dish. Homogenates were transferred into a 15 ml tube and let sediment for 5 min. Supernatants were transferred into a new tube and centrifuged for 10 min at 300 × g at 4°C. 3. Pellets were resuspended in 10 ml of ACK buffer and incubated for 2 min at room temperature and then centrifuged for 10 min at 300 × g at 4°C. 4. Cellular pellets were saved and washed twice with sterile PBS. Cells were resuspended in 2% PBS–BSA and stained with antibodies specific for CD11c, FcgRIIb/FcgRIII (clone 2.4G2), and FcgRIIb (clone Ly17.2). 5. Stained cells were analyzed by flow cytometry and the ratio FcgRIIb/FcgRIII was calculated using the formula (MFI 2.4G2 − MFI Ly17.2)/MFI Ly17.2. Flow cytometry data indicated that FcgRIIb/FcgRIII ratio is higher in NZB/ NZW SLE-prone mice than C57BL/6 mice. MFI, Mean fluorescence intensity.

3.9. Determination of Activating/ Inhibitory Fc g Receptor Ratio on Human DCs

1. Peripheral blood mononuclear cells (PBMC) were obtained from blood samples of healthy donors and SLE patients after separation from whole blood by centrifugation in a Ficoll gradient. Then, PBMC were washed twice with XVIVO-15 medium (Bio-Whittaker), resuspended at 3 × 106 cells/ml in the same

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medium supplemented with 1% autologous serum (herein, hDC medium), and incubated at 37°C for 2 h in 10 ml flasks (19). 2. After 2 h of incubation, nonadherent cells were washed out for the flasks four times with hDC medium and then remaining adherent cells were cultured in the same medium at 37°C. Adherent cells (mainly monocytes) were differentiated to DCs during 6 days by adding 1,000 U/ml IL-4 and 1,000 U/ml GM-CSF every 2 days. DC maturation was induced by adding 5 mg/ml of LPS for 48 h at day 6 of culture. DC phenotype was analyzed and confirmed by FACS using specific antibodies against CD11c, CD40, CD80, CD83, and CD86 (19). 3. After hDC differentiation, cells were harvested and stained with antibodies against CD11c, anti-CD32a, and anti-CD32b and then the relative expression was analyzed by flow cytometry. The activating/inhibitory Fcg receptor ratio was calculated using the formula CD32a MFI/CD32b MFI. Our data suggest that CD32a/CD32b expression ratio is higher on SLE patients than healthy donors (18).

4. Notes 1. OT-I mice were used to track antigen-specific CD8+ T-cell responses. (Also, OT-II mice were used to track CD4 T-cell responses, but for simplicity these data were excluded from this chapter.) These methods are not restrictive to these mice and another transgenic mice can be used to follow antigenspecific CD8+ T-cell responses. 2. We highly recommend the recombinant GM-CSF purchased from Prepotech, it worked consistently well in our hands to promote the efficient differentiation of CD11c+ DCs. 3. We have found that antibodies purchased from Pharmingen worked well most of the times for flow cytometry experiments. 4. When the yield of CD11c+ cells was less than 90%, we proceeded to purify CD11c+ cells before continuing with the experimental protocol.

Acknowledgments Authors would like to thank all the trainees who contributed to the articles from our group cited in this publication. This work was supported by Grants FONDECYT 1070352 and 1085281 and Millennium Nucleus on Immunology and Immunotherapy (P07/088-F)

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References 1. Banchereau, J., Briere, F., Caux, C., Davoust, J., Lebecque, S., Liu, Y. J., Pulendran, B., and Palucka, K. (2000) Immunobiology of dendritic cells, Annu Rev Immunol 18, 767–811. 2. Iruretagoyena, M. I., Wiesendanger, M., and Kalergis, A. M. (2006) The dendritic cell-T cell synapse as a determinant of autoimmune pathogenesis, Curr Pharm Des 12, 131–147. 3. Iruretagoyena, M. I., Lezana, J. P. and Kalergis, A. M. (2005) Interactions at the Dendritic Cell/ T-Cell Interface Define the Balance between Immunity and Tolerance. Transfus Med Hemother, 32:373–383. 4. Ouaaz, F., Arron, J., Zheng, Y., Choi, Y., and Beg, A. A. (2002) Dendritic cell development and survival require distinct NF-kappaB subunits, Immunity 16, 257–270. 5. Zanetti, M., Castiglioni, P., Schoenberger, S., and Gerloni, M. (2003) The role of relB in regulating the adaptive immune response, Ann N Y Acad Sci 987, 249–257. 6. Saccani, S., Pantano, S., and Natoli, G. (2003) Modulation of NF-kappaB activity by exchange of dimers, Mol Cell 11, 1563–1574. 7. Iruretagoyena, M. I., Sepulveda, S. E., Lezana, J. P., Hermoso, M., Bronfman, M., Gutierrez, M. A., Jacobelli, S. H., and Kalergis, A. M. (2006) Inhibition of nuclear factor-kappa B enhances the capacity of immature dendritic cells to induce antigen-specific tolerance in experimental autoimmune encephalomyelitis, J Pharmacol Exp Ther 318, 59–67. 8. Kalergis, A. M., Iruretagoyena, M. I., Barrientos, M. J., González, P. A., Herrada, A. A., Leiva, E. D., Gutiérrez, M. A., Riedel, C. A., Bueno, S. A., and Jacobelli, S. H. (2009) Modulation of nuclear factor-kB activity can influence the susceptibility to systemic lupus erythematosus. Immunology 128, e306–e314. 9. Iruretagoyena, M. I., Tobar, J. A., Gonzalez, P. A., Sepulveda, S. E., Figueroa, C. A., Burgos, R. A., Hancke, J. L., and Kalergis, A. M. (2005) Andrographolide interferes with T cell activation and reduces experimental autoimmune encephalomyelitis in the mouse, J Pharmacol Exp Ther 312, 366–372.

10. Kelly, D., Campbell, J. I., King, T. P., Grant, G., Jansson, E. A., Coutts, A. G., Pettersson, S., and Conway, S. (2004) Commensal anaerobic gut bacteria attenuate inflammation by regulating nuclear-cytoplasmic shuttling of PPARgamma and RelA, Nat Immunol 5, 104–112. 11. Nimmerjahn, F., and Ravetch, J. V. (2007) Fc-receptors as regulators of immunity, Adv Immunol 96, 179–204. 12. Ravetch, J. V., and Bolland, S. (2001) Igg fc receptors, Annu Rev Immunol 19, 275–290. 13. Nimmerjahn, F., Bruhns, P., Horiuchi, K., and Ravetch, J. V. (2005) FcgammaRIV: a novel FcR with distinct IgG subclass specificity, Immunity 23, 41–51. 14. Kalergis, A. M. (2003) Modulation of T cell immunity by TCR/pMHC dwell time and activating/inhibitory receptor pairs on the antigen-presenting cell, Curr Pharm Des 9, 233–244. 15. Kalergis, A. M., and Ravetch, J. V. (2002) Inducing tumor immunity through the selective engagement of activating Fcgamma receptors on dendritic cells, J Exp Med 195, 1653–1659. 15. Iruretagoyena, M. I., Riedel, C. A., Leiva, E. D., Gutierrez, M. A., Jacobelli, S. H., and Kalergis, A. M. (2008) Activating and inhibitory Fcgamma receptors can differentially modulate T cell-mediated autoimmunity, Eur J Immunol 38, 2241–2250. 17. Bolland, S., and Ravetch, J. V. (2000) Spontaneous autoimmune disease in Fc(gamma) RIIB-deficient mice results from strain-specific epistasis, Immunity 13, 277–285. 18. Clynes, R., Dumitru, C., and Ravetch, J. V. (1998) Uncoupling of immune complex formation and kidney damage in autoimmune glomerulonephritis, Science 279, 1052–1054. 19. Carreño L.J., Pacheco, R., Gutierrez, M. A., Jacobelli, S., and Kalergis, A. M. (2009) Disease activity in systemic lupus erythematosus is associated with an altered expression of low-affinity-FcgRs and costimulatory molecules on dendritic cells. Immunology 128, 334–341.

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Chapter 23 Fine-Tuning Antitumor Responses Through the Control of Galectin–Glycan Interactions: An Overview Mariana Salatino and Gabriel A. Rabinovich Abstract In recent years, we have witnessed critical advances in genomics and proteomics which contributed to delineate the “tumor progression signature”. This includes the altered expression of genes and proteins not only in tumor cells, but also in tumor-associated stromal, endothelial, and immune cells. Adding more complexity to this bewildering information, efforts are being made to define the “glycosylation signature” of the tumor microenvironment, which results from the abnormal expression and activity of glycosyltransferases, glycosidases, and enzyme chaperons. The multiple combinatorial possibilities of glycan structures expressed by neoplastic versus normal tissue provide enormous potential for information display and expand potential therapeutic opportunities. The responsibility of deciphering the biological information encoded by the tumor-associated glycome is partially assigned, to distinct families of endogenous glycanbinding proteins or lectins, whose expression and function are regulated in cancerous tissues. Galectins, a family of evolutionarily conserved glycan-binding proteins, can control tumor progression by directly influencing tumor growth or by modulating cell migration, angiogenesis, and tumor–immune escape. In this review, we will highlight recent findings on how galectin–glycan lattices control the dialogue between tumor and immune cells and how these interactions could be exploited for therapeutic purposes. Key words: Glycosylation, Cancer, Tumor microenvironment, Tumor immunity, Galectins

1. Glycomics of the Tumor Microenvironment

Glycans decorate eukaryotic cell surfaces, where they are poised to mediate a variety of cell surface recognition events including host–pathogen and host–tumor interactions, leading to a wide variety of signaling processes and cellular responses (1). Glycan structures are incorporated to macromolecules such as proteins and lipids through a coordinated process termed “glycosylation” that involves the synchronized action of glycanmodifying enzymes; namely glycosyltransferases and glycosidases.

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_23, © Springer Science+Business Media, LLC 2011

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The expression and activity of these enzymes are exquisitely regulated according to cell fate and microenvironmental stimuli. Hence, cell surface glycosylation is altered not only during physiological processes such as immune cell activation, differentiation, and trafficking, but also during pathological settings including inflammation and cancer (1, 2). Thus, the information encoded by the “glycome” (i.e., the entire repertoire of sugar structures expressed in cells and tissues in physiological and pathological settings) may provide clues to define critical issues still unresolved by the “genome” or “proteome,” including the capacity of the same cytokine receptor to trigger opposing effects, the differential trafficking patterns of immune cells, and the ability of tumors or microbes to elicit divergent signaling events (2, 3). Studies on glycosylation have been hampered by the lack of straightforward approaches to study glycan structures in the basic “non-specialized” laboratory. However, these difficulties could be overcome in the past years by the identification of reliable and versatile strategies capable of profiling glycosylation changes; these include lectin cytometric analysis, a routine method which could be complemented by “glycan-gene” chip arrays and mass spectrometric analysis of glycan structures. Finally, definitive confirmation of the relevance of glycosylation in a given physiologic or pathologic settings may be achieved by the careful examination of mice transgenic or knockout for individual genes linked to the “glycosylation machinery” (namely glycosyltransferases, glycosidases, enzyme chaperons, or lectins) (more information is available at http://www.functionalglycomics.org). Abnormal glycosylation tightly correlates with the development of cancer and metastasis (4). These structural alterations are often the result of changes in the activity of one or more glycosyltransferases during the process of tumor transformation or metastasis (4). Notably, changes in the glycophenotype are also apparent in the tumor-associated stroma, endothelium, and infiltrating cells (2). Abnormal expression of glycosyltransferases or glycosidases can result in the modification of N-linked and O-linked glycans (Fig. 1). An example illustrating this concept is represented by N-glycan elongation which is strongly linked to an increased activity of N-acetylglucosaminiltransferase V (Mgat5) which leads to b1,6GlcNAc branching (5). This dynamic process also creates sites for incorporation of terminal sialic acid residues by sialyltransferases which are also upregulated during tumor growth (6). Programmed remodeling of tumor-associated N- and O-glycans can influence cell–cell and cell–matrix interactions, which results in dramatic changes in cell motility, invasiveness, and metastasis. In this regard, the metastatic potential of tumor cells has been extensively correlated with increased sialylation of cell surface glycoproteins, which is consistent with the known ability of sialic acid-binding lectins, such as selectins to mediate cell adhesion and extravasation during the metastatic process (7–10).

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Fig. 1. Protein–glycan interactions in the tumor microenvironment. Examples illustrating glycosylation changes associated with cancer progression are shown here. Increased activity of N-acetylglucosaminiltransferase V (Mgat5) promotes b1,6GlcNAc branching which creates new sites for incorporation of sialic acid residues. In addition, MUC-1 binds to the C-type lectin MGL which instructs DCs to drive TH2-mediated responses leading to anti-inflammatory responses. Galectins, a family of glycan-binding proteins, capable of recognizing multiple poly-LacNAc ligands, are secreted by tumor, stromal, and endothelial cells to modulate the survival and proliferation of effector T cells, skew the cytokine balance toward a TH2-type profile, modulate the physiology of APCs, and/or induce the differentiation, expansion, and/or recruitment of TReg cells to favor tumorimmune escape.

The aberrant activity of glycosyltransferases not only promotes alterations in the elongation and branching of glycans structures, but also favors the incorporation of particular terminal residues in the transformed cells. For example, several carbohydrate structures, such as Tn (GalNAc-a-Ser/Thr, CD175), sialyl Tn (N-acetylneuraminic acid-a6-GalNAc-a-Ser/Thr, CD175s), Thomsen–Friedenreich disaccharide (Gal-b1-3-GalNAc, CD176), and (sialylated) Lewis antigens (CD15s) are highly upregulated in malignant cells and have been broadly used as diagnostic and prognostic markers (8). Interestingly, glycophenotypic differences between normal and transformed cells have been exploited in the clinics, using monoclonal antibodies as probes, to detect circulating tumor metastatic cells (2). In addition, a common feature of the tumor cell microenvironment is the abundant production of mucin glycoproteins, which are distinguished by the prevalence of high density of O-linked glycans. Mucins are often found in neoplastic tissues and metastatic lesions, and have been proposed as potential prognostic markers of many tumor types (11). Interestingly, Tn glycans on MUC-1 bind the C-type lectin receptor MGL and instruct dendritic cells (DCs) to drive TH2mediated responses, which, unlike TH1, TH17, or CD8+ cytotoxic

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T cells, do not appear to contribute to tumor eradication. For that reason, the expression of Tn epitopes on MUC-1 is considered as a poor prognosis factor in several tumor types (4). In addition, transformed cells can also express a distinct pattern of gangliosides, sialic acid-containing glycosphingolipids which play critical roles in cell recognition and immunity. In this regard, complex gangliosides are often elevated in a plethora of tumors including small-cell lung carcinomas, neuroblastomas, and melanomas (12). Altogether these examples illustrate the relevance of altered glycosylation in the tumor microenvironment and suggest the potential development of novel therapeutic and diagnostic approaches based on the positive or negative regulation of protein–glycan interactions.

2. Biochemistry and Cell Biology of Galectins

The responsibility of deciphering the biological information encoded by the “glycome” is assigned, at least in part, to a large number of endogenous glycan-binding proteins or lectins, whose expression and function are regulated during tumor progression. This includes distinct families including C-type lectins (e.g., DC-SIGN, MGL, and selectins), siglecs, and galectins which are extremely divergent from either biochemical or functional standpoints (2). Galectins are evolutionarily conserved glycans-binding proteins with emerging roles in a wide variety of physiological and pathological processes (13, 14). To date, 15 galectins have been identified in mammals, with relative homologs widely distributed in the animal kingdom. Although some galectins (e.g., galectin-5, -10, and -12) are expressed with restricted tissue specificity, most of them have a wide tissue distribution (14). Galectins share a common structure and at least one conserved carbohydrate recognition domain (CRD) of approximately 130 amino acids that mediates carbohydrate binding. Traditionally, galectins are classified based on structural similarities in “proto-type” galectins (galectin-1, -2, -5, -7, -10, -11, 13, -14, and -15), which have one CRD and exist as monomers or dimers, “tandem repeattype” galectins (galectin-4, -6, -8, -9, and -12), which contain two different CRDs separated by a linker of up to 70 amino acids; and the “chimera-type” galectin-3, which contains one CRD connected to a nonlectin amino-terminal region (13, 14). With regard to their carbohydrate-binding activities, galectins are either bivalent or multivalent which allow the recognition of multiple binding partners and the activation of distinct signaling pathways. “Proto-type” galectins can dimerize, “tandem repeat-type” galectins are at least bivalent, and galectin-3 can form oligomers upon binding to multivalent glycoproteins (13, 14).

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In spite of the absence of a classical secretory signal in their primary sequence, most galectins are exported to the extracellular milieu through an unusual route that involves the glycanbinding activity of the secreted protein (14). Upon secretion, galectins can bind multiple glycosylated partners (glycoproteins or glycolipids) and can convey glycan-encoded information into immune cell activation, differentiation, and homeostatic programs (3, 15). How do galectins decode the biological information encrypted by glycan structures? Although much remains to be learned, research over the past years put forward the idea that galectins can transduce intracellular signals by forming ordered arrays of protein–glycan structures – termed lattices – on the surface of a variety of cell types. Yet, they may also function by engaging specific cell surface glycoconjugates and forming traditional ligand–receptor interactions (14, 16, 17). However, galectins are also active within the intracellular compartment through mechanisms that remain poorly understood (13, 14). Examples illustrating this concept are galectin-3 and galectin-10, which function intracellularly either to modulate cell survival and pre-mRNA splicing or to control the immunosuppressive activity of CD4+CD25+FoxP3+ human regulatory T (TReg) cells (14, 18). A general consensus exists in the notion that secreted galectins, in contrast to cytokines or chemokines, do not have specific receptors, but can mediate immune cell communication through the recognition of a preferred set of cell surface glycoconjugates (3, 14). In this context, the minimal structure recognized by galectins is the disaccharide N-acetyllactosamine (LacNAc), which is found in N- and O-glycans and can be presented as multiple units (poly-LacNAc) on cell surface glycoproteins (19). However, research over the past few years revealed substantial differences among the glycan-binding preferences of individual members of the galectin family (3, 19, 20), which represents the basis of functional divergences in their biological activity. These variations in glycan recognition are mainly associated with the extent of N-glycan branching, the multiplicity of LacNAc residues, and/or the modification of terminal saccharides including sialylation or fucosylation (19, 20). Interestingly, differences in carbohydrate recognition of individual galectins can be even more pronounced, as the specific binding of galectin-10 to mannose is of much more higher affinity than its binding to LacNAc or terminal galactose (14). In addition, selective binding of galectins to different glycoproteins can result from the particular spatial orientation of individual CRDs and the unique glycoprotein topologies determined by the number of attached N-glycans (16). Thus, in spite of their shared sequence homology and evolutionary conservation, galectins may exhibit diverse carbohydrate specificity and play divergent roles in biology.

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3. Galectin–Glycan Interactions in the Tumor Microenvironment

Although overlooked for a long time, the key importance of galectin–glycan interactions in cancer progression is now undisputed. In addition to glycophenotypic changes, gene and protein screening have repeatedly led to the identification of galectins as proteins that are up- or downregulated in neoplastic and metastatic lesions (21). Hence, given their abundant expression in tumor microenvironment, it is not surprising that the regulated expression of galectins may also contribute to delineate the “poor prognosis signature” (21). In fact, galectins can influence tumor progression through many different mechanisms, including the direct control of neoplastic transformation and/or the modulation of tumor cell survival, angiogenesis, and migration. In addition, galectins may also contribute to tumor growth by tilting the balance toward an immunosuppressive microenvironment that favors escape from T-cell-mediated immunity (21). Hence, a detailed analysis of the repertoire of galectins and their specific glycan partners may contribute to further understand the dialogue between tumor, stromal, and immune cells and delineate the potential role of protein–carbohydrate systems in cancer immunoediting. To date, galectins-1 and -3 are the most widely studied lectins with respect to their role in tumor progression (22). Galectin-1 modulates cancer progression by influencing cell–cell and cell– matrix interactions (23), by inducing apoptosis of effector T cells (24), or by contributing to tumor cell migration and angiogenesis (25, 26). Moreover, and in keeping with its immunosuppressive functions, research from our laboratory has identified a crucial role for galectin-1 in tumor cell evasion of immune response. Interestingly, blockade of galectin-1 expression in melanoma cells resulted in heightened T-cell-mediated tumor rejection, decreased frequency of apoptotic T cells, and increased secretion of TH1type cytokines (27). Moreover, Reed Sternberg cells in Hodgkin lymphoma selectively overexpressed galectin-1, which contributed to the immunosuppressive activity of these cells through induction of a TH2-type cytokine pattern, promotion of TReg cell expansion, and suppression of Epstein–Barr virus (EBV)-specific T-cell responses (28, 29). Furthermore, prostate cancer cells that had low expression of the core 2 N-acetylglucosaminyltransferase 1 (C2GnT1) were resistant to galectin-1-mediated cell death, although these cells expressed substantially higher amounts of this protein to selectively dampen effector T-cell responses (30). This immune inhibitory activity was further confirmed in human cancerous tissues, in which a strong inverse correlation was found between galectin-1 expression and the presence of tumor-infiltrating T cells in head and neck squamous cell carcinomas (31).

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This effect appears to be a common feature of distinct members of the galectin family as local delivery of galectin-3 efficiently promoted apoptosis of tumor-reactive CD8+ T cells and sustained tumor growth in a mouse model of colorectal cancer (32). Furthermore, galectin-3 expression correlated with apoptosis of tumor-associated lymphocytes in human melanoma biopsies (33). This was also true for galectin-9 as Klibi et al. recently reported the immunosuppressive effects of galectin-9-containing exosomes, which induced massive apoptosis in EBV-specific CD4+ T cells from patients with nasopharyngeal carcinoma (34). This effect was prevented using an anti-Tim-3 blocking antibody, consistent with the reported role of galectin-9 in selectively eliminating Tim-3+ T cells (35). In keeping with these findings, galectin-9 also modulates tumor immunity by facilitating Tim-3-dependent interactions between DCs and effector T cells (36). Hence, galectin– glycan interactions can influence immune tolerance in tumor microenvironments through the control of several mechanisms including the promotion of T-cell apoptosis, modulation of T-helper cytokine balance, regulation of DC physiology, and selective expansion of TReg cells. The availability of mice knock out for galectin genes as well as the possibility to manipulate galectin–glycan lattices in vivo has sparked a keen interest in studying galectin functions during tumor growth and metastasis. We will focus here on the cellular and molecular mechanisms underlying galectin-induced immunosuppression in vivo and the manipulation of galectin–glycan interactions for expanding and/ or improving existing anticancer therapies. 3.1. Galectin–Glycan Interactions in the Control of T-Cell Survival

Several members of the galectin family can bind to glycoprotein receptors on the surface of mature T cells (including CD45, CD43, CD2, CD3, and CD7) and trigger distinct signaling events which act in concert to regulate T-cell survival, thus promoting contraction and/or modulation of different immune cell compartments (3). In addition, galectins can also regulate the fate of other cells in the tumor microenvironment including cancerous and stromal cells (21). Galectin-1 can induce the upregulation of a- and b-chains of the IFN-g receptor on activated T lymphocytes, rendering these cells sensitive to IFN-g-induced apoptosis (37). Although the intracellular signaling pathways triggered by galectin-1 remain poorly understood, galectin-1-induced T-cell death has been shown to proceed through a caspase-independent pathway that involves rapid translocation of endonuclease G from mitochondria to the nucleus (38). However, galectin-1 was also shown to induce T-cell apoptosis through mechanisms involving sensitization to Fas (CD95) pathway and caspase-8 activation (39, 40). Moreover, exposure to galectin-1 triggers a disbalance of the Bcl-2/Bax ratio with a predominance of pro-apoptotic Bax and

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activation of the ERK-1/2 and AP-1 signaling pathways (41, 42). However, and in spite of these findings, other observations suggested that galectin-1 is not capable by itself to initiate a full death program, but instead induces phosphatidylserine exposure which prepares immune cells for phagocytic removal (43). Moreover, a recent report showed that galectin-1-induced T-cell death involves degradation of fodrin, a cytoskeletal adaptor that links CD45 to actin cytoskeleton (44). More importantly, the repertoire of Nand O-glycan structures expressed by activated or differentiated T cells (TH1, TH2, or TH17 cells), as well as the regulated expression and activity of particular glycosyltransferases are critical for galectin-1-induced T-cell death (45, 46). Supporting this notion, expression of the C2GnT1 can determine the susceptibility to galectin-1. This enzyme is responsible of creating and elongating the core 2 branch on O-glycans, thus allowing the incorporation of N-acetyllactosamine sequences, which are the preferred saccharide ligands of galectin-1 (47). On the other hand, the regulated expression and activity of the a2,6 sialyltransferase 1 (ST6Gal 1) can modify LacNAc ligands by the addition of sialic acid in a2-6-position of terminal galactose, which substantially blocks galectin-1 binding and abrogates galectin-1-induced cell death (45, 48). Therefore, susceptibility to galectin-1-induced T-cell death may be regulated at two distinct levels: (a) the presence of a restricted set of cell surface glycoproteins (e.g.,: CD43, CD45, or CD7) (49), whose segregation into membrane microdomains allows signaling events and activation of specific downstream effector molecules (49) and (b) the regulated expression of a set of glycosyltransferases responsible for creating or masking cell surface glycoconjugates (47, 48). Similar to galectin-1, galectin-2 binds to b1 integrins and triggers the death of activated T cells. This “proto-type” lectin triggers the activation of the intrinsic apoptotic pathway which involves caspases-3 and -9, cytochrome c release, disruption of the mitochondrial membrane potential, and increase in the Bax/ Bcl-2 ratio (50). Also, the “tandem-repeat” lectin galectin-9 can induce T-cell apoptosis, an effect which is prevented by enforced expression of the anti-apoptotic protein Bcl-2 (51) and occurs via the Ca2+-calpain-caspase-1 pathway (52). On the other hand, contrasting results have been reported for galectin-3 depending on its subcellular localization (14). Intracellular expression of galectin-3 has been mainly linked to an intrinsic anti-apoptotic effect of tumor and immune cells (14). In vivo evidence has been obtained through the analysis of galectin-3-deficient (Lgals3−/−) mice, whose peritoneal macrophages were much more prone to IFN-g- or LPS-induced apoptosis than with their wild-type counterpart (53). Within the tumor microenvironment, phosphorylated galectin-3 has been shown to protect BT549 human breast carcinoma cells from anoikis (a type of cell death elicited by the

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loss of cell anchorage) (54, 55). Interestingly, galectin-3 can display either anti- or pro-apoptotic activities against the same tumor necrosis factor-related (TRAIL) apoptotic stimuli depending on the target cell type. Overexpression of galectin-3 in J82 human bladder carcinoma cells rendered these cells resistant to TRAILinduced apoptosis, an effect which resulted from the activation of PI3K/Akt pathway (56), whereas galectin-3-transfected BT549 human breast carcinoma cells were paradoxically more sensitive to TRAIL-induced apoptosis through Akt inactivation (57). The mechanisms underlying these contrasting effects are still not clear. Similarly, the effects of galectin-3 on T-cell survival were found to be strongly dependent on its subcellular localization (14, 58). Exogenous galectin-3 can induce the formation of “lectin– glycoprotein lattices” which engage a T-cell apoptotic program by rising intracellular (Ca2+) levels, augmenting cytochrome c release, and promoting caspase-3 activation (58–60). On the contrary, intracellular expression of galectin-3 conferred resistance to apoptosis induced by a variety of agents including Fas ligand (CD95L) and chemotherapeutic agents through regulation of mitochondrial integrity and reduction of reactive oxygen species (ROS) (61–63). In fact, galectin-3-transfected Jurkat T cells survive significantly longer when treated with a pro-apoptotic antiCD95 (APO-1/Fas) monoclonal antibody (61). In this regard, two primary CD95-mediated apoptotic signaling routes have been described: (a) type I cells in which apoptosis is regulated through large amounts of caspase-8 activated by the death-inducing signaling complex (DISC) and (b) type II cells in which DISC and activated caspase 8 favor the apoptogenic activity of mitochondria through the release of cytochrome c and activation of caspase 3. Although type I cells express high amounts of galectin-3, type II cells appear to be negative for this protein. Notably, transfection with galectin-3 converted type II into type I apoptotic tumor cells, suggesting that galectin-3 can act directly at early signaling events of the CD95 pathway by promoting DISC formation and/or recruitment (64). An interesting observation is that galectin-3 translocates into the mitochondria when cells are exposed to apoptotic stimuli through a mechanism that involves binding to sinexin (62). Interestingly, a careful analysis of the galectin-3 primary sequence revealed the presence of four amino acids resembling the “anti-death-motif NWGR” (Asp-Trp-GlyArg) that is present in the BH1 domain of the Bcl-2 protein. This sequence is highly conserved among galectin-3 from different species and appears to be essential for its carbohydrate-binding activity (65). Even when galectin-3 was not capable of regulating the expression levels of any member of the Bcl-2 family (65), this lectin specifically interacted with Bcl-2 in a lactose-inhibitable fashion (61). Further studies are required to fully understand the way galectin-3 can differentially influence cell survival.

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Also galectin-7 is engaged in the regulation of cell fate. Galectin-7 gene (Lgals7) is an early transcriptional target of the tumor suppressor p53 (66). Galectin-7 is upregulated in UVBirradiated epidermal keratinocytes, and ectopic expression of galectin-7 made these cells more prone to undergo apoptosis compared with their normal counterpart (67). The pro-apoptotic properties of galectin-7 have been demonstrated using different cell types and distinct stimuli: a squamous cell line (67), HeLa cells, and the colon carcinoma cell line DLD-1 (68). Upon apoptosis induction, all galectin-7 transfectants displayed upregulation of c-Jun N-terminal kinase (JNK) activity, caspase-3 activation, and cytochrome c release (68). Remarkably, galectin-7-deficient keratinocytes were found to be protected from irradiation-induced apoptosis (69). Completing this picture, galectin-8 recently has been shown to modulate T-cell survival (70, 71) and even a galectin homologous isolated from liver chicken (CLL-I) can promote T-cell death (72), suggesting that galectin–glycan lattices are highly conserved molecular systems endowed with an intrinsic ability to regulate cell fate. 3.2. Galectins in the Control of T-Helper Cytokine Balance

Glycosylation can change dramatically not only in cancerous tissues, but also during physiological processes including immune cell activation, homing, and differentiation, resulting in the creation or masking of specific carbohydrate ligands for endogenous lectins (2, 73). A clear example illustrating this concept is the differential glycosylation of cell surface glycoproteins which can selectively control the survival of T-helper cells by modulating their susceptibility to galectin-1 (45). Although TH1 and TH17 differentiated cells express the repertoire of cell surface glycans that are required for galectin-1 binding and subsequent cell death, TH2 cells are protected from galectin-1 through a2-6 sialylation of cell surface glycoproteins (45). In keeping with this finding, galectin-1-deficient (Lgals1−/−) mice showed an increased frequency of TH1 and TH17 cells and enhanced the susceptibility to autoimmune neuroinflammation (45). These observations unveiled a molecular link among differential glycosylation of T-helper cells, susceptibility to cell death, and termination of the inflammatory response. Accordingly, recent studies showed that TH2 cells promote TH1 cell apoptosis through the secretion of galectin-1 (74), suggesting a galectin-1-dependent mechanism of counter-regulation between distinct T-helper subsets. In addition, galectin-1 can modulate the cytokine balance independently of its ability to modulate the lifespan of T cells. While exposure to galectin-1 downregulates IFN-g production, this glycan-binding protein favors the synthesis of T-cell-derived IL-10, IL-5, and TGF-b1 (75–78). Tim-3 (T-cell immunoglobulin mucin 3) was identified as a TH1-specific cell surface molecule that controls TH1 responses and regulates T-cell tolerance. In search for specific Tim-3 ligands,

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Zhu et al. identified galectin-9 as a specific binding partner capable of stimulating intracellular calcium flux, promoting aggregation, and inducing selective death of TH1 cells (35, 79). Interestingly, galectin-9 can also suppress the differentiation of TH17 cells both in vitro and in vivo independently of its ability to induce T-cell apoptosis (80). Given the established role of IFN-g-producing TH1 cells and the controversial activity of IL-17-producing TH17 cells in tumor growth, understanding the function of galectin–glycan lattices in the regulation of cytokine balance may contribute to define the hierarchical function of these cell subsets in controlling different stages of cancer immunoediting. 3.3. Galectins in T-Cell Signaling, Activation, and Anergy

Tumor cells use multiple tolerogenic strategies to subvert immune responses including inhibition of T-cell signaling and the promotion of T-cell anergy. Cell surface inhibitory receptors including CTLA-4, PD-1, and other molecules associated to immunoreceptor tyrosine-based inhibition motifs (ITIMs) play a crucial role in delivering negative signals that regulate the balance between T-cell activation, tolerance, and immunopathology (81). Although limited information is available on the role of galectin-1 in T-cell receptor (TCR)-mediated T-cell activation, this protein has been reported to modulate T-cell signaling at sites of immunological synapse (82). Liu et al. found that galectin-1 acts as an autocrine negative regulator of TCR binding, signal transduction, and burst size of CD8+ T cells (82). Moreover, Demetriou and colleagues provided elegant evidence demonstrating that galectin-3–Nglycan lattices can restrict spontaneous TCR clustering and downmodulate TCR responses by interacting with N-glycans modified by the enzyme N-acetylglucosaminyltransferase 5 (Mgat5). In this regard, galectin-3 co-localized with CD45 suppressed Lck activity and TCR signaling (83). These effects may have critical implications at the cross-roads of T-cell responsiveness and tolerance during tumor progression. In keeping with this notion, delivery of high doses of recombinant galectin-3 suppressed the activation of tumor-reactive T cells and promoted tumor growth in mice receiving tumor-reactive CD8+ T cells (32). In addition, at late stages of T-cell activation, galectin–glycan lattices can contribute to the termination of immune responses by promoting cell surface retention of the inhibitory molecule CTLA-4, thus favoring T-cell growth arrest (84). In this regard, a very elegant study highlighted an essential role for galectin-3– N-glycan interactions in mediating anergy of tumor-specific cytotoxic T lymphocytes by favoring the segregation of CD8 from the TCR (85), suggesting the possibility to bypass T-cell anergy by interfering with lectin-glycan lattices. Supporting a role for this protein in immune cell silencing, both galectin-1 and galectin-3 were found to be upregulated in anergic B cells (86).

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Therefore, galectin–glycan lattices may have evolved as endogenous homeostatic systems to prevent spontaneous T-cell activation and “turn-off ” T-cell effector functions after the completion of an immune response. In turn, these interactions may contribute to delineate the typical tolerogenic microenvironment usually found at sites of tumor growth and metastasis. 3.4. Galectins and the Function of TReg Cells

In the past years, a subset of TReg cells expressing CD4 and CD25 and the transcription factor FoxP3 have gained considerable attention and popularity as key regulators of T-cell tolerance and homeostasis (87). This population of T cells is specifically engaged in the maintenance of immune self-tolerance and the control of exuberant immune responses to foreign antigens. In addition, TReg cells have been proposed to be critical obstacles that hinder antitumor immunity and favor tumor–immune escape (88, 89). Investigation of gene and protein expression profiles has shown the upregulation of galectin-1 in human and mouse CD4+CD25+Foxp3+ TReg cells which substantially contributes to the immunosuppressive activity of these cells (90, 91). Investigation of the mechanisms underlying this inhibitory activity revealed a critical function of GM1 as a potential receptor for galectin-1 capable of mediating TRPC5 channel activation on effector T cells (92). Interestingly another “proto-type member” of the galectin family, galectin-10, was also identified as a marker of human CD4+CD25+FoxP3+ TReg cells which appeared to be essential for the suppressive activity of these cells (18). In addition to its upregulated expression in regulatory versus effector T cells, recent findings underscored the capacity of galectin-1 to increase the relative abundance and/or expansion of peripheral TReg cells in vivo (77). Administration of recombinant galectin-1 during the efferent phase of ocular inflammatory disease resulted in a remarkable increase of the immunosuppressive cytokines IL-10 and TGF-b1, which in turn promoted the expansion and/or activation of IL-10-producing TReg cells (77). This effect was also confirmed in a model of stress-induced fetal rejection, as injection of galectin-1 restored tolerance by promoting the expansion of IL-10-producing CD4+CD25+ TReg cells (93). Also, in Hodgkin lymphoma, tumor-derived galectin-1 could induce the differentiation of CD4+CD25+ TReg cells in vitro (28). This effect was not limited to galectin-1, as galectin-9 was also capable of modulating the TReg cell compartment. Exposure to galectin-9 in vitro induced the differentiation of FoxP3+ TReg cells, while simultaneously counteracted the generation TH17 pathogenic cells. The possible therapeutic benefits of these findings were evident in mouse model of collagen-induced arthritis (CIA) where galectin-9 administration ameliorated the arthritogenic process and induced decreased levels of the pro-inflammatory cytokines IL-17, IL-12, and IFN-g in the joint (80). In line with these

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findings, blockade of the Tim-3-galectin-9 pathway also resulted in substantial attenuation of the suppressive activity of TReg cells (94). Collectively, these findings highlight the potential role of galectin–glycan interactions in controlling TReg cell differentiation, expansion, and recruitment to the tumor microenvironments. 3.5. Galectins in the Control of AntigenPresenting Cells

In spite of their critical role in orchestrating adaptive immunity, bone marrow-derived antigen-presenting cells (APCs), particularly DCs, are now considered the pivotal cell type involved in the induction and maintenance of T-cell tolerance in vivo (95, 96). DCs can promote peripheral tolerance by promoting the differentiation of TReg cells, including CD4+CD25+FoxP3+ TReg cells and type-1 TReg (Tr1) cells (96, 97). Multiple factors can influence the decision of DCs to become tolerogenic, including the recognition of apoptotic cells (98), interaction with stromal cells (99), and exposure to an immunosuppressive tumor microenvironment (81). Furthermore, DCs modified by CD4+CD25+FoxP3+ TReg cells may become tolerogenic and drive the differentiation of IL-10-producing Tr1 cells (100), suggesting a link among distinct regulatory cell populations. Given the key role of DCs at the interface of innate and adaptive immunity, it is not surprising that lectin–glycan interactions may play an important role in regulating the biological activity of these cells. In this regard, we recently identified an essential function for galectin-1 in the generation of human and mouse tolerogenic DCs. DCs differentiated in the presence of galectin-1 acquired a distinctive regulatory profile, promoted T-cell tolerance in vivo, and terminated autoimmune neuroinflammation through an immunoregulatory circuit involving IL-27 and IL-10 (101). Exposure to galectin-1 during the maturation process induced the generation of DCs with a typical mature cell surface phenotype but dominant regulatory function. In addition, we have identified a pivotal role for endogenous galectin-1 in “fine tuning” the tolerogenic function of DCs (101). Consistent with these findings, progesterone-regulated galectin-1 could restore immune tolerance in failing pregnancies and this effect correlated with the expansion of TReg cells and the appearance of uterine cells with a regulatory DC phenotype (93). Given the plasticity of DCs, induction of a tolerogenic profile might be exploited therapeutically in order to attenuate autoimmune diseases or prevent graft rejection. On the contrary, silencing DC regulatory pathways might augment DC-cell-based vaccination efficiency or potentiate tumor immunotherapeutic strategies (81). In this regard, vaccination with tumor lysate-pulsed DCs holds a promise to treat immunogenic tumors (102). However, the protective function of DCs could be thwarted if these cells are rendered tolerogenic, as often occurs at the sites of tumor growth (81). Hence, it is the balance between immunogenic and tolerogenic signals that determines the effectiveness of immunotherapeutic strategies.

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Supporting this notion, we have recently demonstrated an in vivo neoplasic setting that tumor-pulsed DCs differentiated in the presence of galectin-1 (DCGal1) may become tolerogenic and fail to suppress tumor growth or elicit effective T-cell responses (101). Surprisingly, all mice immunized with tumor-pulsed DCGal1 developed progressively enlarging tumors when challenged with viable melanoma cells at a rate similar to that of mice receiving unpulsed DCs or vehicle control. Interestingly, co-injection of tumor-pulsed DCs and tumor-pulsed DCGal1 resulted in accelerated tumor, confirming a dominant tolerogenic effect of tumor-pulsed DCGal1, which prevented the protective effect of tumor-pulsed control DCs. Consistently, lymph node cells from mice receiving tumorpulsed DCGal1 or a mixture of tumor-pulsed DCs, and tumor-pulsed DCGal1 showed poor proliferative responses, reduced synthesis of IFN-g, and enhanced secretion of IL-10 (101). Thus, DCs differentiated in a galectin-1-enriched microenvironment cannot elicit an effective T-cell response against tumor challenge and instead skew the cytokine balance to foster a tolerant milieu at sites of tumor growth. Adding complexity to this system, recent findings suggested that galectin-1 can also regulate DC migration through modulation of Syk and protein kinase C (PKC) signaling (103). Thus, galectin-1 may control the motility of DCs, which upon arrival to sites of inflammation or tumor growth could be endowed with tolerogenic potential. While galectin-1 expression was found to be selectively upregulated by tolerogenic stimuli including IL-10, vitamin D3, and apoptotic cells (101), the expression of galectin-9 is peaked following exposure to maturation signals such as IFN-g and IL-1b (104). Accordingly, galectin-9 triggered the maturation of human monocyte-derived DCs through activation of the p38 MAPK pathway (105). In this regard, Tim-3, which is expressed at high levels on human and mouse DCs, has been identified as a candidate receptor for galectin-9 (106). Ligation of Tim-3 with galectin-9 synergized with Toll-like receptors (TLRs) initiate TH1-type immunity (106). Because Tim-3 cross-linking also dampens TH1 responses (35, 106), it has been speculated that galectin-9–Tim-3 interactions may have different effects during the initiation and termination of immune response. In this regard, recent findings supported a critical role for galectin-9 in potentiating tumorspecific T-cell responses through enhancement of Tim-3-mediated DC–CD8+ T-cell interactions (36). Finally, DCs from galectin-3deficient (Lgals3−/−) mice had decreased migratory potential, but instead secreted higher amounts of IL-12 and showed increased T-cell stimulatory capacity (107–109). Although these functions have not been studied in tumor settings, they strongly suggest an essential role for these lectins in fine-tuning DC physiology. Thus, galectin–glycan interactions may have evolved to regulate APC homeostasis and control their activation, signaling, and motility.

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4. Concluding Remarks Changes in glycosylation are a typical hallmark of pathological processes including inflammation, infection, autoimmunity, and cancer. These alterations can be of higher magnitude than those displayed by “the proteome” during cancer progression and metastasis. Galectins, a family of endogenous lectins found at sites of tumor growth and inflammation, can recognize and discriminate subtle changes on glycan structures displayed on the surface of tumor, stromal, and immune cells. Although overlooked for a long time, the key importance of galectin–glycan interactions in cancer progression is now undisputed. These interactions can lead to substantial changes in the malignant process including tumor cell adhesion, migration, angiogenesis, and immune escape. In this context, recent findings had shed light to an essential role of these lectins in the regulation of immune tolerance and inflammation. Galectin–glycan lattices can influence multiple tolerogenic mechanisms by modulating T-cell survival and signaling, controlling cytokine synthesis, promoting the differentiation and/or expansion of TReg cells, and “fine-tuning” DC physiology. Although much remains to be learned, it is likely that galectin– glycan lattices can regulate tumor immunoediting by bridging tumor, stromal, and immune cells. Due to the recent breakthrough of “proteomics” and “glycomics,” protein–glycan interactions have become more amenable of therapeutic approaches, suggesting novel anticancer strategies using small glycomimetic inhibitors, siRNA approaches, or galectin-specific blocking antibodies.

Acknowledgments We apologize to the many authors whose papers could not be cited owing to space limitations. We are grateful to D. Croci for helping in the design of figure. Work in authors’ laboratory is supported by grants from The Cancer Research Institute “Elaine R. Shephard” Award (USA), The Prostate Cancer Foundation (UK), Fundación Sales (Argentina), and Agencia Nacional de Promoción Científica y Tecnológica (FONCYT, PICT 2006-603; PICT 2007-903). References 1. Ohtsubo K and Marth JD. (2006) Glycosylation in cellular mechanisms of health and disease. Cell 126: 855–867. 2. van Kooyk Y and Rabinovich GA. (2008) Protein–glycan interactions in the control of

innate and adaptive immune responses. Nat Immunol 9: 593–601. 3. Rabinovich GA and Toscano MA. (2009) Turning ‘sweet’ on immunity: galectin– glycan interactions in immune tolerance

370

4.

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

Salatino and Rabinovich and inflammation. Nat Rev Immunol 9: 338–352. Hakomori S. (2002) Glycosylation defining cancer malignancy: new wine in an old bottle. Proc Natl Acad Sci USA 99: 10231–10233. Lau KS and Dennis JW. (2008) N-Glycans in cancer progression. Glycobiology 18: 750–760. Kim YJ and Varki A. (1997) Perspectives on the significance of altered glycosylation of glycoproteins in cancer. Glycoconj J 14: 569–576. Dube DH and Bertozzi CR. (2003) Metabolic oligosaccharide engineering as a tool for glycobiology. Curr Opin Chem Biol 7: 616–625. Dube DH and Bertozzi CR. (2005) Glycans in cancer and inflammation – potential for therapeutics and diagnostics. Nat Rev Drug Discov 4: 477–488. Keppler OT, Horstkorte R, Pawlita M, et al. (2001) Biochemical engineering of the N-acyl side chain of sialic acid: biological implications. Glycobiology 11: 11R–18R. Prescher JA, Dube DH and Bertozzi CR. (2004) Chemical remodelling of cell surfaces in living animals. Nature 430: 873–877. Hollingsworth MA and Swanson BJ. (2004) Mucins in cancer: protection and control of the cell surface. Nat Rev Cancer 4: 45–60. Hakomori S. (2000) Traveling for the glycosphingolipid path. Glycoconj J 17: 627–647. Rabinovich GA, Toscano MA, Jackson SS, et al. (2007) Functions of cell surface galectin-glycoprotein lattices. Curr Opin Struct Biol 17: 513–520. Yang RY, Rabinovich GA and Liu FT. (2008) Galectins: structure, function and therapeutic potential. Expert Rev Mol Med 10: e17. Rabinovich GA and Ilarregui JM. (2009) Conveying glycan information into T-cell homeostatic programs: a challenging role for galectin-1 in inflammatory and tumor microenvironments. Immunol Rev 230: 144–159. Brewer CF, Miceli MC and Baum LG. (2002) Clusters, bundles, arrays and lattices: novel mechanisms for lectin-saccharidemediated cellular interactions. Curr Opin Struct Biol 12: 616–623. Laderach DJ, Compagno D, Toscano MA, et al. (2010) Dissecting the signal transduction pathways triggered by galectin–glycan interactions in physiological and pathological settings. IUBMB Life 62(1):1–13.

18. Kubach J, Lutter P, Bopp T, et al. (2007) Human CD4+CD25+ regulatory T cells: proteome analysis identifies galectin-10 as a novel marker essential for their anergy and suppressive function. Blood 110:1550–1558. 19. Hirabayashi J, Hashidate T, Arata Y, et al. (2002) Oligosaccharide specificity of galectins: a search by frontal affinity chromatography. Biochim Biophys Acta 1572: 232–254. 20. Stowell SR, Arthur CM, Mehta P, et al. (2008) Galectin-1, -2, and -3 exhibit differential recognition of sialylated glycans and blood group antigens. J Biol Chem 283: 10109–10123. 21. Liu FT and Rabinovich GA. (2005) Galectins as modulators of tumour progression. Nat Rev Cancer 5: 29–41. 22. Salatino M, Croci DO, Bianco GA, et al. (2008) Galectin-1 as a potential therapeutic target in autoimmune disorders and cancer. Expert Opin Biol Ther 8: 45–57. 23. Elola MT, Wolfenstein-Todel C, Troncoso MF, et al. (2007) Galectins: matricellular glycan-binding proteins linking cell adhesion, migration, and survival. Cell Mol Life Sci 64: 1679–1700. 24. Perillo NL, Pace KE, Seilhamer JJ, et al. (1995) Apoptosis of T cells mediated by galectin-1. Nature 378: 736–739. 25. Thijssen VL, Postel R, Brandwijk RJ, et al. (2006) Galectin-1 is essential in tumor angiogenesis and is a target for antiangiogenesis therapy. Proc Natl Acad Sci U S A 103: 15975–15980. 26. Le Mercier M, Fortin S, Mathieu V, et al. (2009) Galectin 1 proangiogenic and promigratory effects in the Hs683 oligodendroglioma model are partly mediated through the control of BEX2 expression. Neoplasia 11: 485–496. 27. Rubinstein N, Alvarez M, Zwirner NW, et al. (2004) Targeted inhibition of galectin-1 gene expression in tumor cells results in heightened T cell-mediated rejection; a potential mechanism of tumor-immune privilege. Cancer Cell 5: 241–251. 28. Juszczynski P, Ouyang J, Monti S, et al. (2007) The AP1-dependent secretion of galectin-1 by Reed Sternberg cells fosters immune privilege in classical Hodgkin lymphoma. Proc Natl Acad Sci U S A 104: 13134–13139. 29. Gandhi MK, Moll G, Smith C, et al. (2007) Galectin-1 mediated suppression of Epstein– Barr virus specific T-cell immunity in classic Hodgkin lymphoma. Blood 110: 1326–1329. 30. Valenzuela HF, Pace KE, Cabrera PV, et al. (2007) O-glycosylation regulates LNCaP

Galectin–Glycan Interactions in Tumor Immunity

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

prostate cancer cell susceptibility to apoptosis induced by galectin-1. Cancer Res 67: 6155–6162. Le QT, Shi G, Cao H, et al. (2005) Galectin-1: a link between tumor hypoxia and tumor immune privilege. J Clin Oncol 23: 8932–8941. Peng W, Wang HY, Miyahara Y, et al. (2008) Tumor-associated galectin-3 modulates the function of tumor-reactive T cells. Cancer Res 68: 7228–7236. Zubieta MR, Furman D, Barrio M, et al. (2006) Galectin-3 expression correlates with apoptosis of tumor-associated lymphocytes in human melanoma biopsies. Am J Pathol 168: 1666–1675. Klibi J, Niki T, Riedel A, et al. (2009) Blood diffusion and Th1-suppressive effects of galectin-9-containing exosomes released by Epstein–Barr virus-infected nasopharyngeal carcinoma cells. Blood 113: 1957–1966. Zhu C, Anderson AC, Schubart A, et al. (2005) The Tim-3 ligand galectin-9 negatively regulates T helper type 1 immunity. Nat Immunol 6: 1245–1252. Nagahara K, Arikawa T, Oomizu S, et al. (2008) Galectin-9 increases Tim-3+ dendritic cells and CD8+ T cells and enhances antitumor immunity via galectin-9-Tim-3 interactions. J Immunol 181: 7660–7669. Allione A, Wells V, Forni G, et al. (1998) Beta-galactoside-binding protein (beta GBP) alters the cell cycle, up-regulates expression of the alpha- and beta-chains of the IFN-gamma receptor, and triggers IFNgamma-mediated apoptosis of activated human T lymphocytes. J Immunol 161: 2114–2119. Hahn HP, Pang M, He J, et al. (2004) Galectin-1 induces nuclear translocation of endonuclease G in caspase- and cytochrome c-independent T cell death. Cell Death Differ 11: 1277–1286. Brandt B, Buchse T, Abou-Eladab EF, et al. (2008) Galectin-1 induced activation of the apoptotic death-receptor pathway in human Jurkat T lymphocytes. Histochem Cell Biol 129: 599–609. Matarrese P, Tinari A, Mormone E, et al. (2005) Galectin-1 sensitizes resting human T lymphocytes to Fas (CD95)-mediated cell death via mitochondrial hyperpolarization, budding, and fission. J Biol Chem 280: 6969–6985. Vespa GN, Lewis LA, Kozak KR, et al. (1999) Galectin-1 specifically modulates TCR signals to enhance TCR apoptosis but

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

371

inhibit IL-2 production and proliferation. J Immunol 162: 799–806. Rabinovich GA, Alonso CR, Sotomayor CE, et al. (2000) Molecular mechanisms implicated in galectin-1-induced apoptosis: activation of the AP-1 transcription factor and downregulation of Bcl-2. Cell Death Differ 7: 747–753. Stowell SR, Karmakar S, Arthur CM, et al. (2009) Galectin-1 induces reversible phosphatidylserine exposure at plasma membrane. Mol Biol Cell 20: 1408–1418. Pang M, He J, Johnson P, et al. (2009) CD45-mediated fodrin cleavage during galectin-1 T cell death promotes phagocytic clearance of dying cells. J Immunol 182: 7001–7008. Toscano MA, Bianco GA, Ilarregui JM, et al. (2007) Differential glycosylation of TH1, TH2 and TH-17 effector cells selectively regulates susceptibility to cell death. Nat Immunol 8: 825–834. Earl LA, Bi S and Baum LG. (2010) N- and O-glycans modulate galectin-1 binding, CD45 signaling and T cell death. J Biol Chem 285: 2232–2244. Galvan M, Tsuboi S, Fukuda M, et al. (2000) Expression of a specific glycosyltransferase enzyme regulates T cell death mediated by galectin-1. J Biol Chem 275: 16730–16737. Amano M, Galvan M, He J, et al. (2003) The ST6Gal I sialyltransferase selectively modifies N-glycans on CD45 to negatively regulate galectin-1-induced CD45 clustering, phosphatase modulation, and T cell death. J Biol Chem 278: 7469–7475. Pace KE, Lee C, Stewart PL, et al. (1999) Restricted receptor segregation into membrane microdomains occurs on human T cells during apoptosis induced by galectin-1. J Immunol 163: 3801–3811. Sturm A, Lensch M, Andre S, et al. (2004) Human galectin-2: novel inducer of T cell apoptosis with distinct profile of caspase activation. J Immunol 173: 3825–3837. Bi S, Earl LA, Jacobs L, et al. (2008) Structural features of galectin-9 and galectin-1 that determine distinct T cell death pathways. J Biol Chem 283: 12248–12258. Kashio Y, Nakamura K, Abedin MJ, et al. (2003) Galectin-9 induces apoptosis through the calcium-calpain-caspase-1 pathway. J Immunol 170: 3631–3636. Hsu DK, Yang RY, Pan Z, et al. (2000) Targeted disruption of the galectin-3 gene results in attenuated peritoneal inflammatory responses. Am J Pathol 156: 1073–1083.

372

Salatino and Rabinovich

54. Yoshii T, Fukumori T, Honjo Y, et al. (2002) Galectin-3 phosphorylation is required for its anti-apoptotic function and cell cycle arrest. J Biol Chem 277: 6852–6857. 55. Kim HR, Lin HM, Biliran H, et al. (1999) Cell cycle arrest and inhibition of anoikis by galectin-3 in human breast epithelial cells. Cancer Res 59: 4148–4154. 56. Oka N, Nakahara S, Takenaka Y, et al. (2005) Galectin-3 inhibits tumor necrosis factorrelated apoptosis-inducing ligand-induced apoptosis by activating Akt in human bladder carcinoma cells. Cancer Res 65: 7546–7553. 57. Lee YJ, Song YK, Song JJ, et al. (2003) Reconstitution of galectin-3 alters glutathione content and potentiates TRAIL-induced cytotoxicity by dephosphorylation of Akt. Exp Cell Res 288: 21–34. 58. Stillman BN, Hsu DK, Pang M, et al. (2006) Galectin-3 and galectin-1 bind distinct cell surface glycoprotein receptors to induce T cell death. J Immunol 176: 778–789. 59. Dong S and Hughes RC. (1996) Galectin-3 stimulates uptake of extracellular Ca2+ in human Jurkat T-cells. FEBS Lett 395: 165–169. 60. Fukumori T, Takenaka Y, Yoshii T, et al. (2003) CD29 and CD7 mediate galectin-3induced type II T-cell apoptosis. Cancer Res 63: 8302–8311. 61. Yang RY, Hsu DK and Liu FT. (1996) Expression of galectin-3 modulates T-cell growth and apoptosis. Proc Natl Acad Sci U S A 93: 6737–6742. 62. Yu F, Finley RL, Jr, Raz A, et al. (2002) Galectin-3 translocates to the perinuclear membranes and inhibits cytochrome c release from the mitochondria. A role for synexin in galectin-3 translocation. J Biol Chem 277: 15819–15827. 63. Matarrese P, Fusco O, Tinari N, et al. (2000) Galectin-3 overexpression protects from apoptosis by improving cell adhesion properties. Int J Cancer 85: 545–554. 64. Fukumori T, Takenaka Y, Oka N, et al. (2004) Endogenous galectin-3 determines the routing of CD95 apoptotic signaling pathways. Cancer Res 64: 3376–3379. 65. Akahani S, Nangia-Makker P, Inohara H, et al. (1997) Galectin-3: a novel antiapoptotic molecule with a functional BH1 (NWGR) domain of Bcl-2 family. Cancer Res 57: 5272–5276. 66. Polyak K, Xia Y, Zweier JL, et al. (1997) A model for p53-induced apoptosis. Nature 389: 300–305. 67. Bernerd F, Sarasin A and Magnaldo T. (1999) Galectin-7 overexpression is associated with

68.

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

the apoptotic process in UVB-induced sunburn keratinocytes. Proc Natl Acad Sci USA 96: 11329–11334. Kuwabara I, Kuwabara Y, Yang RY, et al. (2002) Galectin-7 (PIG1) exhibits proapoptotic function through JNK activation and mitochondrial cytochrome c release. J Biol Chem 277: 3487–3497. Gendronneau G, Sidhu SS, Delacour D, et al. (2008) Galectin-7 in the control of epidermal homeostasis after injury. Mol Biol Cell 19: 5541–5549. Tribulatti MV, Mucci J, Cattaneo V, et al. (2007) Galectin-8 induces apoptosis in the CD4(high)CD8(high) thymocyte subpopulation. Glycobiology 17: 1404–1412. Norambuena A, Metz C, Vicuna L, et al. (2009) Galectin-8 induces apoptosis in Jurkat T cells by phosphatidic acid-mediated ERK1/2 activation supported by protein kinase A down-regulation. J Biol Chem 284: 12670–12679. Rabinovich GA, Modesti NM, Castagna LF, et al. (1997) Specific inhibition of lymphocyte proliferation and induction of apoptosis by CLL-I, a beta-galactoside-binding lectin. J Biochem 122: 365–373. Marth JD and Grewal PK. (2008) Mammalian glycosylation in immunity. Nat Rev Immunol 8: 874–887. Motran CC, Molinder KM, Liu SD, et al. (2008) Galectin-1 functions as a Th2 cytokine that selectively induces Th1 apoptosis and promotes Th2 function. Eur J Immunol 38: 3015–3027. Rabinovich GA, Ariel A, Hershkoviz R, et al. (1999) Specific inhibition of T-cell adhesion to extracellular matrix and proinflammatory cytokine secretion by human recombinant galectin-1. Immunology 97: 100–106. Rabinovich GA, Daly G, Dreja H, et al. (1999) Recombinant galectin-1 and its genetic delivery suppress collagen-induced arthritis via T cell apoptosis. J Exp Med 190: 385–398. Toscano MA, Commodaro AG, Ilarregui JM, et al. (2006) Galectin-1 suppresses autoimmune retinal disease by promoting concomitant Th2- and T regulatory-mediated anti-inflammatory responses. J Immunol 176: 6323–6332. Stowell SR, Qian Y, Karmakar S, et al. (2008) Differential roles of galectin-1 and galectin-3 in regulating leukocyte viability and cytokine secretion. J Immunol 180: 3091–3102. Niwa H, Satoh T, Matsushima Y, et al. (2009) Stable form of galectin-9, a Tim-3 ligand, inhibits contact hypersensitivity and

Galectin–Glycan Interactions in Tumor Immunity

80.

81.

82.

83.

84.

85.

86.

87.

88.

89.

90.

91.

92.

psoriatic reactions: a potent therapeutic tool for Th1 and Th17-mediated skin inflammation. Clin Immunol 132: 184–194. Seki M, Oomizu S, Sakata KM, et al. (2008) Galectin-9 suppresses the generation of Th17, promotes the induction of regulatory T cells, and regulates experimental autoimmune arthritis. Clin Immunol 127: 78–88. Rabinovich GA, Gabrilovich D and Sotomayor EM. (2007) Immunosuppressive strategies that are mediated by tumor cells. Annu Rev Immunol 25: 267–296. Liu SD, Tomassian T, Bruhn KW, et al. (2009) Galectin-1 tunes TCR binding and signal transduction to regulate CD8 burst size. J Immunol 182: 5283–5295. Grigorian A, Torossian S and Demetriou M. (2009) T-cell growth, cell surface organization, and the galectin-glycoprotein lattice. Immunol Rev 230: 232–246. Lau KS, Partridge EA, Grigorian A, et al. (2007) Complex N-glycan number and degree of branching cooperate to regulate cell proliferation and differentiation. Cell 129: 123–134. Demotte N, Stroobant V, Courtoy PJ, et al. (2008) Restoring the association of the T cell receptor with CD8 reverses anergy in human tumor-infiltrating lymphocytes. Immunity 28: 414–424. Clark AG, Chen S, Zhang H, et al. (2007) Multifunctional regulators of cell growth are differentially expressed in anergic murine B cells. Mol Immunol 44: 1274–1285. Sakaguchi S. (2004) Naturally arising CD4+ regulatory t cells for immunologic self-tolerance and negative control of immune responses. Annu Rev Immunol 22: 531–562. Curiel TJ, Coukos G, Zou L, et al. (2004) Specific recruitment of regulatory T cells in ovarian carcinoma fosters immune privilege and predicts reduced survival. Nat Med 10: 942–949. Zou W. (2006) Regulatory T cells, tumour immunity and immunotherapy. Nat Rev Immunol 6: 295–307. Garin MI, Chu CC, Golshayan D, et al. (2007) Galectin-1: a key effector of regulation mediated by CD4+CD25+ T cells. Blood 109: 2058–2065. Sugimoto N, Oida T, Hirota K, et al. (2006) Foxp3-dependent and -independent molecules specific for CD25+CD4+ natural regulatory T cells revealed by DNA microarray analysis. Int Immunol 18: 1197–1209. Wang J, Lu ZH, Gabius HJ, et al. (2009) Cross-linking of GM1 ganglioside by galectin-1 mediates regulatory T cell activity

93.

94.

95.

96.

97. 98.

99.

100.

101.

102.

103.

104.

105.

106.

373

involving TRPC5 channel activation: possible role in suppressing experimental autoimmune encephalomyelitis. J Immunol 182: 4036–4045. Blois SM, Ilarregui JM, Tometten M, et al. (2007) A pivotal role for galectin-1 in fetomaternal tolerance. Nat Med 13: 1450–1457. Wang F, Wan L, Zhang C, et al. (2009) Tim3-Galectin-9 pathway involves the suppression induced by CD4+CD25+ regulatory T cells. Immunobiology 214: 342–349. Guermonprez P, Valladeau J, Zitvogel L, et al. (2002) Antigen presentation and T cell stimulation by dendritic cells. Annu Rev Immunol 20: 621–667. Steinman RM, Hawiger D and Nussenzweig MC. (2003) Tolerogenic dendritic cells. Annu Rev Immunol 21: 685–711. Reis e Sousa C. (2006) Dendritic cells in a mature age. Nat Rev Immunol 6: 476–483. Skoberne M, Beignon AS and Bhardwaj N. (2004) Danger signals: a time and space continuum. Trends Mol Med 10: 251–257. Svensson M, Maroof A, Ato M, et al. (2004) Stromal cells direct local differentiation of regulatory dendritic cells. Immunity 21: 805–816. Awasthi A, Carrier Y, Peron JP, et al. (2007) A dominant function for interleukin 27 in generating interleukin 10-producing antiinflammatory T cells. Nat Immunol 8: 1380–1389. Ilarregui JM, Croci DO, Bianco GA, et al. (2009) Tolerogenic signals delivered by dendritic cells to T cells through a galectin-1driven immunoregulatory circuit involving interleukin 27 and interleukin 10. Nat Immunol 10: 981–991. Kotera Y, Shimizu K and Mule JJ. (2001) Comparative analysis of necrotic and apoptotic tumor cells as a source of antigen(s) in dendritic cell-based immunization. Cancer Res 61: 8105–8109. Fulcher JA, Chang MH, Wang S, et al. (2009) Galectin-1 co-clusters CD43/CD45 on dendritic cells and induces cell activation and migration through Syk and protein kinase C signaling. J Biol Chem 284: 26860–26870. Hirashima M, Kashio Y, Nishi N, et al. (2009) Galectin-9 in physiological and pathological conditions. Glycoconj J 19: 593–600. Dai SY, Nakagawa R, Itoh A, et al. (2005) Galectin-9 induces maturation of human monocyte-derived dendritic cells. J Immunol 175: 2974–2981. Anderson AC, Anderson DE, Bregoli L, et al. (2007) Promotion of tissue inflammation by the immune receptor Tim-3 expressed on innate immune cells. Science 318: 1141–1143.

374

Salatino and Rabinovich

107. Hsu DK, Chernyavsky AI, Chen HY, et al. (2009) Endogenous galectin-3 is localized in membrane lipid rafts and regulates migration of dendritic cells. J Invest Dermatol 129: 573–583. 108. Bernardes ES, Silva NM, Ruas LP, et al. (2006) Toxoplasma gondii infection reveals a novel regulatory role for galectin-3 in the

interface of innate and adaptive immunity. Am J Pathol 168: 1910–1920. 109. Breuilh L, Vanhoutte F, Fontaine J, et al. (2007) Galectin-3 modulates immune and inflammatory responses during helminthic infection: impact of galectin-3 deficiency on the functions of dendritic cells. Infect Immun 75: 5148–5157.

Relevant Websites 110. Consortium for Functional Glycomics (http://www.functionalglycomics.org).

111. The Japanese Consortium for Glycobiology and Glycotechnology (http://www.jcgg.jp).

Chapter 24 Regulation of Lymphocytes by Nitric Oxide Christian Bogdan Abstract Shortly after the identification of nitric oxide (NO) as a product of macrophages, it was discovered that NO generated by inducible NO synthase (iNOS) inhibits the proliferation of T lymphocytes. Since then, it has become clear that iNOS activity also regulates the development, differentiation, and/or function of various types of T cells and B cells and also affects NK cells. The three key mechanisms underlying the iNOS-dependent immunoregulation are (a) the modulation of signaling processes by NO, (b) the depletion of arginine, and (c) the alteration of accessory cell functions. This chapter highlights important principles of iNOS-dependent immunoregulation of lymphocytes and also reviews more recent evidence for an effect of endothelial or neuronal NO synthase in lymphocytes. Key words: Nitric oxide, Inducible nitric oxide synthase, Endothelial nitric oxide synthase, Neuronal nitric oxide synthase, Arginase, Myeloid (-derived) suppressor cells, T lymphocytes, B lymphocytes, Natural killer cells

1. Introduction A protective immune response to viruses, bacteria, and other infectious pathogens requires not only efficient modes of induction but also an adequate restriction of its magnitude and duration to avoid chronic inflammatory events and autoimmune reactions. The immune system, therefore, maintains a complex network of cellular and humoral components that ideally help to prevent overshooting responses and tissue damage and to resolve inflammations. However, during certain infectious diseases, these counterregulatory, immunosuppressive mechanisms might be hyperinduced by the pathogen itself leading to its persistence and to chronic infections. A similar detrimental dysbalance can be observed during some malignancies, where the highly desirable cytotoxic anti-tumor response is partially shut down by tumor-derived metabolites or Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_24, © Springer Science+Business Media, LLC 2011

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tumor-induced products of the immune system that ultimately facilitate the growth and metastasis of the transformed cells. In both situations, various types of regulatory immune cells that dampen the immune reaction operate. These include different subsets of regulatory T cells (e.g., interleukin [IL]-10-expressing type 1 T-helper cells [Th1], inducible forkhead-box-protein P3 [FoxP3]positive regulatory T cells [iTreg], FoxP3+ natural regulatory T cells [nTreg]) [1], regulatory or tolerogenic dendritic cells [2–6], suppressor monocytes and macrophages [7, 8], myeloid suppressor cells [9, 10], or IL-10-expressing neutrophils [11]. Numerous cell-bound or secreted products are known that contribute to the suppressive phenotype of these cells, e.g., galectins [12], CTLA4 [1], IL-10 [13, 14], transforming growth factor-b [1, 15], prostaglandins [16], lipoxins [17], adenosine [18], kynurenines (derived from indoleamine-2,3-dioxygenase) [1, 19], carbon monoxide (derived from heme oxygenase) [20], arginase [9, 21], and nitric oxide (NO) (derived from inducible NO synthase [iNOS, NOS2]) [22–24]. The most versatile molecule of these products in the immune system is NO, which is mainly due to its widespread production, prominent diffusion capacity, high reactivity, and its multidirectional effects on immune cells [25]. This chapter focuses on the regulatory effects of NO on T cells and other lymphocytes. For an in-depth review of the expression, function, and regulation of NO in the entire immune system, the reader is referred to earlier articles [22, 23, 26–28].

2. NO Synthases Mammalian NO synthases (NOS) are flavoprotein-containing oxidoreductases that convert the aminoacid l-arginine and molecular oxygen into citrulline and nitric oxide radical (in the following abbreviated as NO). All NOS isoforms function as homodimers and require the binding of calmodulin, the incorporation of iron protoporphyrin IX (heme) and zinc, and the presence of multiple cofactors and cosubstrates [(6R)-tetrahydrobiopterin (BH4), NADPH, flavin-adenine mononucleotide (FMN), flavin-adenine dinucleotide (FAD), thiol] [29]. Two of the NOS isoforms – the neuronal NOS (nNOS or NOS1) and the endothelial NOS (eNOS or NOS3) – are constitutively expressed enzymes that are already found in strictly resting cells and tissues and subject to regulation by Ca2+ fluxes, which led to their joint name cNOS. They release small amounts of NO upon activation by Ca2+dependent binding of calmodulin. nNOS and eNOS were originally discovered in the central nervous system or cardiovascular system, respectively, where they mainly fulfill homeostatic

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functions, e.g., in the context of neurotransmission or blood pressure regulation [30–33]. However, more recently published studies indicate that both nNOS- and eNOS-derived NO exert regulatory effects during infections and autoimmune or other inflammatory processes [34–36] (see also below). The third isoform of NOS (iNOS or NOS2) was originally discovered in activated macrophages (therefore, sometimes also termed macNOS) and is characterized by the de novo expression in response to interferon (IFN)-g and microbial ligands of pattern recognition receptors (PRR), the production of high amounts of NO, and a predominant role in the immune system compared to other organ systems. iNOS expression is regulated at multiple levels, ranging from the transcriptional induction and mRNA stability, the synthesis, stability, and degradation of the iNOS protein, to the activity of the iNOS enzyme [23, 28, 37]. Key signaling cascades that govern the transcription and mRNA expression of iNOS include the Janus kinase (Jak1 and Tyk2)/signal transducer and activator of transcription (STAT1/2)/ interferon-regulatory factor (IRF1)-pathway, which is triggered by IFN-a/b or IFN-g, as well as the mitogen-activated protein kinases (MAPK) and the nuclear factor (NF) kB, which are activated by tumor necrosis factor (TNF) or pathogen-associated molecular patterns (PAMP) [e.g., bacterial lipopolysaccharide (LPS), fungal zymosan, and microbial nucleic acids] [23, 28]. The expression of iNOS protein in primary macrophages is controlled by proteasomes, which are targets of the iNOS-suppressive transforming growth factor-b1 [38–41]. In addition, iNOS is regulated by the availability of its substrate (l-arginine), which not only affects iNOS enzyme activity, but also the translation of iNOS mRNA [42, 43]. In this context, the enzyme arginase is of particular relevance because it converts l-arginine into ornithine and urea and, therefore, is capable to deplete exogenous arginine supplies if activated prior to the induction of iNOS. Arginase exists in two isoforms (cytosolic “hepatic” arginase 1 and mitochondrial “extrahepatic” arginase 2). Both isoforms are expressed in macrophages, but only arginase 1 is strongly upregulated by IL-3, IL-4, IL-10, IL-13, and TGF-b in macrophages and therefore accounts at least partially for the iNOS-suppressive effects of these cytokines [23, 42–48]. While iNOS-dependent NO production was originally described as a host-protective cytotoxic mechanism against infectious pathogens and tumors, it is now clear that iNOS-derived NO and its congeners [collectively termed reactive nitrogen intermediates (RNI)] also function as immunoregulatory or as tissue-damaging molecules during infections, malignancies, transplantations, and autoimmune diseases (for review see [22, 23, 27, 28, 49]).

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3. NO Producers While nNOS is expressed in different types of neurons and eNOS is found in various types of endothelial cells and in vascular smooth muscle cells, the prototype of an iNOS-positive cell is the macrophage activated by cytokines and/or microbial products [22]. This expression pattern was relevant for the initial naming of the NOS isoforms, but no longer reflects the complexity of the distribution of NOS in mammalian organisms. In addition to classical bone marrow-derived or peritoneal macrophages, iNOS can be readily induced in a broad spectrum of other myeloid cells [22], e.g., in granulocytes, osteoclasts, bone marrow-derived conventional myeloid dendritic cells (cDC; CD11b+CD11chigh)[5, 50–52], in tumor necrosis factor/iNOS-positive splenic DCs (TIP-DC; CD11b+CD11cintGr-1+B220−) [53, 54], in granulocytic (CD11b+ CD11c−Ly6G+Ly6ClowCD31+) or monocytic (CD11b+CD11c−Ly 6G−Ly6ChighCD31+) myeloid-derived suppressor cells (MSC or MDSC; human MDSC: CD11b+CD14−CD33+)[9, 10, 55–57], in microglia [22, 58], and in mast cells [59]. iNOS will be also expressed in cytokine-stimulated hepatocytes, endothelial cells, epithelial cells, keratinocytes, and fibroblasts [22] and has been detected in different types of neuronal cells in the central nervous system during inflammatory responses [60]. Whether T lymphocytes can be activated for the expression of iNOS mRNA and protein has been repeatedly addressed, but yielded conflicting results [23, 61, 62]. nNOS and/or eNOS mRNA or protein have also been reported to be expressed in certain myeloid and lymphoid primary cells and cell lines (for review see [22, 23, 63–65]) (see also Subheading 4.2).

4. NO Targets 4.1. Exogenous iNOS/ NO and T Lymphocytes 4.1.1. T-Cell Differentiation

NO has been implicated in the differentiation of various T-cell populations. Intermediate concentrations of an NO donor [S-nitroso-N-acetyl-penicillamine (SNAP); 1–10 mM] enhanced the development of IFN-g producing CD4+ T cells (Th1 cells) after antigen-specific or polyclonal stimulation in the presence of IL-12 and anti-IL-4, whereas an IL-4-driven Th2 cell line remained unaffected [66]. In accordance with these findings, exogenous NO (generated by the NO-donor NOC-18) upregulated – via activation of soluble guanylyl cyclase – the mRNA expression of the IL-12Rb2 chain, whereas the IL4Ra mRNA remained unaffected [67].

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Recently, NO (NOC-18) was reported to induce a population of regulatory T cells (termed NO-Treg), which carried a characteristic set of markers (CD25+, CD27+, Foxp3−, GITR+, T-betlow, and GATA3high), but were functionally similar to natural Treg (i.e., inhibition of CD4+CD25− T-cell proliferation in vitro, attenuation of colitis, and collagen-induced arthritis in vivo) [68]. Another study demonstrated that the production of NO by IFNg-activated DCs promoted the development of Foxp3+ Treg [69]. The NO-dependent induction of Foxp3− or Foxp3+ Treg could be one of the mechanisms, by which IFN-g helps to protect the organism against exaggerated immune responses and immunopathology. However, the relationship between NO and regulatory T cells appears to be considerably more complex. First, in tumorbearing mice wildtype and iNOS-deficient MDSC equally well supported the development of Tregs, indicating that NO is not a conditio sine qua non for the generation of Tregs in all settings [70]. Second, Brahmachari et al. provided evidence that antigenpriming or stimulation of mouse splenocytes with IL-12p40 homodimers leads to an iNOS/NO-dependent downregulation of Foxp3 in CD4+CD25+ T cells. The authors tacitly assume that the decrease of Foxp3 leads to a loss of suppressor function of the Tregs [71, 72]. In the light of the well-documented existence of Foxp3− Tregs [1] and the above-cited study by Niedbala et al. [68], this is not necessarily the case. Third, a subset of antigeninduced regulatory T cells that express the transcription factor T-bet and exhibit a Th1 phenotype, exert their inhibitory effect on disease-mediating pathogenic target T cells in a cell contact-, IFN-g-, myeloid cell- and NO-dependent manner [24]. Thus, iNOS-derived NO can function as a potential inducer, a modulator, or as a terminal effector molecule of Tregs. 4.1.2. T-Cell Proliferation

The first evidence that the proliferation of lymphocytes is regulated by the iNOS-dependent metabolism of l-arginine, was obtained by Hoffman et al. 20 years ago, when they observed that the alloantigen-induced proliferation and induction of cytotoxic T-cell activity of rat or mouse splenocytes were drastically enhanced in the presence of the NOS inhibitor NG-monomethyll-arginine (l-NMMA), an effect that could be reversed by the addition of l-arginine [73]. The authors excluded a lack of T-cell stimulatory cytokines in the control cultures without l-NMMA, but otherwise did not further analyze the reported phenotype at that time. However, they correctly anticipated and discussed the three possible mechanisms underlying the l-arginine-dependent suppression of T-cell proliferation and activity: (a) a direct cytostatic effect of oxidative l-arginine metabolites (iNOS-derived NO and other RNI) on T cells; (b) a retrograde regulatory or cytotoxic effect of NO (or other RNI) on iNOS-positive

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antigen-presenting myeloid cells; (c) a depletion of l-arginine by the iNOS reaction. During the following years, each of these postulated mechanisms was supported by the data obtained in various experimental systems. iNOS-derived NO was shown to inhibit IL-2 receptor signaling (Jak3 and STAT5 tyrosine phosphorylation) [74], IL-2 mRNA expression [75], ribonucleotide reductase activity [76], and caspase activation in T cells and to promote their expression of heme oxygenase [77]; all of these effects might contribute to the impaired proliferative response of T cells (Fig. 1, mechanism 1). Conventional DC exposed to endogenously produced NO became apoptotic with a subsequent feedback-inhibitory effect on T-cell proliferation [51] or failed to support the secretion of IFN-g by T cells [5]. Also, expression of iNOS/NO by MDSC impeded the development of T-cell-stimulatory DC [78] (Fig. 1, mechanism 2). The consumption of arginine by iNOS-expressing myeloid cells might impair the arginase-dependent generation of ornithine and the ornithine decarboxylase-driven synthesis of polyamines that are necessary for the growth and proliferation of lymphocytes. Alternatively, it is possible that a reduced arginine availability affects the translation of certain mRNAs in T cells as has been described for macrophages, astroglia, and other cell types following amino acid depletion [42] (for review see [43]) (Fig. 1, mechanism 3). In addition to iNOS, T cells can also be deprived of l-arginine by the expression of arginase in myeloid cells (macrophages, DCs, or MDSC), infectious pathogens (e.g., Helicobacter pylori and

Polyamines

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Mechanism 4 •฀uncoupling฀of฀iNOS฀ → H2O2 ↑ •฀CD3z ↓ •฀translation฀↓(?)

Mechanism 1 •฀phosphorylation฀Jak3฀/฀Stat5฀↓ •฀ribonucleotide฀reductase฀↓ •฀caspase฀activity ↓ •฀IL-2฀mRNA ↓

Fig. 1. l-Arginine metabolism and its impact on T-cell proliferation. For details, references, and abbreviations see main text.

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Leishmania major), or certain tumor cells (Fig. 1, mechanism 4). Arginase efficiently impeded T-cell proliferation and the antimicrobial or anti-tumor response in various experimental systems [16, 79–85]. Although iNOS and arginase are induced in myeloid cells by different sets of cytokines, the expression of arginase and iNOS is not mutually exclusive. In fact, certain populations of myeloid suppressor cells that produced IFN-g as well as IL-13 were shown to simultaneously express iNOS and arginase 1 [86, 87]. One well-described molecular effect of arginine depletion is the selective downregulation of the CD3z chain on T lymphocytes, which is critical for T-cell-receptor-mediated signaling events and initiation of T-cell proliferation. The suppression of CD3z protein most likely results from a drastic reduction of the half-life of CD3z mRNA in the absence of arginine [88] and was observed with T cells that were grown in l-arginine-deficient medium or cocultured with macrophages, pathogens, or tumor cells expressing arginase [82, 83, 89]. Arginase might also account for the downregulation of CD3z seen in patients with active tuberculosis [90]. Another well-known effect of arginine depletion by arginase is the uncoupling of the NOS reaction: in the absence of l-arginine, all NOS isoforms discontinue to generate NO and start to produce superoxide (O2−) [91–93], which can be converted by superoxide dismutase to peroxide (H2O2). H2O2 is known as strong suppressor of T-cell responses [94–96]. There is evidence for a differential impact of iNOS-derived NO on T-cell subpopulations. Mouse CD8+ T cells were more susceptible to the anti-proliferative effect of NO than CD4+ T cells [97]. NO generated by MDSC blocked the development of allogeneic cytotoxic T cells [98, 99]. When Th1 or Th2 cell lines were cocultured with antigen-loaded mouse brain microglia, Th1, but not Th2 cells underwent apoptosis [100]. In contrast, when human Th1 and Th2 were cocultured with human bronchial epithelial cells, the induction of iNOS by endogenously produced (i.e., Th1-derived) or exogenously added IFN-g led to a comparable inhibition of Th1 and Th2 cell proliferation [101]. Likewise, mouse Th1 and Th2 clones were equally susceptible to the growth-inhibitory effect of the NO donor SNAP [102]. Thus, iNOS-derived NO is capable to limit the expansion of either type of T-helper cell, which might be relevant for protection against overshooting T-cell responses. 4.1.3. T-Cell Cytokine Production

There is no uniform picture of the effect of exogenous NO on the production of cytokines by differentiated T cells. While some studies suggested a selectivity of NO for the suppression of IFN-g or IL-2 release by mouse and human T cell lines [103–106], others came to the conclusion that NO donors equally well inhibit

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the release of Th1 and Th2 cytokines in mixed T-cell or leukocyte populations [75, 107] or even questioned an effect of NO at non-toxic concentrations [102, 108]. The main deficit of all these in vitro studies is that they resorted to exogenously added NO donors rather than analyzing the role of NO produced by myeloid cells under coculture conditions. 4.1.4. T-Cell Adhesion and Migration

As has been reported for several other cell types, NO is capable to inhibit the adhesion of Th1 cells to inflamed endothelial cells. The effect is mediated by IFN-g-induced iNOS and therefore represents a feedback mechanism to limit the recruitment of new Th1 cells to inflammatory sites [109]. In line with these results, NO was found to alter the morphology of encephalitogenic T cells and to block their transendothelial migration [110].

4.2. Endogenous NO and T Lymphocytes

iNOS, nNOS, or eNOS have been detected in mouse or human T cell lines, clones, or hybridomas in vitro, but the quality of the published data and the degree of evidence for a cell-autonomous function of NO in these cells are quite variable and require critical assessment (e.g., primary cells vs. transformed cells or cloned cells; purity of the analyzed cells; analysis on a total population vs. single cell level; mRNA vs. protein vs. enzyme activity vs. NO production) (for review see [23]; [103, 104, 111, 112]). To date, only very few studies addressed the question whether primary mouse or human T lymphocytes express NOS isoforms on a single cell level. While several studies failed to detect iNOS or cNOS mRNA and protein in mouse or human T cells on a population or single cell level [61, 107] (See Note 1), some analyses argue for the expression of iNOS, nNOS, or eNOS in purified (>90–99% CD3+ cells) splenic or peripheral blood mouse or human T lymphocytes. Notably, independent of the NOS isoform detected in the T cells, NO was found to modulate T-cell proliferation, survival, or death: 1. iNOS: In mouse and human T cells, iNOS-derived NO contributed to postactivation-induced cell death. Increased levels of the anti-apoptotic proteins Bcl-2 and Bcl-xL were detectable in iNOS−/− T cells compared to iNOS+/+ T cells. In an ovalbumin immunization model, higher frequencies of CD4+ or CD8+ memory T cells occurred in iNOS−/− compared to iNOS+/+ mice, even if MHC-mismatched iNOS+/+ APCs were used for the immunization, supporting a T-cell-autonomous action of NO [62]. Somewhat opposing results were obtained in a study where iNOS mRNA and protein was induced in CD8+ as well as CD4+ human T cells upon coculture with allogeneic endothelial cells and preferentially expressed in

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non-proliferating T cells. Surprisingly, inhibition of iNOS reduced the number of proliferated T cells, suggesting that iNOS expression in bystander T cells is a prerequisite for the proliferation of alloreactive T cells [113, 114]. A similar growth-supportive role of iNOS was also observed in BW5147 T-lymphoma cells [111]. 2. nNOS: In the murine T-cell hybridoma 2B4, human Jurkat T cells, and primary human T-cell blasts, nNOS-mediated NO production was implicated in the TCR-triggered death of mature T lymphocytes [115]. 3. eNOS: Confirming earlier results [116], eNOS mRNA and protein (but no iNOS protein) was found in mouse and human T lymphoblasts or in TCR-transfected Jurkat T cells [65]. Stimulation with anti-CD3, antigen-, or superantigenexpressing APCs led to the translocation of eNOS along with the Golgi complex to the immunological synapse, the production of NO, the phosphorylation of TCR signaling molecules (CD3z, ZAP-70), and the activation of MAPK via S-nitrosylation of the Golgi-localized GTPase N-Ras. Transfection of Jurkat cells with wildtype N-Ras, but not with mutated N-Ras (which lacked a cysteine residue essential for S-nitrosylation), increased TCR-dependent cell death [64, 65]. Activation of human T cells with anti-CD3 plus antiCD28 caused an upregulation of eNOS and nNOS protein and a NO-dependent mitochondrial hyperpolarization – a process that is associated with either T-cell proliferation or T-cell death [117]. In human g/d T cells, eNOS activity conferred partial protection against anti-CD95 (Fas)-induced cell death [118]. 4.3. NO and NK Cells

The activation of NK cells for cytotoxic activity and IFN-g production requires an interaction with myeloid cells [119–121]. A series of previous studies revealed that NO can be involved in various aspects of NK cell biology, depending on the species and tissue origin (mouse, rat, or human NK cells; lymph node, splenic, and blood NK cells), the prior activation status of the NK cells [resting NK cells vs. IL-2-expanded NK cells (lymphokine-activated killer cells, LAK)], and the source of NO. The NO was either generated by iNOS-positive myeloid cells or derived from iNOS [122–124] or eNOS expressed within the NK cell itself [125]. Although the picture is far from uniform, three major categories of actions of NO on NK cells have emerged: 1. Induction of NK cell differentiation, activity, and survival: Endogenous expression of iNOS was described as a maturation

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factor in the IL-2-, IL-12-, or NK cell-receptor-triggered activation of rodent NK cells for the production of IFN-g and – in the case of mouse NK cells – also for the development of cytotoxic activity [122, 123, 126]. In mouse uterine NK cells, which accumulate in the placenta during pregnancies, iNOS was necessary for the expression of perforin [127]. In human peripheral blood NK cells, which were positive for eNOS and released NO in response to target cells or antiCD16 stimulation in the presence of IL-2, NO impeded apoptosis and TNF expression and contributed to target cell lysis [125]. 2. Downregulation of NK cell activity: In human peripheral blood NK cells, the induction of iNOS by IL-12 (±TNF) impaired the IFN-g production, the granzyme B expression, and the cytolytic activity against tumor cells [124]. A similar inhibitory effect was also seen in mouse, rat, or human NK cells or IL-2-activated LAK cells that were exposed to exogenous NO generated by iNOS-positive macrophages or released by NO donors (reviewed in [128, 129]; [130, 131]). One underlying mechanism is the inhibition of cytolytic granule exocytosis by NO [131]. 3. NO as cytotoxic effector molecule: A series of studies with mouse, rat, or human NK cells provided evidence that NO generated by NK cells via the iNOS- or eNOS-pathway directly contributes to the killing of tumor targets (reviewed in [129]). 4.4. NO and B Cells

iNOS or eNOS protein was detected in B-cell lines (Burkitt lymphoma cell lines, B-cell chronic lymphocytic leukemia cells, and CD5+ IgM+ WEHI 231 cells) and in primary B cells (human peritoneal B1 cells and IgD+ B cells), but except for a possible anti-apoptotic effect no functional role was reported (for review see [23]; [116]). More recent studies revealed that the immunoglobin expression is under the control of iNOS. After influenza virus A infection, iNOS−/− mice had more viable B cells and plasmablasts and higher levels of virus-specific IgG2a antibodies [132]. In the mucosa-associated lymphatic system of the gut, iNOS expressed by TIP-DC in response to commensal bacteria was essential for the T-cell-dependent as well as T-cellindependent IgA class switch recombination. In the absence of iNOS, the concentrations of IgA in the serum and in the intestine were strongly reduced. iNOS-deficiency caused a drastic reduction of the TGF-b receptor signaling chain (TGF-bR2) and its downstream molecules (Runx3 and Smad3) in stimulated B cells as well as of the B-cell activating factors APRIL and BAFF in intestinal DCs, which explains the defect of the T-cell-dependent or T-cell-independent IgA class switch, respectively [133].

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5. Therapeutic Implications Considering the above-mentioned effects of the l-arginine metabolism on the function and proliferation of lymphocytes, a number of potential therapeutic strategies appear feasible. These include (1) the inhibition of iNOS/NO-production and/or arginase 1 in malignant tumors, where iNOS-derived NO might promote tumor metastasis [134] and different subsets of myeloid suppressor cells (including tumor-associated macrophages and activated neutrophils) or the tumor cells themselves deprive infiltrating T cells of arginine and thereby hinder the anti-tumor T-cell response [16, 80, 86, 135]; (2) the inhibition of iNOS activity/ NO production during autoimmune processes, where NO maintains inflammatory processes and contributes to the tissue damage [49]; (3) the inhibition of iNOS activity/NO production during severe acute infections (e.g., viral pneumonia and bacterial sepsis), where hyperproduction of NO not only accounts for the suppression of the antigen-specific T-cell response, but also for vasodilatation and shock and for fulminant pathology and failure of organs [136]; (4) the activation of iNOS activity/NO production during chronic infections, where iNOS/NO is suppressed in certain host cell subpopulations and thereby allows for lifelong pathogen persistence. The major problem with any of these therapeutic approaches is that the targeted pathway (iNOS or arginase) not only exerts detrimental effects in the setting of a specific disease (e.g., malignant tumor, autoimmune, or infectious disease), but also triggers disease-limiting and host-protective processes. Thus, in malignancies, iNOS-derived NO also acts as a cytotoxic, tumoricidal molecule [23]; in autoimmunity, iNOS/NO is a crucial mechanism to counteract the hyperactive T-cell response and to restrict the influx of inflammatory cells [49, 109, 137]; and in severe bacterial infections, iNOS/NO helps to prevent thrombosis and secondary ischemia of organs [22, 138]. Another critical issue is the specificity of NOS or arginase inhibitors. NOS inhibitors that are used for the modulation of the immune response need to be specific for iNOS and leave eNOS and nNOS unaffected in order to avoid cardiovascular, hepatic, renal, or neurological side effects [139, 140]. In addition, it would be highly desirable to avoid systemic effects of these inhibitors, either by targeting them to defined organs or cells or by designing compounds that only become active at the site of the disease process. Similar considerations and pharmaceutical developments are necessary prior to clinical use of NO donors [141–143]. With respect to arginase inhibitors, the urea cycle in the liver, which is essential for life and dependent on arginase 1 activity [144], currently obviates their prolonged systemic application. Future strategies for the therapeutic application of arginase inhibitors [145] in malignant diseases will have to incorporate modes for tumor-specific targeting of these compounds.

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6. Note 1. In a recently published study (Fritzsche, C., Schleicher, U., and Bogdan, C. 2010. Endothelial nitric oxide synthase limits the inflammatory response in mouse cutaneous leishmaniasis. Immunobiology 215, 826–832) it was reported that naive T lymphocytes, Th1 cells and Th2 cells did not express eNOS after stimulation and that eNOS was not required for Th1 differentiation in vitro. Lymph node-derived T cells from Leishmania major-infected wildtype and eNOS-/- mice released comparable amounts of IFN-g and proliferated equally well.

Acknowledgments I wish to apologize to all researchers whose work could only be cited in form of review articles due to space restrictions. The preparation of this chapter and the conduct of some of the studies reviewed were supported by grants to C.B. from the Deutsche Forschungsgemeinschaft (Bo996/3-3, SFB643 A6) and from the IZKF Erlangen (Project A24). References 1. Vignali, D. A., Collison, L. W., and Workman, C. J. (2008) How regulatory T cells work. Nat Rev Immunol 8, 523–32. 2. Lutz, M. B., and Schuler, G. (2002) Immature, semi-mature and fully mature dendritic cells: which signals induce tolerance or immunity? Trends Immunol 23, 445–9. 3. Kuang, D. M., Zhao, Q., Xu, J., Yun, J. P., Wu, C., and Zheng, L. (2008) Tumoreducated tolerogenic dendritic cells induce CD3epsilon down-regulation and apoptosis of T cells through oxygen-dependent pathways. J Immunol 181, 3089–98. 4. Zhang, M., Tang, H., Guo, Z., An, H., Zhu, X., Song, W., Guo, J., Huang, X., Chen, T., Wang, J., and Cao, X. (2004) Splenic stroma drives mature dendritic cells to differentiate into regulatory dendritic cells. Nat Immunol 5, 1124–33. 5. Ren, G., Su, J., Zhao, X., Zhang, L., Zhang, J., Roberts, A. I., Zhang, H., Das, G., and Shi, Y. (2008) Apoptotic cells induce immunosuppression through dendritic cells: critical roles of IFN-gamma and nitric oxide. J Immunol 181, 3277–84. 6. Norian, L. A., Rodriguez, P. C., O’Mara, L. A., Zabaleta, J., Ochoa, A. C., Cella, M.,

7.

8.

9.

10.

11.

12.

and Allen, P. M. (2009) Tumor-infiltrating regulatory dendritic cells inhibit CD8+ T cell function via l-arginine metabolism. Cancer Res 69, 3086–94. Schechter, G. P., Wahl, L. M., and Oppenheim, J. J. (1979) Suppressor monocytes in human disease: a review. Adv Exp Med Biol 121B, 283–98. Elgert, K. D., Alleva, D. G., and Mullins, D. W. (1998) Tumor-induced immune dysfunction: the macrophage connection. J Leukoc Biol 64, 275–90. Bronte, V., and Zanovello, P. (2005) Regulation of immune responses by l-arginine metabolism. Nat Rev Immunol 5, 641–53. Gabrilovich, D. I., and Nagaraj, S. (2009) Myeloid-derived suppressor cells as regulators of the immune system. Nat Rev Immunol 9, 162–74. Zhang, X., Majlessi, L., Deriaud, E., Leclerc, C., and Lo-Man, R. (2009) Coactivation of Syk kinase and MyD88 adaptor protein pathways by bacteria promotes regulatory properties of neutrophils. Immunity 31, 761–71. Blois, S. M., Ilarregui, J. M., Tometten, M., Garcia, M., Orsal, A. S., Cordo-Russo, R., Toscano, M. A., Bianco, G. A., Kobelt, P.,

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13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

Handjiski, B., Tirado, I., Markert, U. R., Klapp, B. F., Poirier, F., Szekeres-Bartho, J., Rabinovich, G. A., and Arck, P. C. (2007) A pivotal role for galectin-1 in fetomaternal tolerance. Nat Med 13, 1450–7. Bogdan, C., Vodovotz, Y., and Nathan, C. (1991) Macrophage deactivation by IL-10. J Exp Med 174, 1549–55. Mosser, D. M., and Zhang, X. (2008) Interleukin-10: new perspectives on an old cytokine. Immunol Rev 226, 205–18. Bogdan, C., and Nathan, C. (1993) Modulation of macrophage function by transforming growth factor-b, interleukin 4 and interleukin 10. Ann N Y Acad Sci 685, 713–39. Rodriguez, P. C., Hernandez, C. P., Quiceno, D., Dubinett, S. M., Zabaleta, J., Ochoa, J. B., Gilbert, J., and Ochoa, A. C. (2005) Arginase I in myeloid suppressor cells is induced by COX-2 in lung carcinoma. J Exp Med 202, 931–9. Serhan, C. N., Chiang, N., and Van Dyke, T. E. (2008) Resolving inflammation: dual antiinflammatory and pro-resolution lipid mediators. Nat Rev Immunol 8, 349–61. Deaglio, S., Dwyer, K. M., Gao, W., Friedman, D., Usheva, A., Erat, A., Chen, J. F., Enjyoji, K., Linden, J., Oukka, M., Kuchroo, V. K., Strom, T. B., and Robson, S. C. (2007) Adenosine generation catalyzed by CD39 and CD73 expressed on regulatory T cells mediates immune suppression. J Exp Med 204, 1257–65. Romani, L., Fallarino, F., De Luca, A., Montagnoli, C., D’Angelo, C., Zelante, T., Vacca, C., Bistoni, F., Fioretti, M. C., Grohmann, U., Segal, B. H., and Puccetti, P. (2008) Defective tryptophan catabolism underlies inflammation in mouse chronic granulomatous disease. Nature 451, 211–5. Song, R., Mahidhara, R. S., Zhou, Z., Hoffman, R. A., Seol, D. W., Flavell, R. A., Billiar, T. R., Otterbein, L. E., and Choi, A. M. (2004) Carbon monoxide inhibits T lymphocyte proliferation via caspase-dependent pathway. J Immunol 172, 1220–6. Munder, M. (2009) Arginase: an emerging key player in the mammalian immune system. Br J Pharmacol 158, 638–51. Bogdan, C. (2000) The function of nitric oxide in the immune system, in Handbook of experimental pharmacology. Volume: nitric oxide (Mayer, B., Ed.) pp 443–92, Springer, Heidelberg. Bogdan, C. (2001) Nitric oxide and the immune response. Nat Immunol 2, 907–16.

387

24. Chen, C., and Liu, C. P. (2009) Regulatory function of a novel population of mouse autoantigen-specific Foxp3 regulatory T cells depends on IFN-gamma, NO, and contact with target cells. PLoS One 4, e7863. 25. Bogdan, C. (2001) Nitric oxide and the regulation of gene expression. Trends Cell Biol 11, 66–75. 26. Fang, F. C. (2004) Antimicrobial reactive oxygen and nitrogen species: concepts and controversies. Nat Rev Immunol 2, 820–32. 27. Bogdan, C. (2004) Reactive oxygen and reactive nitrogen metabolites as effector molecules against infectious pathogens, in The innate immune response to infection (Kaufmann, S. H. E., Medzhitov, R., and Gordon, S., Eds.) pp 357–96, ASM Press, Washington, DC. 28. Bogdan, C. (2009) Regulation and antimicrobial function of inducible nitric oxide synthase in macrophages, in Phagocyte–pathogen interaction (Russell, D. and Gordon, S., Eds.) pp 367–78, ASM Press, Washington. 29. Stuehr, D. J., Tejero, J., and Haque, M. M. (2009) Structural and mechanistic aspects of flavoproteins: electron transfer through the nitric oxide synthase flavoprotein domain. FEBS J 276, 3959–74. 30. Nathan, C. (1992) Nitric oxide as a secretory product of mammalian cells. FASEB J. 6, 3051–64. 31. Li, H., and Forstermann, U. (2009) Prevention of atherosclerosis by interference with the vascular nitric oxide system. Curr Pharm Des 15, 3133–45. 32. Forstermann, U., and Munzel, T. (2006) Endothelial nitric oxide synthase in vascular disease: from marvel to menace. Circulation 113, 1708–14. 33. Zhou, L., and Zhu, D. Y. (2009) Neuronal nitric oxide synthase: structure, subcellular localization, regulation, and clinical implications. Nitric Oxide 20, 223–30. 34. Durand, J. L., Mukherjee, S., Commodari, F., De Souza, A. P., Zhao, D., Machado, F. S., Tanowitz, H. B., and Jelicks, L. A. (2009) Role of NO synthase in the development of Trypanosoma cruzi-induced cardiomyopathy in mice. Am J Trop Med Hyg 80, 782–7. 35. Cui, X., Besch, V., Khaibullina, A., Hergen, A., Quezado, M., Eichacker, P., and Quezado, Z. M. (2007) Neuronal nitric oxide synthase deficiency decreases survival in bacterial peritonitis and sepsis. Intensive Care Med 33, 1993–2003. 36. Kajiya, K., Huggenberger, R., Drinnenberg, I., Ma, B., and Detmar, M. (2008) Nitric oxide mediates lymphatic vessel activation via

388

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

Bogdan soluble guanylate cyclase alpha1beta1-impact on inflammation. FASEB J 22, 530–7. Nathan, C. (1997) Inducible nitric oxide synthase: what difference does it make? J Clin Invest 100, 2417–23. Vodovotz, Y., Bogdan, C., Paik, J., Xie, Q. W., and Nathan, C. (1993) Mechanisms of suppression of macrophage nitric oxide release by transforming growth factor-b. J Exp Med 178, 605–13. Mitani, T., Terashima, M., Yoshimura, H., Nariai, Y., and Tanigawa, Y. (2005) TGFbeta1 enhances degradation of IFN-gammainduced iNOS protein via proteasomes in RAW 264.7 cells. Nitric Oxide 13, 78–87. Takaki, H., Minoda, Y., Koga, K., Takaesu, G., Yoshimura, A., and Kobayashi, T. (2006) TGF-beta1 suppresses IFN-gammainduced NO production in macrophages by suppressing STAT1 activation and accelerating iNOS protein degradation. Genes Cells 11, 871–82. Chen, L., Kong, X., Fu, J., Xu, Y., Fang, S., Hua, P., Luo, L., and Yin, Z. (2009) CHIP facilitates ubiquitination of inducible nitric oxide synthase and promotes its proteasomal degradation. Cell Immunol 258, 38–43. El-Gayar, S., Thüring-Nahler, H., Pfeilschifter, J., Röllinghoff, M., and Bogdan, C. (2003) Translational control of inducible nitric oxide synthase by IL-13 and arginine availability in inflammatory macrophages. J Immunol 171, 4561–8. König, T., Bogdan, C., and Schleicher, U. (2009) Translational repression of inducible NO synthase in macrophages by l-arginine depletion is not associated with an increased phosphorylation of eIF2alpha. Immunobiology 214, 822–7. Schneider, E., and Dy, M. (1985) The role of arginase in the immune response. Immunol Today 6, 136–9. Rutschman, R., Lang, R., Hesse, M., Ihle, J. N., Wynn, T. A., and Murray, P. J. (2001) Stat6-dependent substrate depletion regulates nitric oxide production. J Immunol 166, 2173–7. El Kasmi, K. C., Qualls, J. E., Pesce, J. T., Smith, A. M., Thompson, R. W., HenaoTamayo, M., Basaraba, R. J., Konig, T., Schleicher, U., Koo, M. S., Kaplan, G., Fitzgerald, K. A., Tuomanen, E. I., Orme, I. M., Kanneganti, T. D., Bogdan, C., Wynn, T. A., and Murray, P. J. (2008) Toll-like receptor-induced arginase 1 in macrophages thwarts effective immunity against intracellular pathogens. Nat Immunol 9, 1399–406.

47. Morris, S. M., Jr. (2009) Recent advances in arginine metabolism: roles and regulation of the arginases. Br J Pharmacol 157, 922–30. 48. Kuroda, E., Ho, V., Ruschmann, J., Antignano, F., Hamilton, M., Rauh, M. J., Antov, A., Flavell, R. A., Sly, L. M., and Krystal, G. (2009) SHIP represses the generation of IL-3-induced M2 macrophages by inhibiting IL-4 production from basophils. J Immunol 183, 3652–60. 49. Bogdan, C. (1998) The multiplex function of nitric oxide in (auto)immunity. J Exp Med 187, 1361–5. 50. Stenger, S., Donhauser, N., Thüring, H., Röllinghoff, M., and Bogdan, C. (1996) Reactivation of latent leishmaniasis by inhibition of inducible nitric oxide synthase. J Exp Med 183, 1501–14. 51. Lu, L., Bonham, C. A., Chambers, F. G., Watkins, S. C., Hoffman, R. A., Simmons, R. L., and Thomson, A. W. (1996) Induction of nitric oxide synthase in mouse dendritic cells by IFN-g, endotoxin, and interaction with allogeneic T cells. Nitric oxide production is associated with dendritic cell apoptosis. J Immunol 157, 3577–86. 52. Eriksson, S., Chambers, B. J., and Rhen, M. (2003) Nitric oxide produced by murine dendritic cells is cytotoxic for intracellular Salmonella enterica sv. Typhimurium. Scand J Immunol 58, 493–502. 53. Serbina, N. V., Kuziel, W., Flavell, R., Akira, S., Rollins, B., and Pamer, E. G. (2003) Sequential MyD88-independent and -dependent activation of innate immune responses to intracellular bacterial infection. Immunity 19, 891–901. 54. Tam, M. A., and Wick, M. J. (2006) Differential expansion, activation and effector functions of conventional and plasmacytoid dendritic cells in mouse tissues transiently infected with Listeria monocytogenes. Cell Microbiol 8, 1172–87. 55. Angulo, I., Rodriguez, R., Garcia, B., Medina, M., Navarro, J., and Subiza, J. L. (1995) Involvement of nitric oxide in bone marrow-derived natural suppressor activity. Its dependence on IFN-gamma. J Immunol 155, 15–26. 56. Angulo, I., de las Heras, F. G., Garcia-Bustos, J. F., Gargallo, D., Munoz-Fernandez, M. A., and Fresno, M. (2000) Nitric oxide-producing CD11b(+)Ly-6G(Gr-1)(+) CD31(ER-MP12)(+) cells in the spleen of cyclophosphamide-treated mice: implications for T-cell responses in immunosuppressed mice. Blood 95, 212–20. 57. Pelaez, B., Campillo, J. A., Lopez-Asenjo, J. A., and Subiza, J. L. (2001) Cyclophosphamide

Regulation of Lymphocytes by Nitric Oxide

58.

59.

60.

61.

62.

63.

64.

65.

66.

induces the development of early myeloid cells suppressing tumor cell growth by a nitric oxide-dependent mechanism. J Immunol 166, 6608–15. Chao, C. C., Hu, S., Molitor, T. W., Shaskan, E. G., and Peterson, P. K. (1992) Activated microglia mediate neuronal cell injury via a nitric oxide mechanism. J Immunol 149, 2736–41. Gilchrist, M., Savoie, M., Nohara, O., Wills, F. L., Wallace, J. L., and Befus, A. D. (2002) Nitric oxide synthase and nitric oxide production in in vivo-derived mast cells. J Leukoc Biol 71, 618–24. Amrouni, D., Gautier-Sauvigne, S., Meiller, A., Vincendeau, P., Bouteille, B., Buguet, A., and Cespuglio, R. (2010) Cerebral and peripheral changes occurring in nitric oxide (NO) synthesis in a rat model of sleeping sickness: identification of brain iNOS expressing cells. PLoS One 5, e9211. Thüring, H., Stenger, S., Gmehling, D., Röllinghoff, M., and Bogdan, C. (1995) Lack of inducible nitric oxide synthase activity in T cell-clones and T lymphocytes from naive and Leishmania major-infected mice. Eur J Immunol 25, 3229–34. Vig, M., Srivastava, S., Kandpal, U., Sade, H., Lewis, V., Sarin, A., George, A., Bal, V., Durdik, J. M., and Rath, S. (2004) Inducible nitric oxide synthase in T cells regulates T cell death and immune memory. J Clin Invest 113, 1734–42. Kamimura, Y., Fujii, T., Kojima, H., Nagano, T., and Kawashima, K. (2003) Nitric oxide (NO) synthase mRNA expression and NO production via muscarinic acetylcholine receptor-mediated pathways in the CEM, human leukemic T-cell line. Life Sci 72, 2151–4. Ibiza, S., Perez-Rodriguez, A., Ortega, A., Martinez-Ruiz, A., Barreiro, O., GarciaDominguez, C. A., Victor, V. M., Esplugues, J. V., Rojas, J. M., Sanchez-Madrid, F., and Serrador, J. M. (2008) Endothelial nitric oxide synthase regulates N-Ras activation on the Golgi complex of antigen-stimulated T cells. Proc Natl Acad Sci U S A 105, 10507–12. Ibiza, S., Victor, V. M., Bosca, I., Ortega, A., Urzainqui, A., O’Connor, J. E., SanchezMadrid, F., Esplugues, J. V., and Serrador, J. M. (2006) Endothelial nitric oxide synthase regulates T cell receptor signaling at the immunological synapse. Immunity 24, 753–65. Niedbala, W., Wei, X.-Q., Piedrafita, D., Xu, D., and Liew, F. Y. (1999) Effects of nitric oxide on the induction and differentiation of Th1 cells. Eur J Immunol 29, 2498–505.

389

67. Niedbala, W., Wei, X.-q., Campbell, C., Thomson, D., Komai-Koma, M., and Liew, F. Y. (2002) Nitric oxide preferentially induces type 1 T cell differentiation by selectively up-regulating IL-12 receptor b2 expression via cGMP. Proc Natl Acad Sci U S A 99, 16186–91. 68. Niedbala, W., Cai, B., Liu, H., Pitman, N., Chang, L., and Liew, F. Y. (2007) Nitric oxide induces CD4+CD25+ Foxp3 regulatory T cells from CD4+CD25 T cells via p53, IL-2, and OX40. Proc Natl Acad Sci U S A 104, 15478–83. 69. Feng, G., Gao, W., Strom, T. B., Oukka, M., Francis, R. S., Wood, K. J., and Bushell, A. (2008) Exogenous IFN-gamma ex vivo shapes the alloreactive T-cell repertoire by inhibition of Th17 responses and generation of functional Foxp3+ regulatory T cells. Eur J Immunol 38, 2512–27. 70. Huang, B., Pan, P. Y., Li, Q., Sato, A. I., Levy, D. E., Bromberg, J., Divino, C. M., and Chen, S. H. (2006) Gr-1+CD115+ immature myeloid suppressor cells mediate the development of tumor-induced T regulatory cells and T-cell anergy in tumor-bearing host. Cancer Res 66, 1123–31. 71. Brahmachari, S., and Pahan, K. (2009) Suppression of regulatory T cells by IL-12p40 homodimer via nitric oxide. J Immunol 183, 2045–58. 72. Brahmachari, S., and Pahan, K. (2010) Myelin basic protein priming reduces the expression of Foxp3 in T cells via nitric oxide. J Immunol 184, 1799–809. 73. Hoffman, R. A., Langrehr, J. M., Billiar, T. R., Curran, R. D., and Simmons, R. L. (1990) Alloantigen-induced activation of rat splenocytes is regulated by the oxidative metabolism of l-arginine. J. Immunol 145, 2220–6. 74. Bingisser, R. M., Tilbrook, P. A., Holt, P. G., and Kees, U. R. (1998) Macrophagederived nitric oxide regulates T cell activation via reversible disruption of the Jak3/ Stat5 signaling pathway. J Immunol 160, 5729–34. 75. Macphail, S. E., Gibney, C. A., Brooks, B. M., Booth, C. G., Flanagan, B. F., and Coleman, J. W. (2003) Nitric oxide regulation of human peripheral blood mononuclear cells: critical time dependence and selectivity for cytokine versus chemokine expression. J Immunol 171, 4809–15. 76. Henson, S. E., Nichols, T. C., Holers, V. M., and Karp, D. R. (1999) The ectoenzyme g-glutamyl transpeptidase regulates antiproliferative effects of S-nitrosoglutathione on

390

77.

78.

79.

80.

81.

82.

83.

84.

85.

Bogdan human T and B lymphocytes. J Immunol 163, 1845–52. Mahidhara, R. S., Hoffman, R. A., Huang, S., Wolf-Johnston, A., Vodovotz, Y., Simmons, R. L., and Billiar, T. R. (2003) Nitric oxide-mediated inhibition of caspasedependent T lymphocyte proliferation. J Leukoc Biol 74, 403–11. Greifenberg, V., Ribechini, E., Rossner, S., and Lutz, M. B. (2009) Myeloid-derived suppressor cell activation by combined LPS and IFN-gamma treatment impairs DC development. Eur J Immunol 39, 2865–76. Bronte, V., Serafini, P., de Santo, C., Marigo, I., Tosello, V., Mazzoni, A., Segal, D. M., Staib, C., Lowel, M., Sutter, G., Colombo, M. P., and Zanovello, P. (2003) IL-4-induced arginase 1 suppresses alloreactive T cells in tumor-bearing mice. J Immunol 170, 270–8. Rodriguez, P. C., Ernstoff, M. S., Hernandez, C., Atkins, M., Zabaleta, J., Sierra, R., and Ochoa, A. C. (2009) Arginase I-producing myeloid-derived suppressor cells in renal cell carcinoma are a subpopulation of activated granulocytes. Cancer Res 69, 1553–60. Rodriguez, P. C., Quiceno, D. G., and Ochoa, A. C. (2007) l-arginine availability regulates T-lymphocyte cell-cycle progression. Blood 109, 1568–73. Rodriguez, P. C., Quiceno, D. G., Zabaleta, J., Ortiz, B., Zea, A. H., Piazuelo, M. B., Delgado, A., Correa, P., Brayer, J., Sotomayor, E. A., Antonia, S., Ochoa, J. B., and Ochoa, A. C. (2004) Arginase I production in the tumor microenvironment by mature myeloid cells inhibits T cell-receptor expression and antigen-specific T cell responses. Cancer Res 64, 5839–49. Zabaleta, J., McGee, D. J., Zea, A. H., Hernandez, C. P., Rodriguez, P. C., Sierra, R. A., Correa, P., and Ochoa, A. C. (2004) Helicobacter pylori arginase inhibits T cell proliferation and reduces the expression of the TCR zeta-chain (CD3zeta). J Immunol 173, 586–93. Modolell, M., Choi, B. S., Ryan, R. O., Hancock, M., Titus, R. G., Abebe, T., Hailu, A., Muller, I., Rogers, M. E., Bangham, C. R., Munder, M., and Kropf, P. (2009) Local suppression of T cell responses by arginaseinduced l-arginine depletion in nonhealing leishmaniasis. PLoS Negl Trop Dis 3, e480. Munder, M., Choi, B. S., Rogers, M., and Kropf, P. (2009) l-arginine deprivation impairs Leishmania major-specific T-cell responses. Eur J Immunol 39, 2161–72.

86. Bronte, V., Kasic, T., Gri, G., Gallana, K., Borsellino, G., Marigo, I., Battistini, L., Iafrate, M., Prayer-Galetti, T., Pagano, F., and Viola, A. (2005) Boosting antitumor responses of T lymphocytes infiltrating human prostate cancers. J Exp Med 201, 1257–68. 87. Gallina, G., Dolcetti, L., Serafini, P., De Santo, C., Marigo, I., Colombo, M. P., Basso, G., Brombacher, F., Borrello, I., Zanovello, P., Bicciato, S., and Bronte, V. (2006) Tumors induce a subset of inflammatory monocytes with immunosuppressive activity on CD8+ T cells. J Clin Invest 116, 2777–90. 88. Rodriguez, P. C., Zea, A. H., Culotta, K. S., Zabaleta, J., Ochoa, J. B., and Ochoa, A. C. (2002) Regulation of T cell receptor CD3zeta chain expression by l-arginine. J Biol Chem 277, 21123–9. 89. Rodriguez, P. C., Zea, A. H., DeSalvo, J., Culotta, K. S., Zabaleta, J., Quiceno, D. G., Ochoa, J. B., and Ochoa, A. C. (2003) l-arginine consumption by macrophages modulates the expression of CD3z chain in T lymphocytes. J Immunol 17, 1232–9. 90. Zea, A. H., Culotta, K. S., Ali, J., Mason, C., Park, H. J., Zabaleta, J., Garcia, L. F., and Ochoa, A. C. (2006) Decreased expression of CD3zeta and nuclear transcription factor kappa B in patients with pulmonary tuberculosis: potential mechanisms and reversibility with treatment. J Infect Dis 194, 1385–93. 91. Xia, Y., Roman, L. J., Masters, B. S. S., and Zweier, J. L. (1998) Inducible nitric oxide synthase generates superoxide from the reductase domain. J Biol Chem 273, 22635–9. 92. Gao, Y. T., Panda, S. P., Roman, L. J., Martasek, P., Ishimura, Y., and Masters, B. S. (2007) Oxygen metabolism by neuronal nitric-oxide synthase. J Biol Chem 282, 7921–9. 93. Gao, Y. T., Roman, L. J., Martasek, P., Panda, S. P., Ishimura, Y., and Masters, B. S. (2007) Oxygen metabolism by endothelial nitric-oxide synthase. J Biol Chem 282, 28557–65. 94. Schmielau, J., and Finn, O. J. (2001) Activated granulocytes and granulocytederived hydrogen peroxide are the underlying mechanism of suppression of t-cell function in advanced cancer patients. Cancer Res 61, 4756–60. 95. Kusmartsev, S., Nefedova, Y., Yoder, D., and Gabrilovich, D. I. (2004) Antigen-specific inhibition of CD8+ T cell response by imma-

Regulation of Lymphocytes by Nitric Oxide

96.

97.

98.

99.

100.

101.

102.

103.

104.

ture myeloid cells in cancer is mediated by reactive oxygen species. J Immunol 172, 989–99. Brys, L., Beschin, A., Raes, G., Ghassabeh, G. H., Noel, W., Brandt, J., Brombacher, F., and De Baetselier, P. (2005) Reactive oxygen species and 12/15-lipoxygenase contribute to the antiproliferative capacity of alternatively activated myeloid cells elicited during helminth infection. J Immunol 174, 6095–104. Hoffman, R. A., Mahidhara, R. S., WolfJohnston, A. S., Lu, L., Thomson, A. W., and Simmons, R. L. (2002) Differential modulation of CD4 and CD8 T-cell proliferation by induction of nitric oxide synthesis in antigen presenting cells. Transplantation 74, 836–45. Langrehr, J. M., Dull, K. E., Ochoa, J. B., Billiar, T. R., Ildstad, S. T., Schraut, W. H., Simmons, R. L., and Hoffman, R. A. (1992) Evidence that nitric oxide production by in vivo allosensitized cells inhibits the development of allospecific CTL. Transplantation 53, 632–40. Medot-Pirenne, M., Heilman, M. J., Saxena, M., McDermott, P. E., and Mills, C. D. (1999) Augmentation of an antitumor CTL response in vivo by inhibition of suppressor macrophage nitric oxide. J Immunol 163, 5877–82. Monsonego, A., Imitola, J., Zota, V., Oida, T., and Weiner, H. L. (2003) Microgliamediated nitric oxide cytotoxicity of T cells following amyloid beta-peptide presentation to Th1 cells. J Immunol 171, 2216–24. Eriksson, U., Egermann, U., Bihl, M. P., Gambazzi, F., Tamm, M., Holt, P. G., and Bingisser, R. M. (2005) Human bronchial epithelium controls TH2 responses by TH1induced, nitric oxide-mediated STAT5 dephosphorylation: implications for the pathogenesis of asthma. J Immunol 175, 2715–20. van der Veen, R. C., Dietlin, T. A., Pen, L., and Gray, J. D. (1999) Nitric oxide inhibits the proliferation of T-helper 1 and 2 lymphocytes without reduction in cytokine secretion. Cell Immunol 193, 194–201. Taylor-Robinson, A. W., Liew, F. Y., Severn, A., Xu, D., McScorley, S. J., Garside, P., Padron, J., and Phillips, R. S. (1994) Regulation of the immune response by nitric oxide differentially produced by T helper type 1 and T helper type 2 cells. Eur J Immunol 24, 980–4. Benbernou, N., Esnault, S., Shin, H. C., Fekkar, H., and Guenounou, M. (1997) Differential regulation of IFN-gamma, IL-10

105.

106.

107.

108.

109.

110.

111.

112.

113.

114.

391

and inducible nitric oxide synthase in human T cells by cyclic AMP-dependent signal transduction pathway. Immunology 91, 361–8. Chang, R.-H., Lin Feng, M.-H., Liu, W.-H., and Lai, M.-Z. (1997) Nitric oxide increased interleukin-4 expression in T lymphocytes. Immunology 90, 364–9. Fiorucci, S., Antonelli, E., Distrutti, E., Del Soldato, P., Flower, R. J., Clark, M. J., Morelli, A., Perretti, M., and Ignarro, L. J. (2002) NCX-1015, a nitric-oxide derivative of prednisolone, enhances regulatory T cells in the lamina propria and protects against 2,4,6-trinitrobenzene sulfonic acid-induced colitis in mice. Proc Natl Acad Sci U S A 99, 15770–5. Bauer, H., Jung, T., Tsikas, D., Stichtenoth, D. O., Fröhlich, J. C., and Neumann, C. (1997) Nitric oxide inhibits the secretion of T-helper 1- and T-helper 2-associated cytokines in activated human T cells. Immunology 90, 205–11. Marcinkiewicz, J., Grabowska, A., and Chain, B. M. (1996) Is there a role for nitric oxide in regulation of T cell secretion of IL-2? J Immunol 156, 4617–21. Norman, M. U., Zbytnuik, L., and Kubes, P. (2008) Interferon-gamma limits Th1 lymphocyte adhesion to inflamed endothelium: a nitric oxide regulatory feedback mechanism. Eur J Immunol 38, 1368–80. Staykova, M. A., Berven, L. A., Cowden, W. B., Willenborg, D. O., and Crouch, M. F. (2003) Nitric oxide induces polarization of actin in encephalitogenic T cells and inhibits their in vitro trans-endothelial migration in a p70S6 kinase-independent manner. FASEB J 17, 1337–9. Barreiro Arcos, M. L., Gorelik, G., Klecha, A., Goren, N., Cerquetti, C., and Cremaschi, G. A. (2003) Inducible nitric oxide synthasemediated proliferation of a T lymphoma cell line. Nitric Oxide 8, 111–8. Barreiro Arcos, M. L., Gorelik, G., Klecha, A., Genaro, A. M., and Cremaschi, G. A. (2006) Thyroid hormones increase inducible nitric oxide synthase gene expression downstream from PKC-zeta in murine tumor T lymphocytes. Am J Physiol Cell Physiol 291, C327–36. Choy, J. C., Wang, Y., Tellides, G., and Pober, J. S. (2007) Induction of inducible NO synthase in bystander human T cells increases allogeneic responses in the vasculature. Proc Natl Acad Sci U S A 104, 1313–8. Choy, J. C., and Pober, J. S. (2009) Generation of NO by bystander human CD8 T cells augments allogeneic responses by

392

115.

116.

117.

118.

119.

120.

121.

122.

123.

124.

125.

Bogdan inhibiting cytokine deprivation-induced cell death. Am J Transplant 9, 2281–91. Williams, M. S., Noguchi, S., Henkart, P. S., and Osawa, Y. (1998) Nitric oxide synthase plays a signalling role in TCR-triggered apoptotic death. J Immunol 161, 6526–31. Reiling, N., Kröncke, R., Ulmer, A. J., Gerdes, J., Flad, H.-D., and Hauschildt, S. (1996) Nitric oxide synthase: expression of the endothelial, Ca2+/calmodulin-dependent isoform in human B and T lymphocytes. Eur J Immunol 26, 511–16. Nagy, G., Koncz, A., and Perl, A. (2003) T cell activation-induced mitochondrial hyperpolarization is mediated by Ca2+- and redoxdependent production of nitric oxide. J Immunol 171, 5188–97. Sciorati, C., Rovere, P., Ferrarini, M., Heltai, S., Manfredi, A. A., and Clementi, E. (1997) Autocrine nitric oxide modulates CD95induced apoptosis in gd T lymphocytes. J Biol Chem 272, 23211–5. Degli-Esposti, M. A., and Smyth, M. J. (2005) Close encounters of different kinds: dendritic cells and NK cells take centre stage. Nat Rev Immunol 5, 112–24. Lucas, M., Schachterle, W., Oberle, K., Aichele, P., and Diefenbach, A. (2007) Dendritic cells prime natural killer cells by trans-presenting interleukin-15. Immunity 26, 1–15. Schleicher, U., Liese, J., Knippertz, I., Kurzmann, C., Hesse, A., Heit, A., Fischer, J. A., Weiss, S., Kalinke, U., Kunz, S., and Bogdan, C. (2007) NK cell activation in visceral leishmaniasis requires TLR9, myeloid DCs, and IL-12, but is independent of plasmacytoid DCs. J Exp Med 204, 893–906. Diefenbach, A., Schindler, H., Röllinghoff, M., Yokoyama, W., and Bogdan, C. (1999) Requirement for type 2 NO-synthase for IL-12 responsiveness in innate immunity. Science 284, 951–55. Cifone, M. G., D’Alo, S., Parroni, R., Millimaggi, D., Biordi, L., Martinotti, S., and Santoni, A. (1999) Interleukin-2 activated rat natural killer cells express inducible nitric oxide synthase that contributes to cytotoxic function and interferon-g production. Blood 93, 3876–84. Salvucci, O., Kolb, J. P., Dugas, B., Dugas, N., and Chouaib, S. (1998) The induction of nitric oxide by interleukin-12 and tumor necrosis factor-alpha in human natural killer cells: relationship with the regulation of lytic activity. Blood 92, 2093–102. Furuke, K., Burd, P. R., Horvath-Arcidiacono, J. A., Hori, K., Mostowski, H., and Bloom,

126.

127.

128.

129.

130.

131.

132.

133.

134.

135.

136.

E. T. (1999) Human NK cells express endothelial nitric oxide synthase, and nitric oxide protects them from activation-induced cell death by regulating expression of TNF-a. J Immunol 163, 1473–1480. Diefenbach, A., Schindler, H., Donhauser, N., Lorenz, E., Laskay, T., MacMicking, J., Röllinghoff, M., Gresser, I., and Bogdan, C. (1998) Type 1 interferon (IFN-a/b) and type 2 nitric oxide synthase regulate the innate immune response to a protozoan parasite. Immunity 8, 77–87. Burnett, T. G., and Hunt, J. S. (2000) Nitric oxide synthase-2 and expression of perforin in uterine NK cells. J Immunol 164, 5245–50. Bogdan, C., Röllinghoff, M., and Diefenbach, A. (2000) The role of nitric oxide in innate immunity. Immunol Rev 173, 17–26. Cifone, M. G., Ulisse, S., and Santoni, A. (2001) Natural killer cells and nitric oxide. Int Immunopharmacol 1, 1513–24. Kurose, I., Miura, S., Saito, H., Tada, S., Fukumura, D., Higuchi, H., and Ishii, H. (1995) Rat Kupffer cell-derived nitric oxide modulates induction of lymphokine-activated killer cell. Gastroenterology 109, 1958–68. Ferlito, M., Irani, K., Faraday, N., and Lowenstein, C. J. (2006) Nitric oxide inhibits exocytosis of cytolytic granules from lymphokine-activated killer cells. Proc Natl Acad Sci U S A 103, 11689–94. Jayasekera, J. P., Vinuesa, C. G., Karupiah, G., and King, N. J. (2006) Enhanced antiviral antibody secretion and attenuated immunopathology during influenza virus infection in nitric oxide synthase-2-deficient mice. J Gen Virol 87, 3361–71. Tezuka, H., Abe, Y., Iwata, M., Takeuchi, H., Ishikawa, H., Matsushita, M., Shiohara, T., Akira, S., and Ohteki, T. (2007) Regulation of IgA production by naturally occurring TNF/iNOS-producing dendritic cells. Nature 448, 929–33. Lechner, M., Lirk, P., and Rieder, J. (2005) Inducible nitric oxide synthase (iNOS) in tumor biology: the two sides of the same coin. Semin Cancer Biol 15, 277–89. Singh, R., Pervin, S., Karimi, A., Cederbaum, S., and Chaudhuri, G. (2000) Arginase activity in human breast cancer cell lines: Nw-hydroxyl-arginine selectively inhibits cell proliferation and induces apoptosis in MDA-MB-468 cells. Cancer Res 60, 3305–12. Pamer, E. G. (2009) Tipping the balance in favor of protective immunity during influenza virus infection. Proc Natl Acad Sci U S A 106, 4961–2.

Regulation of Lymphocytes by Nitric Oxide 137. Dalton, D. K., and Wittmer, S. (2005) Nitricoxide-dependent and independent mechanisms of protection from CNS inflammation during Th1-mediated autoimmunity: evidence from EAE in iNOS KO mice. J Neuroimmunol 160, 110–21. 138. Hauser, B., Bracht, H., Matejovic, M., Radermacher, P., and Venkatesh, B. (2005) Nitric oxide synthase inhibition in sepsis? Lessons learned from large-animal studies. Anesth Analg 101, 488–98. 139. Su, F., Huang, H., Kazuki, A., Occhipinti, G., Donadello, K., Piagnerelli, M., De Backer, D., and Vincent, J. L. (2010) Effects of a selective iNOS inhibitor versus norepinephrine in the treatment of septic shock. Shock. Feb 10. [Epub ahead of print]. 140. Heemskerk, S., Masereeuw, R., Russel, F. G., and Pickkers, P. (2009) Selective iNOS inhibition for the treatment of sepsis-induced acute kidney injury. Nat Rev Nephrol 5, 629–40. 141. Lamontagne, F., Meade, M., Ondiveeran, H. K., Lesur, O., and Robichaud, A. E. (2008) Nitric oxide donors in sepsis: a systematic review of clinical and in vivo preclinical data. Shock 30, 653–9.

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142. Valdez, C. A., Saavedra, J. E., Showalter, B. M., Davies, K. M., Wilde, T. C., Citro, M. L., Barchi, J. J., Jr., Deschamps, J. R., Parrish, D., El-Gayar, S., Schleicher, U., Bogdan, C., and Keefer, L. K. (2008) Hydrolytic reactivity trends among potential prodrugs of the O(2)glycosylated diazeniumdiolate family targeting nitric oxide to macrophages for antileishmanial activity. J Med Chem 51, 3961–70. 143. Singh, R., Manjunatha, U., Boshoff, H. I., Ha, Y. H., Niyomrattanakit, P., Ledwidge, R., Dowd, C. S., Lee, I. Y., Kim, P., Zhang, L., Kang, S., Keller, T. H., Jiricek, J., and Barry, C. E., 3rd. (2008) PA-824 kills nonreplicating Mycobacterium tuberculosis by intracellular NO release. Science 322, 1392–5. 144. Deignan, J. L., Livesay, J. C., Yoo, P. K., Goodman, S. I., O’Brien, W. E., Iyer, R. K., Cederbaum, S. D., and Grody, W. W. (2006) Ornithine deficiency in the arginase double knockout mouse. Mol Genet Metab 89, 87–96. 145. Boucher, J. L., Moali, C., and Tenu, J.-P. (1999) Nitric oxide biosynthesis, nitric oxide synthase inhibitors and arginase competition for l-arginine utilization. Cell Mol Life Sci 55, 1015–28.

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Part III Physiopathological Situations

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Chapter 25 Pregnancy: Tolerance and Suppression of Immune Responses Anne Leber, Maria Laura Zenclussen, Ana Teles, Nadja Brachwitz, Pablo Casalis, Tarek El-Mousleh, Federico Jensen, Katja Woidacki, and Ana Claudia Zenclussen Abstract Presence of foreign tissue in a host’s body would immediately lead to a strong immune response directed to destroy the alloantigens present in fetus and placenta. However, during pregnancy, the semiallogeneic fetus is allowed to grow within the maternal uterus due to multiple mechanisms of immune tolerance, which are discussed in this chapter. Key words: Pregnancy, Tolerance, Regulatory T cells, Dendritic cells, Heme oxygenase-1

1. Introduction Presence of foreign tissue in a host’s body would immediately lead to a strong immune response directed to destroy the alloantigens. During pregnancy, however, the semiallogeneic fetus is allowed to grow within the maternal uterus. It is now known that the maternal immune system becomes aware of the presence of the fetus and actively tolerates it. Most interesting, it has been proposed that the fetus itself contributes to his own tolerance. The contradictory nature of the immunological relationship between a pregnant mother and her antigenically foreign child was first recognized by Peter Medawar in 1953 (1), who postulated that (1) the anatomical separation between embryo and mother through the placenta, (2) the antigenic immaturity of the fetus, and (3) the immunological inertness of the mother during pregnancy would ensure the “acceptance” of the growing fetus. However, it is now known that none of these explanations is valid anymore. Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_25, © Springer Science+Business Media, LLC 2011

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Firstly, maternal and fetal cells are in close contact during pregnancy, thus trophoblasts and cells derived from the fetus itself are exposed to maternal immune cells, which have the potential to develop a strong immune response to reject the foreign tissues. Besides, bidirectional transfer of cells between mother and fetus is well documented for human and murine pregnancy ((2–4) and Zenclussen AC, unpublished data). Secondly, the fetus may not be antigenically immature (5). Lastly, there are clear evidences that maternal T-cell recognition of fetal antigens can lead to antigenspecific tolerance (6–8). The fetal–maternal interface can be therefore considered as a site of immune privilege, being the conceptus partially responsible for the establishment of a maternal immune response toward a protective, tolerant one. However, pregnancy complications of immunological origin, such as spontaneous abortion and pre-eclampsia, do exist and may arise from incomplete tolerance. Many mechanisms have been proposed to explain the survival of the semiallogeneic fetus during pregnancy in the context of the active generation of tolerance. In this chapter, we describe cells and molecules known to be the contributors to pregnancy tolerance, particularly emphasizing on regulatory T cells (Treg) and the enzyme heme oxygenase-1 (HO-1).

2. Cell Populations During Pregnancy 2.1. Cells of the Innate Immune System 2.1.1. Antigen-Presenting Cells: Macrophages and Dendritic Cells

The first maternal immune cells to be in contact with foreign antigens from the semen are antigen-presenting cells (APCs) present in the vaginal fluid. Within APCs, dendritic cells (DCs, CD11c+ cells) are the most effective in antigen presentation. We have been able to detect both maternal CD11c+ APCs and paternal antigens in vaginal lavage immediately after plug detection (Zenclussen ML, (9)). This was accompanied by an expansion of regulatory T cells (Treg) which act specifically protecting paternal antigens (Zenclussen ML, (9)). The fact that antigen-specific responses were generated very early in pregnancy defines pregnancy success (Zenclussen ML, in revision, (10, 11)) and speaks strongly in favor of early antigen presentation. Moreover, molecules like galectin-1 and heme oxygenase-1 (HO-1) were found to be involved in the maturation state of DCs and were described to render them tolerogenic (12, 13). We observed that in vivo modulation of HO-1 during pregnancy greatly modifies the maturation state of DCs which in turn help expanding antigen-specific Treg which finally protect the fetus from abortion (Wafula et al. in revision). There are strong evidences that trophoblasts secrete factors which attract monocytes and “educate” them to produce and secrete cytokines and chemokines which support implantation

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and fetal growth (14). In addition, it has been recently described that DCs can interact with uterine natural killer cells (uNKs) and regulate their function at the fetal-maternal interface (15). 2.1.2. Uterine Natural Killer Cells

In murine pregnancy, uNKs make up to 75% of the lymphocytes within the uterus (16). Beginning on day 5 until day 7 of pregnancy, uNKs begin to proliferate, gain size, and accumulate cytoplasmatic granula (17–19). In addition, they start off with the interferon-g (IFNg) production (19). The differentiation of uNKs is indirectly regulated by the hormones estrogen and progesterone (17, 20). On day 8 of pregnancy, the number of uNKs reaches the maximum and after that their number decreases due to apoptosis (17). It is already known that uNKs are important for trophoblast invasion and successful pregnancy outcome. Interleukin-15 (IL-15) is essential for the survival of uNKs and accordingly, IL-15 knockout mice show no formation of uNKs, which is associated with abnormal development of spiral arteries (21). We have been able to observe that upregulation of HO-1 diminishes the abortion rate of DBA/2J-mated CBA/J females which is associated with elevated numbers of uNKs at the fetal–maternal interface (Brachwitz N and Zenclussen AC, unpublished data). In addition, heterozygous or knockout mice for HO-1 which have a poor pregnancy outcome showed a significant reduction of uNKs at the fetal–maternal interface (Brachwitz N and Zenclussen AC, unpublished data). Our data suggest that HO-1 influences the number of uNKs at the fetal–maternal interface either by favoring the migration of NK cells into the fetal–maternal interface or by promoting their survival and proliferation there.

2.1.3. Mast Cells

Mast cells (MCs) are best known for their key effector functions in allergic diseases (22). However, various other immunological but also non-immunological signals can lead to the activation of MCs (23, 24). It is further known that hormones, e.g., estradiol and progesterone are able to modulate MC degranulation (25), therefore a role in reproductive processes would be expected. MCs are also able to produce a variety of preformed and newly synthesized mediators which can induce various pro-inflammatory, anti-inflammatory, and/or immunoregulatory effects (24, 26–28), as well as regulate extracellular matrix (ECM) degradation and tissue remodelling through activation and production of matrix metalloproteinases (29) which play a critical role for the correct embryo implantation. MCs are found in high numbers in myometrium, endometrium, and cervix; they are specially localized around blood vessels (30–32). It has been reported that the number of uterine MCs in mouse and hamster is related to the cyclic phenomenon of estrous being higher in diestrus than in estrus, which can be interpreted as MCs being regulated by endocrine

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mechanisms (33–35). Pregnancy is related with a steady increase in MC density and histamine content (33, 36). It has been proposed that MC degranulation is a necessary process to contribute to normal angiogenesis of the rat cervix during pregnancy (37). The application of a MC-stabilizer (disodium cromoglycate) which inhibits MC degranulation led to decreased endothelial cell proliferation in uterus while reducing VEGF expression (38) (Fig. 1). These observations would predict a positive, even necessary role of MCs for pregnancy to develop normally. Accordingly, Widayati and colleagues described the presence of MCs in large numbers at the site of implantation as well as in myometrium but not in decidua at later stages in normal pregnant mice (39). We observed a large number of MCs in murine decidua at all stages (Popovic M and Zenclussen AC, unpublished data). We further observed diminished implantation rates in C57BL/6J-Kitw-sh/w-sh mice, which lack MCs at reproductive age. This phenotype could be reversed by adoptive transfer of bone-marrow-derived MCs. Our results suggest that MCs and their released mediators exert positive influence with regard to nidation of the blastocyst in female and seem to be essential for a successful pregnancy outcome (Popovic M and Zenclussen AC, unpublished data). This may be mainly due to their crucial function in tissue remodelling processes and angiogenesis in which molecules such as uPA, VEGF, and MMP-9 are involved (Woidacki K, Popovic M, and Zenclussen AC, unpublished data). A further studied role for MCs is their contribution to uterine contractility in pregnancy (40). Release of MC mediators in the uterus is a stimulus to trigger and/or maintain myometrial contractions during pre-term and term labor (41–43). However, mastocytosis does not seem to imply any risk for pregnant women during labor and delivery, and their newborns were shown to be healthy (44). Finally, placental MCs were also proposed to release vasoconstrictive substances which could in turn impair blood flow and result, e.g., in growth restriction in fetuses of women with type I hypersensitivity reactions (41). No reports exist in the literature indicating a possible role for MCs in pregnancy-induced tolerance. MCs emerge now with a positive

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Extracellular matrix degradation VEGF Angiogenesis + Blastocyst Histamine implantation Tissue remodelling uPA

Fig. 1. Mediators released by activated mast cells are crucial for extracellular matrix degradation, angiogenesis, and tissue remodelling, processes that are essential for a successful implantation of the blastocyst in the endometrium of the uterus.

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physiological role, ensuring self-tolerance and/or tissue integrity through the maintenance of “privileged” microenvironments where adaptive immune responses are damped down (45). In 2006, Lu and colleagues described MCs as essential intermediaries in regulatory T-cell-dependent allograft tolerance (46). We were able to confirm that in the CBA × DBA model, the transfer of Treg is associated with an increase in the number of MCs as well as in the expression of MC-associated genes (Popovic M and Zenclussen AC, unpublished data). To understand the mechanisms as to why MCs reach the pregnant uterus and the placenta in humans, we performed a migration assay and analyzed the ability of HMC-1 MCs to migrate toward primary human trophoblast cells or choriocarcinoma cells (JEG-3 cells). We confirmed that MCs strongly migrate to both human primary trophoblast cells and JEG-3 cells. As for the molecules implied, we could confirm that CCR4, CCR5, and CXCR4 expressed in HMC-1 cells are involved in the migration of MCs toward CCL11, CCL14, CCL16, and CCL22 secreted by uterine cells upon hormone variations (Jensen F and Zenclussen AC, unpublished data). Physiological concentrations of estradiol and progesterone as observed, e.g., during the menstrual cycle or in the beginning of pregnancy led to MC degranulation while they significantly upregulated the expression of CCR4, CCR5, and CXCR4 in HMC-1 cells. The modulatory effects of estradiol on chemokine receptor expression clearly bring to light a novel mechanism as to how MCs may migrate to the uterus and fetal– maternal interface (Jensen F and Zenclussen AC, unpublished data). Estradiol and progesterone increase chemokines secreted from the uterus, which then modulate the expression of the respective chemokine receptors in MCs, ending in the migration of MCs to the uterus. These hormones further induce MC maturation, which help preparing the uterus for a possible implantation by upregulating the levels of the metalloprotease MMP-9 and tryptase (Fig. 2). 2.2. Cells of the Adaptive Immune System 2.2.1. B Cells, Asymmetric Antibodies, and T Cells

Little is known about the contribution of B cells and their products, antibodies, to pregnancy success. Besides acting as APCs, B cells get activated after antigen presentation and become plasma cells, which produce antibodies. Anti-paternal antibodies have been described being present in both maternal sera and at the fetal-maternal interface (47–49). Different studies suggest that development of alloantibodies may be a normal requirement for successful pregnancy, e.g., to inhibit NK cytotoxicity against trophoblast cells (50, 51). Because of their molecular asymmetry, these so-called asymmetric antibodies (AAbs) are not able to provoke immune reactions which destroy antigens (52). During pregnancy, they were found to be elevated in the sera as well as in the placenta, where they were blocking paternal antigens specifically

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Fig. 2. Possible scenario for mast cell migration, maturation, and degranulation under hormone influence.

and found to participate in complex immune mechanisms for the protection of the fetus (53–57). Soluble factors (e.g. cytokines) secreted by human, mouse, and rat placentas were able to enhance AAb synthesis in vivo and in vitro (57, 58). In this regard, IL-6 augments AAb production in a murine hybridoma and participates in vivo and in vitro in the “abnormal” glycosylation of AAbs (59, 60). In human pregnancies, low IL-6 levels were associated with high AAbs levels and normal pregnancy, while high IL-6 levels were accompanied by low AAbs and miscarriage (54, 61). In the last decades, many investigations have focused on the production of cytokines by T cells from pregnant woman, even though it has been hotly discussed whether pregnancy can be seen as an antigen-specific response. The “Th1/Th2/Th3 paradigm” proposed that a balance between Th1 and Th2/Th3 cytokines is critical for pregnancy development. Th2- and Th3-type cytokines such as IL-4, IL-10, and TGF-b were suggested to favor the maintenance of mammalian pregnancy (62), whereas the excessive production of Th1 cytokines (IL-2, IFN-g, and TNF-a) would mediate the rejection of the fetus at the fetal–maternal interface (63–65). However, several studies already showed that mice which are deficient for Th2 cytokines have normal pregnancies (66, 67). Contrary to these evidences, there are strong evidences that IL-4 and IL-10 contribute to successful pregnancy outcome in human (reviewed in (68, 69)). Other immunosuppressive molecules produced by T-helper cells, especially TGF-b have been proposed

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to play an important role during pregnancy (70, 71). There is clear evidence of complex selective mechanisms for regulating leukocyte access to the fetal–maternal interface (72). On the assumption that miscarriage is a Th1 event and that selectins are expressed at the murine fetal–maternal interface by the time of rejection, we investigated whether blocking P- and E-selectins before implantation would inhibit the migration of immune cells into the fetal– maternal interface and thus prevent fetal rejection in a murine model of spontaneous abortion. We observed that blocking the initial step of lymphocytes extravasation via anti-P-selectin mAb positively influences pregnancy outcome by significantly diminishing the abortion rate (73). Interestingly, decidual lymphocytes were predominantly of the Th2 type, suggesting a selective blockage of Th1 cells (73). There are clear evidences of augmented number of Th1 cells at the fetal–maternal interface from mice and human during miscarriage. However, it is still a matter of discussion whether these cells are reactive against paternal antigens or not. The first evidences of an increased Th1 response toward male antigens were provided by Tangri and Raghupathy (74). In a more recent study, we confirmed the existence of specific regulatory processes, which control Th1 cells specific against paternal alloantigen (75). In addition, the transfer of activated Th1 cells into pregnant recipients leads to Preeclampsia-like symptoms (76). Recently, another subset of T-helper cells has been discovered and named as Th17 cells because of their ability to produce IL-17 (77, 78). IL-17 is a proinflammatory cytokine which was described to be important for immunity against extracellular bacteria (77, 78). It stimulates the production of IL-6 (79) which has been shown to be elevated in women with reproductive failure (80). Moreover, together with other inflammatory cytokines such as TNF-a and IFN-g, IL-17 amplifies local inflammation and might therefore favor fetal rejection (81). In transplantation settings, it has been shown that renal and lung allograft rejection is associated with elevated IL-17 levels (82–84). Thus, it is tempting to speculate that IL-17 produced by Th17 cells is involved in fetal loss. On the other hand, it has been demonstrated that IL-17 and IL-10 which are produced by Th17 cells suppress proliferation and cytokine production by Th1 cells (85) which might support a successful pregnancy outcome. These data suggest an ambiguous role for Th17 cells in pregnancy which has to be elucidated in the future. 2.2.2. Regulatory T Cells in Pregnancy

CD4+CD25+ Treg were first described as a specialized subpopulation of T cells responsible for the maintenance of immunological self-tolerance by actively suppressing self-reactive lymphocytes (85). They were confirmed to play a major role in preventing autoimmunity and tolerating allogeneic organ grafts (86, 87). Being specifically expressed in mouse CD4+ Treg, the forkhead box

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transcription factor Foxp3 acts as a master switch in the regulation of their development and function (88–90). Several types of Treg were described from which the majority is naturally produced in the thymus in a continuous mode expressing both glucorticoidinduced TNF receptor family related gene (GITR) and cytotoxic T lymphocyte-associated antigen 4 (CTLA-4) and produce soluble TGF-b and IL-10 (91). Two other subtypes (Tr1 and Th3 cells) are generated extra-thymically and develop from naïve T cells after exposure to antigens in the periphery (92). During human and murine pregnancy, an expansion of Treg can be observed already at very early pregnancy stages and has been confirmed to be essential for a normal pregnancy outcome (75, 93, 94). Using the well-established murine abortion-prone model, we observed that the number of Treg in DBA/2J-mated CBA/J females was significantly diminished compared to BALB/ c-mated CBA/J females. By adoptive transfer of Treg isolated from thymus and spleen of normal pregnant females into abortion-prone females on days 0–2 of pregnancy, we were able to prevent abortion completely (75). As only Treg obtained from pregnant but not from non-pregnant animals were capable in preventing abortion, we propose that Treg from both normal pregnant and non-pregnant mice are able to infiltrate the fetal–maternal interface but only Treg isolated from pregnant females could ensure total fetal protection which suggests that Treg need to be previously exposed to paternal alloantigens in order to gain protective regulatory activity in vivo (75). This could be confirmed by an adoptive transfer of Treg obtained from a third party combination which was not able to protect the fetus (95). Moreover, since Treg transfer from normal pregnant mice failed to prevent abortion when made on days 4–5 of pregnancy and anti-CD25 applied during day 0–2 resulted in implantation failure, this may imply that Treg are necessary locally before implantation (75). Supposing that paternal antigens are necessary for Treg expansion and protective function, we next aimed to investigate the origin of the paternal antigens. We found paternal antigens being present very early in murine pregnancy suggesting that Treg are exposed to paternal antigens at very early pregnancy stages which might explain the increase of Treg observed at this early time point (9). We speculated that paternal antigens coming from the semen meaning sperms or seminal fluid might be responsible for Treg expansion. Using different pairing combinations in which CBA/J females were mated either with intact BALB/c males, vasectomized BALB/c (lacking sperms) or BALB/c without seminal vesicles (lacking seminal fluid), we showed that both sperms and components of the seminal fluid are essential for Treg expansion, confirming our hypothesis of a paternal antigen-driven Treg expansion (Leber A, unpublished data). Our data are supported

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by studies from Sarah Robertson and colleagues which also showed that seminal fluid drives Treg expansion (10). Although it is known that Treg are present in decidual tissue already very early in pregnancy, the mechanisms by which Treg get there are still under investigation. Knowing that IL-2, which is essential for Treg survival and proliferation in vivo, is not available at the fetal–maternal interface in mice and humans (96–98), we hypothesized that Treg need to migrate to the decidua after generation or expansion in draining lymphoid tissue. For migration studies of Treg into the fetal–maternal interface, we first concentrated on chemokines, known to be potent attractors of lymphocytes into tissues. We found chemokine receptors being expressed on the surface of Treg which ligands are present at the fetal–maternal interface. Using in vitro migration assays, we were able to show that Treg are attracted by some of these ligands suggesting that chemokines might be involved in Treg migration into the fetal–maternal interface (Teles A, unpublished data). In addition, we investigated the ability of the pregnancy hormone human chorionic gonadotropin (hCG) to attract Treg knowing that this hormone is expressed at the fetal–maternal interface at very early pregnancy stages. Having confirmed the presence of LH/CG receptors on the surface of Treg, we went on using migration assays to show that both hCG-producing JEG-3 cells and first trimester trophoblasts efficiently attract Treg. In contrast, non-hCG-producing cells like keratinocytes (HaCat cells) could not attract Treg. Our observation of hCG-dependent Treg migration was confirmed by a transfection of non-hCG-producing colon carcinoma cells with hCG vectors (99). Our data clearly show that beside chemokines, hCG is one of the attractor of Treg into the fetal– maternal interface. Treg have been shown to secrete IL-10 and TGF-b and thereby suppress the effector functions of activated leukocytes (100, 101). The blockage of IL-10 but not TGF-b after adoptive transfer of Treg into abortion-prone mice was able to abrogate the Treg protective effect (95), suggesting that IL-10 but not TGF-b might be one of the molecules by which Treg mediate their protective function during pregnancy. Moreover, blockage of CTLA-4 or PD-1 which also have been described to play a role for Treg function led to an abolishment of Treg protection (102). Further data suggest that Treg protect the allogeneic fetus by creating a “tolerant” privileged microenvironment at the fetal–maternal interface characterized by high expression of HO-1, TGF-b, and leukemia inhibitory factor (LIF) (103). Figure 3 shows a hypothetical scenario, which summarizes all data obtained by our working group regarding generation, migration, and function of Treg during pregnancy.

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Fig. 3. Hypothetical scenario on generation, migration, and function of Treg during pregnancy.

3. Molecules Dictating Pregnancy Success 3.1. Molecules Implied in Implantation

Several molecules were described to play an important role during the pre-implantation and implantation period of pregnancy. In this regard, the cytokine LIF has been shown to be highly and specifically expressed on day 4 of pregnancy and its expression coincides with the regulation of the growth and implantation of blastocysts (104). It has been shown that animals lacking the LIF gene produce normal blastocysts that fail to implant into LIF-deficient uteri but are capable to implant in a wild-type uterus (105). It is also known that intraperitoneal injection of LIF on day 4 of pregnancy into LIF−/− mice restored implantation (106). In addition, in human pregnancy, a decreased production of LIF in the uterine microenvironment was found in states of impaired fertility (107). Wenwei and colleagues showed that expression of LIF is regulated by the tumor suppressor p53. Lower LIF levels were observed in the uteri of p53 KO mice than those of p53 wild-type mice, particularly at day 4 of pregnancy when transiently induced high LIF levels are crucial for embryonic implantation. The same research group observed a significant decrease in embryonic implantation, pregnancy rate, and litter size in matings with p53−/− females (108). Altogether these results suggest a function for p53 in maternal reproduction through the regulation of LIF (108). The immune suppressive cytokine TGF-b has been shown to be present in male seminal fluids. Immediately after insemination,

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TGF-b is activated in the female reproductive tract which leads to an immune activation necessary for implantation. (109). In addition, HO-1 has been shown to be involved in implantation processes and formation of the placenta. Recently, Zhao and colleagues found that HO-1 heterozygote cross-breedings have an extremely low birth rate in HO-1 homozygote (HO-1−/−) offspring (2.4%) and small litter sizes were observed. Placentas and fetuses from heterozygote cross-breedings were relatively smaller and weighed less than those from wild-type cross-breedings. Having these results, Zhao and colleagues concluded that a partial deficiency in HO-1 is associated with morphological changes in the placenta (110). 3.1.1. Heme Oxygenase-1 in Pregnancy

In 1964, Wise and Drabkin (111) first described an enzymatic reaction that converted heme to biliverdin and carbon monoxide. They described that this reaction required NAD and ATP, and that the enzyme source was the light mitochondrial fraction of hemophagous organ of the dog placenta. This report was followed by reports of Tenhunen and colleagues (112–114) who described a bile-pigment producing system located in the microsomal fraction of the rat liver, and they named it “microsomal heme oxygenase.” Nowadays it is widely know that heme oxygenase (HO), encoded by the Hmox1 gene, is the enzyme catalyzing the first and rate limiting step in the degradation of heme, to yield equimolecular quantities of biliverdin, CO, and free iron. Biliverdin is then converted to bilirubin via the action of biliverdin reductase, and free iron is sequestered into ferritin (reviewed in (115)). To date, three different mammalian isoforms have been identified: HO-1, HO-2, and HO-3 (116–118). HO-1, also known as heat-shock protein (HSP) 32, is very sensitive to several stimuli and agents that cause oxidative stress and pathological conditions, such as heat shock, ischemia, radiation, hypoxia, hyperoxia (119), cytokines (IL-1, IL-6, or TNF-a), heavy metals, and nitric oxide. Beneficial effects of HO-1 have been described in different fields of medicine, such as transplantation (120–123), atherosclerosis (124), sepsis (125), autoimmune neuroinflammation (126), and infections such as malaria (127). At the fetal–maternal interface, inflammatory processes can occur due to the invasion of microorganisms, but also due to an immune reaction against alloantigens on the fetus or trophoblast. Many studies in animals and humans indicate that some degree of systemic or uterine inflammation is necessary for both normal implantation and pregnancy, but if this inflammation becomes too excessive it can cause pregnancy complications such as abortion (reviewed in (128)). As pointed out in previous reports, HO-1 plays a key role in inflammatory processes (reviewed in (129)). Pregnancy is viewed by many from a histoincompatibility point of view. Many propose that the histoincompatibility between maternal and paternal antigens provokes a semiallogeneic fetus as

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one of the main causes for fetal acceptance or rejection. In this regard, it is thought that an exacerbated immunological reaction toward the fetus may provoke an immunological misbalance that would lead to fetal rejection. According to this, fetal rejection mechanisms would resemble the mechanisms involved in the rejection of an allograft. In the transplantation field, HO-1 has also been shown by others to play an important role and to have beneficial effects when upregulated (120–122). Considering the important role of HO-1 in inflammatory processes as well as in transplantation, and regarding the resemblance of both entities to different stages of pregnancy, we aimed to analyze the role of HO-1 in the different processes related to pregnancy by means of functional studies employing in vivo as well as in vitro models. Using an in vivo approach, we aimed to clarify whether an upregulation of HO-1 may be able to rescue mice from spontaneous abortion using a well-established abortion model. The HO-1 upregulation was performed either chemically by Cobalt protoporphyrin (CoPPIX) or specifically by applying an adenoviral vector containing HO-1. We could show that a specific increase in the systemic HO-1 expression following gene transfer of 1 × 105 PFU AdHO-1/GFP improves pregnancy outcome by diminishing fetal rejection rate (130). We also induced HO-1 upregulation by means of CoPPIX and observed diminished abortion rates as well. Interestingly, the application of Zinc protoporphyrin (ZnPPIX), known to downregulate HO-1, provoked a significant augmentation in the abortion rates (131). Regarding the mechanisms by which the adenoviral vector containing HO-1 is pregnancy-protective in this model, one of them seems to be a Th2 polarization. In this work, augmented levels of IL-4 as well as diminished levels of IFN-g were found systemically and locally after the application of AdHO-1/GFP. We have shown that mice that were rescued from abortion by the adoptive transfer of Treg presented augmented levels of HO-1 at the fetal–maternal interface (102). This suggests that Treg are able to induce HO-1 expression at the fetal–maternal interface. On the other hand, we could also show that HO-1 upregulation by means of CoPPIX augmented the levels of the Treg marker Neuropilin-1 (131), suggesting a bidirectional relationship between HO-1 and Treg in pregnancy. HO-1 is also known to have anti-apoptotic properties (120). We observed a significant diminution in the number of apoptotic cells in placenta from mice receiving low doses of AdHO-1/GFP when compared to PBS- or EGFP-treated abortion-prone mice, confirming therefore the anti-apoptotic effect of the HO-1-therapy at the fetal–maternal interface. Accordingly, we found augmented mRNA levels of the anti-apoptotic molecule Bag-1 (130, 131) which reinforces an anti-apoptotic/cytoprotection hypothesis. Previous studies from our group and from others support the concept that, during pregnancy, different types of trophoblast

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cells are important sources of HOs (132–136) and would participate in the catabolism of the heme proteins, avoiding accumulation or recirculation of free heme, which could be extremely toxic for the mother and for the fetus. After implantation, the trophectoderm surrounding the blastocyst goes on to differentiate into a variety of trophoblast cell subtypes with different functions. Trophoblast stem cells emerge from the polar trophectoderm that overlies the inner cell mass of the blastocyst, and they proliferate in response to close contact to the inner cell mass (136). From the different types of trophoblasts in the mouse, the giant cells are the first to arise, already at the blastocyst stage. The function of trophoblast giant cells is to first mediate the process of implantation and invasion of the conceptus into the uterus. Later, they produce several hormones and cytokines to promote local and systemic physiological adaptations in the mother (reviewed in (136)). Due to the vital importance of the giant cells in the formation of the placenta and in the establishment of a successful pregnancy, we also analyzed in vitro whether HO-1 is necessary for trophoblast stem cell differentiation into giant cells. The Rcho-1 cell line is usually used as an in vitro model for studying trophoblast cell differentiation (137). These cells can be manipulated to proliferate or differentiate into trophoblast giant cells by altering culture conditions. Rcho-1 trophoblast stem cell differentiation recapitulates in vivo trophoblast giant cell development, and is a valuable in vitro tool for studying the process of trophoblast cell differentiation (138). The downregulation of HO-1 expression by ZnPPIX led already to a significantly diminished viability of precursor stem cells, suggesting that HO-1 is necessary even for the survival of trophoblast stem cells. When analyzing the ability of these cells to differentiate into giant cells, the application of ZnPPIX led them unable to transform into giant cells, whereas cells treated with CoPPIX did not show any problem in the differentiation process. These results suggest that HO-1 may play an important role in the differentiation process of trophoblast cells (Zenclussen ML, unpublished data). No results have been found in the literature on this regard, making this a novel discovery in the role of HO-1 in formation of placenta. The fact that mice lacking Hmox1 do not get pregnant also suggests that without HO-1 no placentation is able to occur. As already described by others (139, 140), no progeny can be achieved when mating Hmox1−/− mice. Interestingly, the mating of heterozygous females and males leads to only 3–10% of knockout progeny, instead of the 25% expected Mendelian rate. Both facts suggest that Hmox1 plays a very important role in pregnancy, but so far no explanation for this problem in the maintenance of the colony was found in the literature. Preliminary work form our group mating different combinations of Hmox1+/+, Hmox1+/−, and Hmox1−/− females and males led

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us to hypothesize that HO-1 is very important in the maternal site for a successful pregnancy, whereas its absence in the paternal site may not be as important as like in the maternal site (Zenclussen ML, unpublished data). Further data from our group suggest that Hmox-1 may also be necessary (but not indispensable) for egg fertilization (Zenclussen ML, unpublished data). So far, HO-1 in pregnancy was thought to be important for implantation. We could confirm that Hmox-1−/− females are not able to produce as many oocytes as wild-type females, indicating that HO-1 is necessary even at very early stages of reproduction. The difference in the oocyte production may be due to differences in the reaction of the Hmox1−/− females to hormonal treatment, since ovaries from Hmox1+/+ and Hmox1−/− females showed the same total number of follicles per ovary (Zenclussen ML, unpublished data). The changes that take place on the ovary at the site of follicular rupture are pathophysiological in nature. This local damage induces hemorrhage in the vicinity of the lesion on the surface of the ovary (reviewed in (141)). Considering the importance of the upregulation of HO-1 in inflammatory processes, especially in events related to hemorrhage, it is tempting to speculate that the lack of HO-1 may impair the ovulation process in a way that these animals do not manage to induce the follicular rupture. Without HO-1, the ability to counteract the inflammatory process related to it is missing. In other words, due to the inflammatory nature of ovulation, mature follicles lacking HO-1 may find it more difficult to induce the rupture of the tissue due to the inflammatory nature of the process and may be then less secreted than follicles expressing HO-1 (Zenclussen ML and Casalis PA, unpublished data). Interestingly, NO, an inflammatory mediator normally associated with HO-1, has been described to be involved in ovulation. Studies using eNOS−/− mice have shown that they have reduced fertility due to impaired ovulatory efficiency, abnormalities in meiotic maturation, increased oocyte apoptosis, and altered estrous cyclicity compared to their wild-type littermates (142–144). Studies using iNOS−/− mice show that iNOS deficiency does not alter ovulatory capacity, but it may play a role in fertilization (145). Furthermore, the cyclooxygenase (COX) pathway, which is responsible for prostaglandin (PG) synthesis, is also important in inflammatory responses, as the HO and NO pathways are. In addition, they have constitutive and inducible isoforms (146), and they are closely related (147). The two enzymes responsible for the synthesis of PGs are COX1 and COX2, being the first constitutive and the second inducible form (148). Although Cox1−/− mice have normal fertility, Cox2−/− females are infertile and exhibit abnormalities in ovulation due to PG deficiency (149). Regarding the HO system, interestingly, Hmox2−/− mice do not present reproductive problems (150), whereas Hmox1−/− do present problems,

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as previously described. These results open the possibility to analyze further the problem in a more accurate way, and these studies are being currently done in our group. In summary, we present in this review several important pathways for normal pregnancy outcome, which are probably all working together in synergy to ensure that the maternal immune system supports fetal growth and development. References 1. Medawar PB (1953) Some immunological and endocrinological problems raised by evolution of viviparity in vertebrates. In: Symposia of the society for experimental biology, vol 7, Evolution, R. Brown and JF Danielli, Eds. Syndics of the Cambridge University Press, London, pp. 320–338 2. Yan Z, Lambert NC, Guthrie KA, Porter AJ, Loubiere LS et al (2005) Male microchimerism in women without sons: quantitative assessment and correlation with pregnancy history. Am J Med 118:899–906 3. Khosrotehrani K, Johnson KL, Guégan S, Stroh H, Bianchi DW (2005) Natural history of fetal cell microchimerism during and following murine pregnancy. J Reprod Immunol 66:1–12 4. Tan XW, Liao H, Sun L, Okabe M, Xiao ZC et al (2005) Fetal microchimerism in the maternal mouse brain: a novel population of fetal progenitor or stem cells able to cross the bloodbrain barrier? Stem Cells 23:1443–1452 5. Elbe-Bürger A, Mommaas AM, Prieschl EE, Fiebiger E, Baumruker T, Stingl G (2000) Major histocompatibility complex class II – fetal skin dendritic cells are potent accessory cells of polyclonal T-cell responses. Immunol 101(2):242–253 6. Tafuri A, Alferink J, Möller P, Hämmerling GJ, Arnold B (1995) T cell awareness of paternal alloantigens during pregnancy. Science 270(5236):630–633 7. Jiang SP, Vacchio MS (1998) Multiple mechanisms of peripheral T cell tolerance to the fetal “allograft”. J Immunol 160(7):3086– 3090 8. Moldenhauer LM, Diener KR, Thring DM, Brown MP, Hayball JD, Robertson SA (2009) Cross-presentation of male seminal fluid antigens elicits T cell activation to initiate the female immune response to pregnancy. J Immunol 182(12):8080–8093 9. Zenclussen ML, Thuere C, Ahmad N, Wafula PO, Fest S, Teles A, Leber A, Casalis PA, Bechmann I, Priller J, Volk HD, Zenclussen

10.

11.

12.

13.

14.

15.

16.

AC (2010) The persistence of paternal antigens in the maternal body is involved in regulatory T-cell expansion and fetal-maternal tolerance in murine pregnancy. Am J Reprod Immunol 63(3):200-208 Robertson SA, Guerin LR, Bromfield JJ, Branson KM, Ahlström AC, Care AS (2009) Seminal fluid drives expansion of the CD4+CD25+ T regulatory cell pool and induces tolerance to paternal alloantigens in mice. Biol Reprod 80(5):1036–1045 Tilburgs T, Roelen DL, van der Mast BJ, de Groot-Swings GM, Kleijburg C, Scherjon SA, Claas FH (2008) Evidence for a selective migration of fetus-specific CD4+CD25 bright regulatory T cells from the peripheral blood to the decidua in human pregnancy. J Immunol 180(8):5737–5745 Ilarregui JM, Croci DO, Bianco GA, Toscano MA, Salatino M, Vermeulen ME, Geffner JR, Rabinovich GA (2009) Tolerogenic signals delivered by dendritic cells to T cells through a galectin-1-driven immunoregulatory circuit involving interleukin 27 and interleukin 10. Nat Immunol 10(9):981–991 Rémy S, Blancou P, Tesson L, Tardif V, Brion R, Royer PJ, Motterlini R, Foresti R, Painchaut M, Pogu S, Gregoire M, Bach JM, Anegon I, Chauveau C (2009) Carbon monoxide inhibits TLR-induced dendritic cell immunogenicity. J Immunol 182(4):1877–1884 Fest S, Aldo PB, Abrahams VM, Visintin I, Alvero A, Chen R, Chavez SL, Romero R, Mor G (2007) Trophoblast-macrophage interactions: a regulatory network for the protection of pregnancy. Am J Reprod Immunol 57(1):55–66 Karsten CM, Behrends J, Wagner AK, Fuchs F, Figge J, Schmudde I, Hellberg L, Kruse A (2009) DC within the pregnant mouse uterus influence growth and functional properties of uterine NK cells. Eur J Immunol 39(8):2203–2214 Murphy SP, Fast LD, Hanna NN, Sharma S (2005) Uterine NK cells mediate inflammation

412

17. 18.

19.

20.

21.

22. 23.

24.

25.

26.

27.

28.

29.

Zenclussen et al. induced fetal demise in IL-10-null mice. J Immunol 175:4084–4090 Peel S (1989) Granulated metrial gland cells. Adv Anat Embryol Cell Biol 115:1–112 Parr EL, Parr MB, Zheng LM, Young JD (1991) Mouse granulated metrial gland cells originate by local activation of uterine natural killer lymphocytes. Biol Reprod 44:834–841 Croy BA, Ashkar AA, Foster RA, DiSanto JP, Magram J, Carson D, Gendler SJ, Grusby MJ, Wagner N, Muller W, Guimond MJ (1997) Histological studies of geneablated mice support important functional roles for natural killer cells in the uterus during pregnancy. J Reprod Immunol 35:111–133 Sharma R, Bulmer D, Peel S (1986) Effects of exogenous progesterone following ovariectomy on the metrial glands of pregnant mice. J Anat 144:189–199 Greenwood JD, Minhas K, Di Santo JP, Makita M, Kiso Y, Croy BA (2000) Ultrastructural studies of implantation sites from mice deficient in uterine natural killer cells. Placenta 21:693–702 Metzger H (1992) The receptor with high affinity for IgE. Immunol Rev 125:37–48 Gilfillan AM, Tkaczyk C (2006) Integrated signalling pathways for mast-cell activation. Nat Rev Immunol 6:218–230 Galli SJ, Nakae S, Tsai M (2005) Mast cells in the development of adaptive immune responses. Nat Immunol 6:135–142 Zaitsu M, Narita A, Lambert KC, Grady JJ, Estes DM, Curran EM, Brooks EG, Watson CS, Goldblum RM, Midoro-Horiuti T (2007) Estradiol activates mast cells via a non-genomic estrogen receptor-alpha and calcium influx. Mol Immunol 44(8):1977–1985 Grimbaldeston MA, Metz M, Yu M, Tsai, M, Galli SJ (2006) Effector and potential immunoregulatory roles of mast cells in IgEassociated acquired immune responses. Curr Opin Immunol 18:751–760 Theoharides TC, Kalogeromitros D (2006) The critical role of mast cells in allergy and inflammation. Ann N Y Acad Sci 1088:78–99 Boesiger J, Tsai M, Maurer M, Yamaguchi M, Brown LF, Claffey KP, Dvorak HF, Galli SJ (1998) Mast cells can secrete vascular permeability factor/vascular endothelial cell growth factor and exhibit enhanced release after immunoglobulin E-dependent upregulation of Fce receptor I expression. J Exp Med 188:1135–1145 Baram D, Vaday GG, Salamon P, Drucker I, Hershkoviz R, Mekori YA (2001) Human mast cells release metalloproteinase-9 on contact with activated T cells: juxtacrine reg-

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

ulation by TNF-alpha. J Immunol 167(7): 4008–4016 Jeziorska M, Salamonsen LA, Woolley DE (1995) Mast cell and eosinophil distribution and activation in human endometrium throughout the menstrual cycle. Biol Reprod 53(2):312–320 Mori A, Zhai YL, Toki T, Nikaido T, Fujii S (1997) Distribution and heterogeneity of mast cells in the human uterus. Hum Reprod 12(2):368–372 Cabanillas-Saez A, Schalper JA, Nicovani SM, Rudolph MI (2002) Characterization of mast cells according to their content of tryptase and chymase in normal and neoplastic human uterine cervix. Int J Gynecol Cancer 12(1):92–98 Padilla L, Reinicke K, Montesino H, Villena F, Asencio H, Cruz M, Rudolph MI (1990) Histamine content and mast cells distribution in mouse uterus: the effect of sexual hormones, gestation and labor. Cell Mol Biol 36(1):93–100 Rudolph MI, de los Angeles García M, Sepulveda M, Brandan E, Reinicke K, Nicovani S, Villan L (1997) Ethodin: pharmacological evidence of the interaction between smooth muscle and mast cells in the myometrium. J Pharmacol Exp Ther 282(1):256–261 Harvey EB (1964) Mast cell distribution in the uterus of cycling and pregnant hamsters. Anat Rec 148:507–516 Tabb NT (1994) Immune control of myometrial contractility: Role of mast cells. In: Control of uterine contractility, RE Garfield, NT Tabb, Eds. CRC Press, Boca de Raton, FL, pp. 355–373 Varayoud J, Ramos JG, Bosquiazzo VL, Muñoz-de-Toro M, Luque EH (2004) Mast cells degranulation affects angiogenesis in the rat uterine cervix during pregnancy. Reproduction 127(3):379–387 Bosquiazzo VL, Ramos JG, Varayoud J, Muñoz-de-Toro M, Luque EH (2007) Mast cell degranulation in rat uterine cervix during pregnancy correlates with expression of vascular endothelial growth factor mRNA and angiogenesis. Reproduction 133(5):1045–1055 Widayati DT, Ohmori Y, Fukuta KJ (2004) Distribution patterns of immunocompetent cells in the pregnant mouse uteri carrying allogeneic mouse and xenogeneic vole embryos. J Anat 205(1):45–55 Garfield RE, Irani AM, Schwartz LB, Bytautiene E, Romero R (2006) Structural and functional comparison of mast cells in the pregnant versus nonpregnant human uterus. Am J Obstet Gynecol 194(1):261–267

Pregnancy: Tolerance and Suppression of Immune Responses 41. Bytautiene E, Vedernikov YP, Saade GR, Romero R, Garfield RE (2008) IgEindependent mast cell activation augments contractility of nonpregnant and pregnant guinea pig myometrium. Int Arch Allergy Immunol 147(2):140–146 42. Walsh LJ, Trinchieri G, Waldorf HA, Whitaker D, Murphy GF (1991) Human dermal mast cells contain and release tumor necrosis factor alpha, which induces endothelial leukocyte adhesion molecule 1. Proc Natl Acad Sci USA 88(10):4220–4224 43. Schmith A, Goepfert C, Feitsma K, Buddecke E (2002) Lovastatin-stimulated superinduction of E-selectin, ICAM-1 and VCAM in TNF alpha-activated human vascular endothelial cells. Atherosclerosis 164(1):57–64 44. Worobec AS, Akin C, Scott LM, Metcalfe DD (2000) Mastocytosis complicating pregnancy. Obstet Gynecol 95(3):391–395 45. Waldmann H (2006) Immunology: protection and privilege. Nature 442(7106):987–988 46. Lu L-F, Lind EF, Gondek DC, Bennett KA, Gleeson MW, Pino-Lagos K, Scott ZA, Coyle AJ, Reed JL, Snick JV, Strom TB, Zheng XX, Noelle RJ (2006) Mast cells are essential intermediaries in regulatory T-cell tolerance. Nature 31:997–1002 47. Mowbray JF, Underwood JL (1985) Immunology of abortion. Clin Exp Immunol 60:1–7 48. Innes A, Cunningham C, Power DA, Catto GRD (1989) Fetus as an allograft: noncytotoxic maternal antibodies to HLA-linked paternal antigens. Am J Reprod Immunol 19(4):146–150 49. Malan Borel I, Gentile T, Angelucci J, Pividori J, Margni R (1991) IgG asymmetric molecules with anti-paternal activity isolated from sera and placenta of pregnant human. J Reprod Immunol 20(2):129–140 50. Jalali GR, Rezai A, Underwood JL, Mowbray JF, Surridge SH, AllenWR, Matthias S (1995) An 80-kDa syncytiotrophoblast alloantigen bound to maternal alloantibody in term placenta. Am J Reprod Immunol 33:213–220 51. Jalali GR, Arck P, Surridge S, Markert U, Chaouat G, Clark DA, Underwood JL, Mowbray JF (1996) Immunosuppressive properties of monoclonal antibodies and human polyclonal alloantibodies to the R80K protein of trophoblast. Am J Reprod Immunol 36:129–134 52. Margni RA, Malan Borel I (1998) Paradoxical behavior of asymmetric IgG antibodies. Immunol Rev 163:77–87

413

53. Malan Borel I, Gentile T, Angelucci J, Pividori J, Guala MC, Binaghi RA, Margni RA (1991) IgG asymmetric molecules with antipaternal activity isolated from sera and placenta of pregnant human. J Reprod Immunol 20(2):129–140 54. Zenclussen AC, Gentile T, Kortebani G, Mazzolli A, Margni R (2001) Asymmetric antibodies and pregnancy. Am J Reprod Immunol 45(5):289–294 55. Gentile T, Malan Borel I, Angelucci J, Miranda S, Margni R (1992) Preferential synthesis of asymmetric antibodies in rat immunized with paternal particulate antigens. Effects on pregnancy. J Reprod Immunol 22:173–183 56. Gentile T, Llambias P, Dokmetjian J, Margni RA (1998) Effect of pregnancy and placental factors on the quality of humoral immune response. Immunol Lett 62:151–157 57. Margni RA, Malan Borel I (1999) Role of asymmetric antibodies in fetal maintenance. Curr Trends Immunol 2:53–163 58. Canellada A, Färber A, Zenclussen AC, Gentile T, Dokmetjian J, Keil A, Blois S, Miranda S, Berod L, Gutiérrez G, Markert UR, Margni RA (2002) Interleukin regulation of asymmetric antibody synthesized by isolated placental B cells. Am J Reprod Immunol 48(4):275–282 59. Gutierrez G,Malan Borel I, Margni RA (2001) The placental regulatory factor involved in the asymmetric IgG antibody synthesis responds to IL-6 features. J Reprod Immunol 49:21–32 60. Margni RA, Zenclussen AC (2001) During pregnancy, in the context of a Th2-type cytokine profile, serum IL-6 levels might condition the quality of the synthesized antibodies. Am J Reprod Immunol 46(3): 181–187 61. Zenclussen AC, Kortebani G, Mazzolli A, Margni R, Malan Borel I (2000) Interleukin-6 and soluble interleukin-6 receptor serum levels in recurrent spontaneous abortion women immunized with paternal white cells. Am J Reprod Immunol 44(1):22–29 62. Piccinni MP, Beloni L, Livi C, Maggi E, Scarselli G, Romagnani S (1998) Defective production of both, leukemia inhibitory factor and type 2 T-helper cytokines by decidual T cells in unexplained recurrent abortions. Nat Med 4:1020–1024 63. Lin H, Mossmann TR, Guilbert L, Tuntipopipat S, Wegmann TG (1993) Synthesis of T helper 2-type cytokines at the feto–maternal interface. J Immunol 151:4562–4573

414

Zenclussen et al.

64. Raghupathy R, Makhseed M, Azizieh F, Hassan N, Al-Azemi M, Al-Shamali E (1999) Maternal Th1- and Th2-type reactivity to placental antigens in normal human pregnancy and unexplained recurrent spontaneous abortions. Cell Immunol 196:122–130 65. Saito S (2001) Cytokine network at the feto– maternal interface. J Reprod Immunol 47:87–103 66. Svensson L, Arvola M, Sallstrom MA, Holmdahl R, Mattsson R (2001) The Th2 cytokines IL-4 and IL-10 are not crucial for the completion of allogeneic pregnancy in mice. J Reprod Immunol 51:3–7 67. Fallon PG, Jolin HE, Smith P, Emson CL, Townsend MJ, Fallon R, Smith P, McKenzie AN (2002) IL-4 induces characteristic Th2 even in the combined absence of IL-5, IL-9 and IL-13. Immunity 17:7–17 68. Dealtry GB, O’Farrell MK, Fernandez N (2000) The Th2 cytokine environment of the placenta. Int Arch Allergy Immunol 123(2):107–119 69. White CA, Johansson M, Roberts CT, Ramsay AJ, Robertson SA (2003) Effect of interleukin-10 null mutation on maternal immune response and reproductive outcome in mice. Biol Reprod 70(1):123–131 70. Gorivodsky M, Torchinsky A, Zemliak I, Savion S, Fein A, Toder V (1999) TGF beta 2 mRNA expression and pregnancy failure in mice. Am J Reprod Immunol 42(2):124–133 71. Dünker N, Krieglstein K (2002) Tgfbeta2−/− Tgfbeta3−/− double knockout mice display severe midline fusion defects and early embryonic lethality. Anat Embryol (Berl) 206(1–2):73–83 72. Kruse A, Merchant MJ, Hallmann R, Butcher EC (1999) Evidence of specialized leukocyte-vascular homing interactions at the maternal/fetal interface. Eur J Immunol 29(4):1116–1126 73. Zambon Bertoja A, Zenclussen ML, Wollenberg I, Paeschke S, Sollwedel K, Gerlof K, Woiciechosky C, Volk HD, Zenclussen AC (2005) Upregulation of Bcl-2 at the fetal– maternal interface from mice undergoing abortion. Scand J Immunol 61:492–502 74. Tangri S, Raghupathy R (1993) Expression of cytokines in placentas of mice undergoing immunologically mediated spontaneous fetal resorptions. Biol Reprod 49(4):850–856 75. Zenclussen AC, Gerlof K, Zenclussen ML, Sollwedel A, Zambon Bertoja A, Ritter T, Kotsch K, Leber J, Volk HD (2005) Abnormal T cell reactivity against paternal antigens in spontaneous abortion: adoptive transfer of pregnancy-induced CD4+CD25+

76.

77.

78.

79. 80.

81.

82.

83.

84.

85.

T regulatory cells prevents fetal rejection in a murine abortion model. Am J Pathol 166:811–822 Zenclussen AC, Fest S, Joachim R, Klapp BF, Arck PC (2004) Introducing a mouse model for pre-eclampsia: adoptive transfer of activated Th1 cells leads to pre-eclampsia-like symptoms exclusively in pregnant mice. Eur J Immunol 34(2):377–387 Harrington LE, Hatton RD, Mangan PR, Turner H, Murphy TL, Murphy KM, Weaver CT (2005) Interleukin 17-producing CD4 + effector T cells develop in a lineage distinct from the T helper type 1 and 2 lineages. Nat Immun 6:1123–1132 Park H, Li Z, Yang XO, Chang SH, Nurieva R, Wang YH, Wang Y, Hood L, Zhu Z, Tian Q, Dong C (2005) A distinct lineage of CD4 T cells regulates tissue inflammation by producing interleukin 17. Nat Immun 6:1133–1141 Moller G (1988) Do suppressor T cells exist? Scand J Immunol 27:247–250 Arruvito L, Billordo A, Capucchio M, Prada ME, Fainboim L (2009) IL-6 trans-signaling and the frequency of CD4+FOXP3+ cells in women with reproductive failure. J Reprod Immunol 82(2):158–165 Albanesi C, Cavani A, Girolomoni G (1999) IL-17 is produced by nickelspecific T lymphocytes and regulates ICAM-1 expression and chemokine production in human keratinocytes: synergistic or antagonist effects with IFN-gamma and TNF-alpha. J Immunol 162:494–502 Loong CC, Hsieh HG, Lui WY, Chen A, Lin CY (2002) Evidence for the early involvement of interleukin 17 in human and experimental renal allograft rejection. J Pathol 197:322–332 Vanaudenaerde BM, Dupont LJ, Wuyts WA Verbeken EK, Meyts I, Bullens DM, Dilissen E, Luyts L, Van Raemdonck DE, Verleden GM (2006) The role of interleukin-17 during acute rejection after lung transplantation. Eur Respir J 27:779–787 Yoshida S, Haque A, Mizobuchi T, Iwata T, Chiyo M, Webb TJ, Baldridge LA, Heidler KM, Cummings OW, Fujisawa T, Blum JS, Brand DD, Wilkes DS (2006) Anti-type V collagen lymphocytes that express IL-17 and IL-23 induce rejection pathology in fresh and well-healed lung transplants. Am J Transplant 6:724–735 Del Prete G, De Carli M, Almerigogna F, Giudizi MG, Biagiotti R, Romagnani S (1993) Human IL-10 is produced by both type 1 helper (Th1) and type 2 helper (Th2) T cell clones and inhibits their antigen-specific pro-

Pregnancy: Tolerance and Suppression of Immune Responses

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

liferation and cytokine production. J Immunol 150:353–360 Sakaguchi S, Sakaguchi N, Asano M, Itoh M, Toda M (1995) Immunological self-tolerance maintained by activated T-cells expressing IL-2 receptor alpha-chains (CD25). Breakdown of a single mechanism of self-tolerance causes various auto-immune diseases. J Immunol 155:1151–1164 Kingsley CL, Karim M, Bushell AR, Wood K (2002) CD25+CD4+ regulatory T cells prevent graft rejection: CTLA4- and IL-10-dependent immunoregulation of alloresponses. J Immunol 168:1080–1086 Hori S, Nomura T, Sakaguchi S (2003) Control of regulatory T cell development by the transcription factor Foxp3. Science 299:1057–1061 Fontenot JD, Rudensky AY (2005) A well adapted regulatory contrivance: regulatory T cell development and the forkhead family transcription factor Foxp3. Nat Immunol 6:331–337 Fontenot JD, Gavin MA, Rudensky AY (2003) Foxp3 programs the development and function of CD4+CD25+ regulatory T cells. Nat Immunol 4:330–336 Sakaguchi S (2004) Naturally arising CD4+ regulatory t cells for immunologic self-tolerance and negative control of immune responses. Annu Rev Immunol 22:531–562 Apostolou I, Sarukhan A, Klein L, von Boehmer H (2002) Origin of regulatory T cells with known specificity for antigen. Nat Immunol 3:756–763 Somerset DA, Zheng Y, Kilby MD, Sansom DM, Drayson MT (2004) Normal human pregnancy is associated with an elevation in the immune suppressive CD25+ CD4+ regulatory T-cell subset. Immunology 112:38–43 Aluvihare V, Kallikourdis M, Betz A (2004) Regulatory T cells mediate maternal tolerance to the fetus. Nat Immunol 3:266–271 Schumacher A, Wafula PO, Bertoja AZ, Sollwedel A, Thuere C, Wollenberg I, Yagita H, Volk HD, Zenclussen AC (2007) Mechanisms of action of regulatory T cells specific for paternal antigens during pregnancy. Obstet Gynecol 110:1137–1145 Bulmer JN, Johnson PM (1986) The T-lymphocyte population in first trimester human decidua does not express the interleukin-2 receptor. Immunology 58:685–687 Athanassakis I, Iconomidou B (1996) Cytokine production in the serum and spleen of mice from day 6 to 14 of gestation: cytokines/placenta/spleen/serum. Dev Immunol 4:247–255

415

98. Zhu XY, Zhou YH, Wang MY, Jin LP, Yuan MM, Li DJ (2005) Blockade of CD86 signaling facilitates a Th2 bias at the maternal–fetal interface and expands peripheral CD4+CD25+ regulatory T cells to rescue abortion-prone fetuses. Biol Reprod 72:338–345 99. Schumacher A, Brachwitz N, Sohr S, Engeland K, Langwisch S, Dolaptchieva M, Alexander T, Taran A, Malfertheiner SF, Costa SD, Zimmermann G, Nitschke C, Volk HD, Alexander H, Gunzer M, Zenclussen AC (2009) Human chorionic gonadotropin attracts regulatory T cells into the fetal– maternal interface during early human pregnancy. J Immunol 182(9):5488–5497 100. Groux H, O’Garra A, Bigler M, Rouleau M, Antonenko S, de Vries JE, Roncarolo MG (1997) A CD4+ T-cell subset inhibits antigen-specific T-cell responses and prevents colitis. Nature 389:737–742 101. Weiner HL (2001) Induction and mechanism of action of transforming growth factor-beta-secreting Th3 regulatory cells. Immunol Rev 182:207–214 102. Wafula PO, Teles A, Schumacher A, Pohl K, Yagita H, Volk HD, Zenclussen AC (2009) PD-1 but not CTLA-4 blockage abrogates the protective effect of regulatory T cells in a pregnancy murine model. Am J Reprod Immunol 62(5):283–292 103. Zenclussen AC, Gerlof K, Zenclussen ML, Ritschel S, Zambon Bertoja A, Fest S, Hontsu S, Ueha S, Matsushima K, Leber J, Volk HD (2006) Regulatory T cells induce a privileged tolerance microenvironment at the fetal– maternal interface. Eur J Immunol 36:82–94 104. Bhatt H, Brunet LJ, Stewart CL (1991) Uterine expression of leukemia inhibitory factor coincides with the onset of blastocyst implantation. Proc Natl Acad Sci USA 88(24):11408–11412 105. Stewart CL, Kaspar P, Brunet LJ, Bhatt H, Gadi I, Kontgen F, Abbondanzo SJ (1992) Blastocyst implantation depends on maternal expression of leukaemia inhibitory factor. Nature 359:76–79 106. Cheng JG, Rodriguez CI, Stewart CL (2002) Control of uterine receptivity and embryo implantation by steroid hormone regulation of LIF production and LIF receptor activity: Towards a molecular understanding of “the window of implantation”. Rev Endocr Metab Disord 3:119–126 107. Mikołajczyk M, Skrzypczak J, Szymanowski K, Wirstlein P (2003) The assessment of LIF in uterine flushing: a possible new diagnostic tool in states of impaired fertility. Reprod Biol 3(3):259–270

416

Zenclussen et al.

108. Hu W, Feng Z, Teresky AK, Levine AJ (2007) p53 regulates maternal reproduction through LIF. Nature 450(7170):721–724 109. Robertson SA (2005) Seminal plasma and male factor signaling in the female reproductive tract. Cell Tissue Res 322(1):43–52 110. Zhao H, Wong RJ, Kalish FS, Nayak NR, Stevenson DK (2009) Effect of heme oxygenase-1 deficiency on placental development. Placenta 30(10):861–868 111. Wise CD, Drabkin DL (1964) Degradation of haemoglobin and hemin to biliverdin by a new cell-free system obtained from the hemophagous organ of dog placenta. Fed Proc 23:323 112. Tenhunen R, Marver HS, Schmid R (1968) The enzymatic conversion of heme to bilirubin by microsomal heme oxygenase. Proc Natl Acad Sci USA 61(2):748–755 113. Tenhunen R, Marver HS, Schmid R (1969) Microsomal heme oxygenase – characterization of the enzyme. J Biol Chem 244(23): 6388–6394 114. Tenhunen R, Marver H, Pimstone NR, Trager WF, Cooper DY, Schmid R (1972) Enzymatic degradation of heme. Oxygenative cleavage requiring cytochrome P-450. Biochemistry 11(9):1716–1720 115. Ryter SW, Otterbein LE, Morse D, Choi AM (2002) Heme oxygenase/carbon monoxide signaling pathways: regulation and functional significance. Mol Cell Biochem 234–235 (1–2):249–263 116. Montellano PR (2000) The mechanism of heme oxygenase. Curr Opin Chem Biol 4(2):221–227 117. Otterbein LE, Choi AM (2000) Heme oxygenase: colors of defense against cellular stress. Am J Physiol Lung Cell Mol Physiol 279(6):L1029–L1037 118. Morse D, Choi AM (2002) Heme oxygenase-1: the “Emerging Molecule” has arrived. Am J Respir Cell Mol Biol 27(1):8–16 119. Maines MD (1997) The heme oxygenase system: a regulator of second messenger gases. Annu Rev Pharmacol Toxico 37:517–554 120. Soares MP, Lin Y, Anrather J, Csizmadia E, Takigami K, Sato K, Grey ST, Colvin RB, Choi AM, Poss KD, Bach FH (1998) Expression of heme oxygenase-1 can determine cardiac xenograft survival. Nat Med 4(9):1073–1077 121. Coito AJ, Buelow R, Shen XD, Amersi F, Moore C, Volk HD, Busuttil RW, KupiecWeglinski JW (2002) Heme oxygenase-1 gene transfer inhibits inducible nitric oxide

122.

123.

124.

125.

126.

127.

128.

129.

130.

synthase expression and protects genetically fat zucker rat livers from ischemia-reperfusion injury. Transplantation 74(1):96–102 Tullius SG, Nieminen-Kelha M, Buelow R, Reutzel-Selke A, Martins PN, Pratschke J, Bachmann U, Lehmann M, Southard D, Iyer S, Schmidbauer G, Sawitzki B, Reinke P, Neuhaus P, Volk HD (2002) Inhibition of ischemia/reperfusion injury and chronic graft deterioration by a single-donor treatment with cobalt-protoporphyrin for the induction of heme oxygenase-1. Transplantation 74(5):591–598 Braudeau C, Bouchet D, Tesson L, Iyer S, Rémy S, Buelow R, Anegon I, Chauveau C (2004) Induction of long-term cardiac allograft survival by heme oxygenase-1 gene transfer. Gene Ther 11(8):701–710 Juan SH, Lee TS, Tseng KW, Liou JY, Shyue SK, Wu KK, Chau LY (2001) Adenovirusmediated heme oxygenase-1 gene transfer inhibits the development of atherosclerosis in apolipoprotein E-deficient mice. Circulation 104(13):519–525 Fujii H, Takahashi T, Nakahira K, Uehara K, Shimizu H, Matsumi M, Morita K, Hirakawa M, Akaqi R, Sassa S (2003) Protective role of heme oxygenase-1 in the intestinal tissue injury in an experimental model of sepsis. Crit Care Med 31(3):893–902 Chora AA, Fontoura P, Cunha A, Pais TF, Cardoso S, Ho PP, Lee LY, Sobel RA, Steinman L, Soares MP (2007) Heme oxygenase-1 and carbon monoxide suppress autoimmune neuroinflammation. J Clin Invest 117(2):438–447 Pamplona A, Ferreira A, Balla J, Jeney V, Balla G, Epiphano S, Chora A, Rodrigues CD, Gregoire IP, Cunha-Rodrigues M, Portugal S, Soares MP, Mota MM (2007) Heme oxygenase-1 and carbon monoxide suppress the pathogenesis of experimental cerebral malaria. Nat Med 13(6):703–710 Christiansen OB, Nielsen HS, Kolte AM (2006) Inflammation and miscarriage. Semin Fetal Neonatal Med 11:302–308 Ryter SW, Alam J, Choi AM (2006) Heme oxygenase-1/carbon monoxide: from basic science to therapeutic applications. Physiol Rev 86(2):583–650 Zenclussen ML, Anegon I, Bertoja AZ, Chauveau C, Vogt K, Gerlof K, Sollwedel A, Volk HD, Ritter T, Zenclussen AC (2006) Over-expression of heme oxygenase-1 by adenoviral gene transfer improves pregnancy outcome in a murine model of abortion. J Reprod Immunol 69(1):35–52

Pregnancy: Tolerance and Suppression of Immune Responses 131. Sollwedel A, Bertoja AZ, Zenclussen ML, Gerlof K, Lisewski U, Wafula P, Sawitzki B, Woiciechowsky C, Volk HD, Zenclussen AC (2005) Protection from abortion by heme oxygenase-1 up-regulation is associated with increased levels of bag-1 and neuropilin-1 at the fetal-maternal interface. J Immunol 175(8):4875–4885 132. Ihara N, Akagi R, Ejiri K, Kudo T, Furuyama K, Fujita H (1998) Developmental changes of gene expression in heme metabolic enzymes in rat placenta. FEBS Lett 439: 163–167 133. Barber A, Robson SC, Myatt L, Bulmer J, Lyall F (2001) Heme oxygenase expression in human placenta and placental bed: reduced expression of placenta endothelial HO-2 in pre-eclampsia and fetal growth restriction. FASEB J 15:1158–1168 134. Zenclussen AC, Lim E, Knoeller S, Knackstedt M, Hertwig K, Hagen E, Klapp BF, Arck PC (2003) Heme oxygenases in pregnancy II: HO-2 is downregulated in human pathologic pregnancies. Am J Reprod Immunol. 50: 66–76 135. Zenclussen AC, Sollwedel A, Zambon Bertoja A, Gerlof K, Zenclussen ML, Woiciechowsky C, Volk HD (2005) Heme oxygenase as a therapeutic target in immunological pregnancy complications. Int Immunopharmacol 5(1):41–51 136. Cross JC (2005) How to make a placenta: mechanisms of trophoblast cell differentiation in mice- a review. Placenta 26(Suppl A, Trophoblast Research 19):S3–S9 137. Faria TN, Soares MJ (1991) Trophoblast cell differentiation: establishment, characterization, and modulation of rat trophoblast cell line expressing members of the placental prolactin family. Endocrinology 129(6): 2895–2906 138. Sahgal N, Canham LN, Konno T, Wolfe MW, Soares MJ (2005) Modulation of trophoblast stem cell and giant cell phenotypes: analyses using the Rcho-1 cell model. Differentiation 73:452–462 139. Poss K, Tonegawa S (1997) Reduced stress defense in heme oxygenase-1 deficient cells. Proc Natl Acad Sci USA 94:10925–10930 140. Yet SF, Perrella MA, Layne MD, Hsieh CM, Maemura K, Kobzik L, Wiesel P, Christou H, Kourembanas S, Lee ME (1999) Hypoxia induces severe right ventricular dilatation

141.

142.

143.

144.

145.

146.

147.

148.

149.

150.

417

and infarction in heme oxygenase-1 null mice. J Clin Invest 103(8):R23–R29 Espey LL, Bellinger AS, Healy JA (2004) Ovulation: An inflammatory cascade of gene expression, In: The ovary, PCK Leung and EY Adashi, Eds. Academic Press, New York, pp. 145–165 Jablonka-Shariff A, Olson LM (1998) The role of nitric oxide in oocyte meiotic maturation and ovulation: meiotic abnormalities of endothelial nitric oxide synthase knockout mouse oocytes. Endocrinology 139: 2944–2954 Drazen DL, Klein SL, Burnett AL, Wallach EE, Crone JK, Huang PL, Nelson RJ (1999) Reproductive function in female mice lacking the gene for endothelial nitric oxide synthase. Nitric Oxide 3:366–374 Hefler LA, Gregg AR (2002) Inducible and endothelial nitric oxide synthase: genetic background affects ovulation in mice. Fertil Steril 77:147–151 Yang J, Ajonuma LC, Rowlands DK, Tsang LL, Ho LS, Lam SY, Chen WY, Zhou CX, Chung YW, Cho CY, Tse JY, James AE, Chan HC (2005) The role of inducible nitric oxide synthase in gamete interaction and fertilization: a comparative study on knockout mice of three NOS isoforms. Cell Biol Int 29:785–791 McGarry HF, Plant LD, Taylor MJ (2005) Diethylcarbamazine activity against Brugia malayi microfilariae is dependent on inducible nitric-oxide synthase and the cyclooxygenase pathway. Filaria J 4:4 Ejima K, Perrella MA (2004) Alteration in heme oxygenase-1 and nitric oxide synthase-2 gene expression during endotoxemia in cyclooxygenase-2 deficient mice. Antioxid Redox Signal 6(5):850–857 Venturini CM, Isakson P, Needleman P (1998) Non-steroidal anti-inflammatory drug-induced renal failure: a brief review of the role of cyclo-oxygenase isoforms. Curr Opin Nephrol Hypertens 7(1):79–82 Matsumoto H, Ma W, Smalley W, Trzaskos J, Breyer RM, Dey SK (2001) Diversification of cyclooxygense-2-derived prostaglandins in ovulation and implantation. Biol Reprod 64:1557–1565 Poss KD, Thomas MJ, Ebralidze AK, O’Dell TJ, Tonegawa S (1995) Hippocampal longterm potentiation is normal in heme oxygenase-2 mutant mice. Neuron 15:867–873

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Chapter 26 Organ Transplantation: Modulation of T-Cell Activation Pathways Initiated by Cell Surface Receptors to Suppress Graft Rejection Kathleen Weatherly and Michel Y. Braun Abstract T-cell activation depends upon two types of signals: a T-cell-receptor-mediated antigen-specific signal and several non-antigen-specific ones provided by the engagement of costimulatory and/or inhibitory T-cell surface molecules. In clinical transplantation, T-cell costimulatory/inhibitory molecules are involved in determining cytokine production, vascular endothelial cell damage, and induction of transplant rejection. Several of the latest new immunotherapeutic strategies being currently developed to control graft rejection aim at inhibiting alloreactive T-cell function by regulating activating and costimulatory/inhibitory signals to T cells. This article describes the recent development and potential application of these therapies in experimental and pre-clinical transplantation. Key words: T-cell receptor, Costimulation, Transplantation, Anergy, Inhibitory receptors

1. Introduction T cells are key players in the rejection of transplanted organs as they tightly modulate the inflammatory processes that lead to graft rejection. Therefore, understanding and controlling the different molecular pathways that regulate T-cell activation has been for decades the main subject of investigation for transplantation immunologists. T-cell activation is a complex process that is orchestrated by the delivery of numerous positive and negative signals to T cells. Attempts to manipulate these signals in the context of transplantation have created great expectations but have yet to deliver significant results for clinical benefits.

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_26, © Springer Science+Business Media, LLC 2011

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The present review presents the main immunotherapeutic approaches currently developed in transplantation to control T-cell activation and promote immune tolerance toward allografted organs. The balance between positive and negative signals delivered by antigen-presenting cells (APC) to T cells determine the outcome of an alloimmune response. Activation of alloreactive T lymphocytes is initiated by the engagement of cell surface CD3/ TcR complexes by recipient’s allogeneic MHC/peptide complexes. The CD3/TcR complex comprises two subunits a and b, responsible for binding allo-MHC/peptide ligands, and a signalization subunit formed by the chains (g, d, e, and z) of the CD3 complex. It is generally presumed that, as in non-alloimmune responses, TcR engagement by allogeneic stimuli induces the phosphorylation of tyrosine residues present in the ITAMs (immunoreceptor tyrosine-based activation motif) of CD3 chains by the Src protein kinases Lck and Fyn (Fig. 1). Phosphorylated tyrosines of the z chain then form a docking site for the protein ZAP-70 (zeta chain-associated protein kinase 70 kD). ZAP-70 then becomes a substrate for Lck and acquires its own tyrosine kinase activity which leads to the phosphorylation of adaptor protein LAT (linker of activation of T cells) that becomes an anchor site for other adaptors or enzymes (Fig. 1). From the LAT protein, three main signaling pathways induced by TcR/CD3 crosslinking lead to the activation of the transcription factors NF-AT, NF-kB, and AP-1 whose activity concurs to the optimal transcription of genes encoding proteins essential for the activation of T cells (Fig. 1). The idea of modulating TcR-mediated signaling to regulate T-cell responses responsible for graft rejection came soon after the observation that anti-CD3/TcR antibody treatment led to a strong suppressive activity (1, 2). The strong in vivo and in vitro immunosuppressive potency of the antibody was evidenced by its capacity to induce activation-induced cell death in T cells and prompted its use in clinical transplantation (3–5). However, it became rapidly clear that this therapy also induced serious nonspecific side effects through uncontrolled activation of T cells. The injection of unaltered anti-CD3 antibody was shown to activate polyclonally T cells with the occurrence of a severe syndrome during which recipients experience the side effects of a consequential production of inflammatory cytokines such as TNF-a (6–9). To avoid these deleterious side effects, anti-CD3 antibodies were engineered to prevent their capacity to bind Fc receptors and, therefore, to reduce their ability to stimulate T cells efficiently by TcR cross-linking (10, 11). This treatment is believed to induce T-cell immune deviation and favors the appearance of alloantigen-specific Th2 cells in vivo (12, 13). It has also been shown to promote the emergence of T-regulatory cells (14).

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Fig. 1. Signaling pathways of T-cell activation.

In vitro experiments have shown that engagement of TcR/ CD3 alone by anti-CD3 antibodies can induce a long-term hyporesponsiveness in T cells, called clonal anergy (15). Anergic T cells proliferate poorly and have low IL-2 production upon antigenic challenge. Anergy depends on calcium uptake and can be prevented by Cyclosporine A, indicating that it is induced in conditions in which signaling by calcium and the transcription factor NF-AT are increased over other signals (16). Anergic T cells

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have a defective activation of the GTPase Ras and a subsequent decreased activation of the MAPKinase pathway, as well as a diminished transactivation of the transcription factor AP-1 (17). Whether in vivo treatment with anti-CD3 antibodies is responsible for inducing anergy in T cells is difficult to assess. One has to presume that it could be the case since T cells from anti-CD3 antibody-treated animals show low proliferative capacity as well as lack of IL-2 secretion upon antigenic challenge. However, the observation that T cells isolated after anti-CD3 antibody treatment can produce Th2 type as well as anti-inflammatory cytokines favors the possibility that in vivo “anergy” is more likely a consequence of immune deviation than specific clonal unresponsiveness (12, 13). Experiments performed in animal transplantation models show indeed that a short treatment with anti-CD3e antibodies promotes the development of Th2-type alloreactive T cells in the recipient’s secondary lymphoid organs and extends allograft survival (12). Though TcR-mediated antigen recognition gives the specificity to T-cell responses, the participation of other receptors is required for the effective transduction of activating signals. During TcR/ MHC/antigen peptide interaction, co-receptors CD4 and CD8 binding to non-polymorphic regions of the class I and II MHC molecules, respectively, ensure the specific recruitment of Lck to the CD3/ZAP-70 complex (18, 19). Interestingly, short-term treatment with non-depleting anti-co-receptor CD4 antibodies enables long-term transplantation tolerance in rodents (20). Co-receptor blockade is presumed to prevent CD4 binding to class II MHC molecules, resulting in a lack of recruitment of Lck to the TcR/CD3/ZAP-70 complex. Interestingly, CD4 blockade was shown to allow TGF-b-dependent conversion of alloantigen-specific naive T cells to T-regulatory cells (21). The use of mouse anti-human CD4 antibodies that have been humanized and engineered in its Fc portion to become non-lytic was shown to induce specific immune unresponsiveness in baboons, while responses against third party antigens were left unaltered (22). These observations preclude that tolerization by co-receptor blockade could be translated in the clinics. It is now also well established that interactions between so-called costimulatory receptors present at the surface of T cells and their ligands expressed by APC facilitate the generation of T-cell responses by lowering the threshold of T-cell activation. Among costimulatory proteins, CD28 has been the most studied molecule. It is constitutively expressed at the surface of T cells and binds to CD80 and CD86 molecules expressed on APCs (23, 24). Its cytoplasmic tail possesses an ITAM whose phosphorylation, by yet unidentified tyrosine kinases, allows the recruitment of proteins such as Grb-2 (growth factor-bound protein 2), Itk (IL-2-inducible T-cell kinase), and PI3K (phosphatidylinositol

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3-hydrokinase) (25–27). The PI3K signaling pathway is involved in numerous functions in T lymphocytes, including cell proliferation and cytokine secretion (26). The pathway leads to the phosphorylation and activation of protein kinase B (PKB-Atk) which is essential for the production of IL-2 and IFN-g (Interferon-g). PI3K signaling is also important for T-cell survival as it activates the synthesis of anti-apoptotic factor Bcl-xL (28). Grb-2 and Itk bind LAT and then initiate signaling pathways that lead to the activation of AP-1 and NF-AT, respectively (29, 30). The critical role played by CD28 in the activation of alloreactive T cells came from studies where selective inhibition of CD28/B7 interactions using a non-activating monoclonal antibody to CD28 was shown to induce transplantation tolerance in a rat model of acute kidney graft rejection (31). A short-term treatment abrogated both acute and chronic rejection. Tolerant recipients presented few alloantibodies against donor MHC class II molecules, whereas untreated graft rejecting controls developed anti-MHC class I and II alloantibodies. T-cell activation induces the expression of a series of cell membrane proteins that play a major role in down-regulating T-cell activity. Among these molecules, CTLA-4 (CD152) is a member of the CD28 family (32). It inhibits T-cell activation by reducing IL-2 synthesis and blocks T-cell proliferation in the G0 phase (33). The inhibitory role of CTLA-4 was definitively unveiled in studies that analyzed mice with deleted CTLA-4 gene (34). These animals died between 3 and 4 weeks of age from an acute autoimmune lymphoproliferative disease. CTLA-4 does not appear to play a major role in thymic selection of T cells, since CTLA-4-deficient animals do not present abnormalities in thymocyte development. As opposed to CD28 which is expressed at the surface of most T cells, CTLA-4 is only expressed by activated T cells. The number of CTLA-4 molecules expressed at the cell surface is tightly regulated. Part of this regulation involves the AP-2 complex (adaptor protein complex 2) (35). This complex binds the YKVM motif of CTLA-4 receptor and allows its internalization from the cell surface to intracellular compartments such as endosomic vesicles and lysosome, where it is rapidly degraded. AP-2/CTLA-4 binding is prevented by phosphorylation of tyrosine Y201 present in CTLA-4 YKVM motif. During T-cell activation, the kinases Lck and Fyn phosphorylate Y201, preventing AP-2 binding and internalization of CTLA-4. Several hypotheses exist about how CTLA-4 inhibits T-cell activity. First, CTLA-4 could compete with CD28 for binding to B7 molecules and prevent T-cell costimulation. These two molecules bind the same B7 ligands, CD80 and CD86 (32). However, CTLA-4 binds B7 ligands with a tenfold higher affinity than does CD28. Thus, CTLA-4 could sequester B7 ligands away from CD28. This concept has progressed to the point where inhibiting CD28-mediated

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T-cell costimulation by soluble CTLA-4-Ig fusion proteins is currently tested to suppress the rejection of allografted organs (36). In fact, the capacity of CTLA-4-Ig to prevent transplant rejection has been demonstrated in various murine and rodent models. In primates, the use of a CTLA-4-Ig protein engineered to increase its affinity to B7 ligands demonstrated a good efficacy to inhibit allograft rejection. Current clinical trials are assessing the treatment in transplanted patients. Experimental evidence supports the idea that CTLA-4 might inhibit directly the TcR signaling cascade. Bluestone and colleagues have documented the capacity of CTLA-4 to associate with the z chain of TcR/CD3 complex, allowing the recruitment of the phosphatase SHP-2 (SH2-domain containing tyrosine phosphatase) and the subsequent dephosphorylation of CD3 z (37). A third mechanism of action involves the inhibition of the CD28 signaling pathway. CTLA-4 activation of the phosphatase PP2A (phosphatase protein 2A) directly inhibits the phosphorylation of the protein Atk, an essential kinase to CD28 signaling (38). CTLA-4 can also interfere with the expression and composition of lipid rafts at the surface of T cells (39). Lipid rafts are microdomains rich in glycolipids that participate in the formation of the immunological synapse. Whatever is the mechanism used by CTLA-4 to inhibit T-cell activity, it results in a decrease in T-cell cytokine production and the inhibition of the transcription factors NF-kB, NF-AT, and AP-1. Tolerance induction may therefore be facilitated by inhibiting selectively the B7/CD28 pathway without blocking of B7/CTLA-4. Recent evidence has revealed the important role played by the protein PD-1 (programmed cell death receptor 1) in the control of T-cell activity (40). PD-1 is a member of the CD28 family of proteins. It was initially identified as a molecule expressed by apoptotic T cells. Recent studies, however, have shown that its expression is associated with the control of cell activation. PD-1 expression is induced on activated T and B lymphocytes as well as at the surface of myeloid cells. It contains an inhibitory ITIM motif as well as an immunoreceptor tyrosine-based switch motif (ITSM) in its cytoplasmic tail and, upon ligand engagement, both motives are phosphorylated and can bind to two phosphatases, SH2-domain containing tyrosine phosphatase 1 (SHP-1) and SHP-2 (41). Recent work has shown that PD-1 suppressing function depends primarily on the tyrosine present in its ITSM (42). These studies also suggest that the role of PD-1 is to recruit SHP-1 and SHP-2 to the TcR signaling complex. Proximity of PD-1 to the TcR seems to be important for inhibition by PD-1. PD-1 ligation inhibits antigen receptor signaling only in cis, indicating that PD-1 ligation must occur close to the site of TcR engagement. The inhibitory activity of SHPs is expressed by the dephosphorylation of TcR proximal signaling

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proteins, ZAP-70 and CD3 z, which results in the inhibition of the different signaling triggered by TcR engagement. Like CTLA4, PD-1 can also inhibit CD28 signaling (43). However, this inhibitory activity targets early events in the CD28 signaling cascade. PD-1 suppresses the activity of the protein kinase Akt by inhibiting PI3K. PD-1 engagement limits the production of IFN-g and IL-2 by T lymphocytes as well as T-cell proliferation. Mice without a functional PD-1 develop spontaneous chronic autoimmune diseases, such as cardiomyopathy, arthritis, and glomerulonephritis (40, 44). These animals are also more sensitive to the induction of experimental autoimmune encephalomyelitis, and in PD-1-deficient NOD mice, the onset of diabetes develops earlier (45, 46). Recent evidences show that blockade of the interaction between PD-1 and its ligand influences the motility of PD-1positive tolerant islet antigen-specific T cells in draining pancreatic lymph nodes, resulting in lower T-cell motility, enhanced T cell–DC contacts, and causing autoimmune diabetes (47). PD-1 can react with two ligands, PD-L1 or PD-L2, but its regulatory activity appears to result mainly from its interactions with PD-L1 (48). PD-L1 is expressed on a set of cells distinct from that of any other members of the B7 family. PD-L1 expression is increased on activated APC, such as dendritic cells, monocytes, and B cells. It is also expressed by a wide variety of cell types and tissues after exposure to inflammatory cytokines, such as type I and type II interferons. PD-L2 is expressed exclusively by dendritic cells and monocytes. Both PD-L1 and PD-L2 inhibit in vitro the proliferation and cytokine production by CD4 and CD8 T cells. Recently, PD-L1 suppressive activity on T cells was shown to involve its interaction with CD80, adding another dimension to the immunoregulatory functions of the B7 protein family (49). The role played by PD-1/PD-L1 pathway in the regulation of alloreactive responses has been examined in animal models of transplantation (50, 51). The rejection of mouse heart allografts transplanted across fully MHC-mismatched barrier was delayed in PD-1-deficient recipients, whereas less-immunogenic MHC class I- or class II-mismatched grafts enjoyed long-term survival in the same recipients. In the same way, transplantation of PD-L1deficient cardiac grafts or blocking PD-L1 with specific antibodies accelerated rejection. Taken together, these results suggested that T-cell responses to graft alloantigens were enhanced when T cells did not receive negative signals through PD-1/PD-L1 interactions. However, though the rejection of PD-L1-deficient hearts could be delayed, rejection of PD-L2-deficient cardiac allograft did not differ from that of PD-L2-sufficient organs, demonstrating that PD-L2 played no or little role in the control of T cells in the context of graft rejection. These results preclude the use of soluble recombinant PD-L1-Ig fusion proteins as potential

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therapeutic tools for the suppression of T-cell alloreactivity. Injection of PD-L1-Ig did not prolong the survival of fully MHCmismatched mouse heart allografts. However, when immunosuppressive agents, such as Cyclosporine A or Rapamycin, were administrated in combination with PD-L1-Ig, substantial prolongation of graft survival was observed (50). Thus, these observations suggested that immunotherapies based on PD-1-mediated suppression of T-cell signaling could be used to lower the level of immunosuppressive drugs in transplanted patients. Interestingly, the observation that PD-L1-Ig treatment was more efficient for the suppression of allograft rejection in CD28-deficient recipients supported the possibility to combine treatments targeting PD-1mediated co-inhibition of T-cell activity with those aimed at blocking CD28-dependent costimulation of T cells (50). Another pathway controlling the T-cell activity, which interests transplantation immunologists, is the CD40/CD40L pathway (reviewed in ref. 52). CD40L is expressed at the surface of activated CD4+ T cells and has also been detected in platelets, B cells, CD8+ T cells, DCs, and ECs. A functional CD40–CD40L pathway is critical for effective T-cell activation, and ligation of CD40 on B cells induces proliferation and antibody production. Blockade of the CD40–CD40L pathway has been shown to promote allograft acceptance in several animal models (53, 54). These mechanisms include deletion of alloreactive T cells, T-cell anergy and the induction of T cells with regulatory activity (Tregs). Though blockade of CD40–CD40L interactions can prevent acute rejection of hearts and kidneys in rodents, they are not as efficient at inhibiting skin rejection in rodents or kidney rejection in primates (55). Poorer results after blockade of CD40–CD40L in primates might be explained by higher numbers of antiviral memory T cells cross-reactive with donor antigens which are more difficult to control since their activity is less dependent on CD40/CD40L interactions. These results prompted the use of CD40/CD40L blockade-based therapies in conjunction with other immunosuppressive agents. The observation that combination of blocking anti-CD40L antibodies with soluble CTLA-4-Ig induced longterm acceptance of kidney allografts in primates could justify testing CD40/CD40L-based immunotherapies in clinical trials (56). Non-depleting antibodies targeting the a-chain of the interleukin-2 receptor (IL-2Ra) are also used to suppress graft rejection (57). Interleukin 2 (IL-2), or T-cell growth factor (TCGF), is produced mainly by CD4 T lymphocytes, stimulates cell-mediated immune responses, controls growth and differentiation of B lymphocytes, and intensifies proliferation and activity of all cytotoxic cell clones. IL-2 is a growth factor in vitro and a mediator of selftolerance in vivo, and therefore interests transplantation immunologists. The role played by IL-2 in tolerance induction and maintenance was shown in IL-2-, or functional IL-2R-, deficient

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mice which develop signs of autoimmunity and lack T-regulatory cells (58, 59). Moreover, failure to induce long-term allograft survival in IL-2-deficient mice indicated that endogenous IL-2 was indeed required for the induction of transplantation tolerance (60). However, numerous evidences indicate that IL-2 is a TCGF and administration of anti-IL-2 receptor a chain (CD25) antibodies in the rat was shown to synergize with Cyclosporine A to induce tolerance to pancreatic islet allografts (61). Tolerance could be achieved presumably because activated T cells which express CD25 were specifically depleted by the antibody treatment. In the clinics, anti-CD25 antibodies are used to control graft rejection. These antibodies exhibit a reduced capacity to deplete T cells and do not seem to inhibit the activity of CD4 T-regulatory cells in patients allografted with heart or kidney. Interestingly, anti-CD25 antibody treatment appears to stimulate the expansion of a subpopulation of NK cells with regulatory activity. These NK cells could be responsible for the inhibitory effect observed on T-cell activity after anti-CD25 antibody treatment (62).

2. Conclusion This is an exciting time for transplantation immunologists as fundamental studies have delivered many therapeutic concepts that are currently translated in the clinics. However, first results from clinical trials seem to indicate that induction of transplantation tolerance will not be achieved in humans with intervention on one single T-cell activation pathway but will certainly require the combined suppression/stimulation of several activating/inhibitory pathways. Several unanswered fundamental questions also remain: (1) will immunotherapies developed for the prevention of acute rejection be efficient to inhibit T cells under permanent exposure to alloantigens expressed by long-term-surviving allografts; (2) how will modulation of T-cell activation applied to induce transplantation tolerance affect the development of other immune responses such as those developed against infection and cancer; (3) will there be a need to target directly T cells responsible for allograft rejection to render these therapies more efficient. There is currently good hope that these questions will be answered in the forthcoming years. References 1. Cosimi, A. B., Colvin, R. B., Burton, R. C., Rubin, R. H., Goldstein, G., Kung, P. C., et al. (1981) Use of monoclonal antibodies to T-cell subsets for immunologic monitoring

and treatment in recipients of renal allografts. N. Engl. J. Med. 305, 308–314. 2. Vigeral, P., Chkoff, N., Chatenoud, L., Campos, H., Lacombe, M., Droz, D., et al.

428

3.

4.

5.

6.

7.

8.

9.

10.

11.

12.

Weatherly and Braun (1986) Prophylactic use of OKT3 monoclonal antibody in cadaver kidney recipients. Utilization of OKT3 as the sole immunosuppressive agent. Transplantation. 41, 730–733. Wesselborg, S., Janssen, O., and Kabelitz, D. (1993) Induction of activation-driven death (apoptosis) in activated but not resting peripheral blood T cells. J. Immunol. 150, 4338–4345. Liu, Y., and Janeway, C. A. (1990) Interferon gamma plays a critical role in induced cell death of effector T cells: a possible third mechanism of self-tolerance. J. Exp. Med. 172, 1735–1739. Russell, J. H., Rush, B. J., Abrams, S. I., Wang, R. (1992) Sensitivity of T cells to antiCD3 stimulated suicide is independent of functional properties. Eur. J. Immunol. 22, 1655–1658. Abramowicz, D., Schandene, L., Goldman, M., Crusiaux, A., Vereerstraeten, P., De Pauw, L., et al. (1989) Release of tumor necrosis factor, interleukin-2, and gamma-interferon in serum after injection of OKT3 monoclonal antibody in kidney transplant recipients. Transplantation. 47, 606–608. Hirsch, R., Gress, R. E., Pluznik, D. H., Eckhaus, M. and Bluestone J. A. (1989) Effects of in vivo administration of anti-CD3 monoclonal antibody on T cell function in mice. II. In vivo activation of T cells. J. Immunol. 142, 737–743. Ferran, C., Sheehan, K., Dy, M., Schreiber, R., Merite, S., Landais, P., et al. (1990) Cytokine-related syndrome following injection of anti-CD3 monoclonal antibody: further evidence for transient in vivo T cell activation. Eur. J. Immunol. 20, 509–515. Alegre, M., Vandenabeele, P., Flamand, V., Moser, M., Leo, O., Abramowicz, D., et al. (1990) Hypothermia and hypoglycemia induced by anti-CD3 monoclonal antibody in mice: role of tumor necrosis factor. Eur. J. Immunol. 20, 707–710. Hirsch, R., Bluestone, J. A., De Nonno, L., , and Gress, R. E. (1990) Anti-CD3 F(ab )2 fragments are immunosuppressive in vivo without evoking either the strong humoral response and morbidity associated with whole mAb. Transplantation. 49, 1117–1123. Parlevliet, K. J., Ten Berge, I. J., Yong, S. L., Surachno, J., Wilmink, J. M., and Schellekens, P. T. (1994) In vivo effects of Iga and IgG2a anti-CD3 isotype switch variants. J. Clin. Invest. 93, 1117–1123. Plain, K. M., Chen, J., Merten, S., He, X. Y., Hall, B. M. (1999) Induction of specific

13.

14.

15.

16. 17.

18.

19.

20.

21.

22.

23.

tolerance to allografts in rats by therapy with non-mitogenic, non-depleting anti-CD3 monoclonal antibody: association with TH2 cytokines not anergy. Transplantation. 67, 605–613. Smith, J. A., Tang, Q., and Bluestone, J. A. (1998) Partial TcR signals delivered by FcRnon-binding anti-CD3 monoclonal antibodies differentially regulate individual Th subsets. J. Immunol. 160, 4841–4849. Belghith, M., Bluestone, J. A., Barriot, S., Megret, J., Bach, J. F., Chatenoud, L. (2003) TGF-beta-dependent mechanisms mediate restoration of self-tolerance induced by antibodies to CD3 in overt autoimmune diabetes. Nat. Med. 9, 1202–1208. Smith, J. A., Tso, J. Y., Clark, M. R., Cole, M.S., and Bluestone, J. A. (1997) Nonmitogenic anti-CD3 monoclonal antibodies deliver a partial T cell receptor signal and induce clonal anergy. J. Exp. Med. 185, 1413–1422. Schwartz, R. H. (2003) T cell anergy. Annu. Rev. Immunol. 21, 305–334. Zha, Y., Marks, R., Ho, A. W., Peterson, A. C., Janardhan, S., Brown, I., et al. (2006) T cell anergy is reversed by active Ras and by diacylglycerol kinase-a. Nat. Immunol. 7, 1166–1173. Veillette, A., Bookman, M. A., Horak, E. M., Bolen, J. B. (1988) The CD4 and CD8 T cell surface antigens are associated with the internal membrane tyrosine-protein kinase p56lck. Cell. 55, 301–308. Glaichenhaus, N., Shastri, N., Littman, D. R., and Turner, J. M. (1991) Requirement for association of p56lck with CD4 in antigenspecific signal transduction in T cells. Cell. 64, 511–520. Qin, S., Cobbold, S. P., Pope, H., Elliott, J., Kioussis, D., Davies, J., and Waldmann, H. (1993) “Infectious” transplantation tolerance. Science. 259, 974–977. Cobbold, S. P., Adams, E., Zelenika, D., Graca, L., Humm, S., Waldmann, H. (2004) Induction of Foxp3+ regulatory T cells in the periphery of T cell receptor transgenic mice tolerized to transplants. J. Immunol. 172, 6003–6010. Winsor-Hines, D., Merrill, C., O’Mahony, M., Rao, P. E., Cobbold, S. P., Waldmann, H., et al. (2004) Induction of immunological tolerance/hyporesponsiveness in baboons with a nondepleting CD4 antibody. J. Immunol. 173, 4715–4723. Linsley, P. S., Brady, W., Grosmaire, L., Aruffo, A., Damle, N. K., and Ledbetter, J. A.

Organ Transplantation: Modulation of T-Cell Activation Pathways Initiated

24.

25.

26.

27.

28.

29.

30.

31.

32.

(1991) Binding of the B cell activation antigen B7 to CD28 costimulates T cell proliferation and interleukin 2 mRNA accumulation. J. Exp. Med. 173, 721–730. Freeman, G. J., Gray, G. S., Gimmi, C. D., Lombard, D. B., Zhou, L. J., White, M., et al. (1991) Structure, expression, and T cell costimulatory activity of the murine homologue of the human B lymphocyte activation antigen B7. J. Exp. Med. 174, 625–631. Schneider, H., Cai, Y. C., Prasad, K. V., Shoelson, S. E., and Rudd, C. E. (1995) T cell antigen CD28 binds to the GRB-2/SOS complex, regulators of p21ras. Eur. J. Immunol. 25, 1044–1050. Cai, Y. C., Cefai, D., Schneider, H., Raab, M., Nabavi, N., and Rudd, C. E. (1995) Selective CD28pYMNM mutations implicate phosphatidylinositol 3-kinase in CD86-CD28-mediated costimulation. Immunity. 3, 417–426. August, A., Gibson, S., Kawakami, Y., Kawakami, T., Mills, G. B., and Dupont, B. (1994) CD28 is associated with and induces the immediate tyrosine phosphorylation and activation of the Tec family kinase ITK/EMT in the human Jurkat leukemic T-cell line. Proc. Natl. Acad. Sci. USA. 91, 9347–9351. Jones, R. G., Elford, A. R., Parsons, M. J., Wu, L., Krawczyk, C. M., Yeh, W. C., et al. (2002) CD28-dependent activation of protein kinase B/Akt blocks Fas-mediated apoptosis by preventing death-inducing signaling complex assembly. J. Exp. Med. 196, 335–348. Parry, R. V., Whittaker, G. C., Sims, M., Edmead, C. E., Welham, M. J., and Ward, S. J. (2006) Ligation of CD28 stimulates the formation of a multimeric signaling complex involving grb-2-associated binder 2 (gab2), SRC homology phosphatase-2, and phosphatidylinositol 3-kinase: evidence that negative regulation of CD28 signaling requires the gab2 pleckstrin homology domain. J. Immunol. 176, 594–602. August, A., and Dupont, B. (1995) Activation of extracellular signal-regulated protein kinase (ERK/MAP kinase) following CD28 crosslinking: activation in cells lacking p56lck. Tissue Antigen. 46, 155–162. Haspot, F., Séveno, S., Dugast, A. -S., Coulon, F., Renaudin, K., Usal, C., et al. (2005) Anti-CD28 antibody-induced kidney allograft tolerance related to tryptophan degradation and TCR class II B7 regulatory cells. Am. J. Transplant. 5, 2339–2348. Linsley, P. S., Brady, W., Urnes, M., Grosmaire, L. S., Damle, N. K., and Ledbetter, J. A. (1991) CTLA-4 is a second receptor for the B

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

429

cell activation antigen B7. J. Exp. Med. 174, 561–569. Walunas, T. L., Bakker, C. Y., and Bluestone, J. A. (1996) CTLA-4 ligation blocks CD28dependent T cell activation. J. Exp. Med. 183, 2541–2550. Waterhouse, P., Penninger, J. M., Timms, E., Wakeham, A., Shahinian, A., Lee, K. P., et al. (1995) Lymphoproliferative disorders with early lethality in mice deficient in Ctla-4. Science. 270, 985–988. Shiratori, T., Miyatake, S., Ohno, H., Nakaseko, C., Isono, K., Bonifacino, J. S., and Saito, T. (1997) Tyrosine phosphorylation controls internalization of CTLA-4 by regulating its interaction with clathrin-associated adaptor complex AP-2. Immunity. 6, 583–589. Gandhi, A. M., Fazli, U., Rodina, V., and Qazi, Y. A. (2007) Costimulation targeting therapies in organ transplantation. Curr. Opin. Organ Transplant. 13, 622–626. Lee, K. M., Chuang, E., Griffin, M., Khattri, R., Hong, D. K., Zhang, W., et al. (1998) Molecular basis of T cell inactivation by CTLA-4. Science. 282, 2263–2266. Chuang, E., Fisher, T. S., Morgan, R.W., Robbins, M. D., Duerr, J. M., Vander Heiden, M. G., et al. (2000) The CD28 and CTLA-4 receptors associate with the serine/ threonine phosphatase PP2A. Immunity. 13, 313–322. Chikuma, S., Imboden, J. B., and Bluestone, J. A. (2003) Negative regulation of T cell receptor-lipid raft interaction by cytotoxic T lymphocyte-associated antigen 4. J. Exp. Med. 197, 129–135. Nishimura, H., Nose, M., Hiai, H., Minato, N., and Honjo, T. (1999) Development of lupuslike autoimmune diseases by disruption of the PD-1 gene encoding an ITIM motif-carrying immunoreceptor. Immunity. 11, 141–151. Chemnitz, J. M., Parry, R. V., Nichols, K. E., June, C. H., and Riley, J. L. (2004) SHP-1 and SHP-2 associate with immunoreceptor tyrosine-based switch motif of programmed death 1 upon primary human T cell stimulation, but only receptor ligation prevents T cell activation. J. Immunol. 173, 945–954. Sheppard, K.-L., Fitz, L. J., Lee, J. M., Benander, C., George, J. A., Wooters, J., et al. (2004) PD-1 inhibits T-cell receptor induced phosphorylation of the ZAP70/CD3zeta signalosome and downstream signaling to PKCtheta. FEBS Lett. 574, 37–41. Parry, R. V., Chemnitz, R. J., Frauwirth, K. A., Lanfranco, A. R., Braunstein, I., Kobayashi, S. V., et al. (2005) CTLA-4 and PD-1 receptors

430

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

Weatherly and Braun inhibit T-cell activation by distinct mechanisms. Mol. Cell. Biol. 25, 9543–9553. Nishimura, H., Okazaki, T., Tanaka, Y., Nakatani, K., Hara, M., Matsumori, A., et al. (2001) Autoimmune dilated cardiomyopathy in PD-1 receptor-deficient mice. Science. 291, 319–322. Salama, A. D., Chitnis, T., Imitola, J., Ansari, M. J. I., Akiba, H., Tushima, F., et al. (2003) Critical role of the programmed death-1 (PD-1) pathway in regulation of experimental autoimmune encephalomyelitis. J. Exp. Med. 198, 71–78. Ansari, M. J. I., Salama, A. D., Chitnis, T., Smith, R. N., Yagita, H., Akiba, H., et al. (2003) The programmed death-1 (PD-1) pathway regulates autoimmune diabetes in nonobese diabetic (NOD) mice. J. Exp. Med. 198, 63–69. Fife, B. T., Pauken, K. E., Eagar, T. D., Obu, T., Wu, J., Tang, Q., et al. (2009) Interactions between PD-1 and PD-L1 promote tolerance by blocking the TCR-induced stop signal. Nat. Immunol. 10, 1185–1192. Latchman, Y., Wood, C. R., Chernova, T., Chaudhary, D., Borde, M., Chernova, I., et al. (2001) PD-L2 is a second ligand for PD-1 and inhibits T cell activation. Nat. Immunol. 2, 261–268 . Butte, M. J., Keir, M. E., Phamduy, T. B., Sharpe, A. H., and Freeman, G. J. (2007) Programmed death-1 ligand 1 interacts specifically with the B7-1 costimulatory molecule to inhibit T cell responses. Immunity. 27, 111–122. Ozkaynak, E., Wang, L., Goodearl, A., McDonald, K., Qin, S., O’Keefe, T., et al. (2002) Programmed death-1 targeting can promote allograft survival. J. Immunol. 169, 6546–6553. Yang, J., Popoola, J., Khandwala, S., Vadivel, N., Vanguri, V., Yuan, X., et al. (2008) Critical role of donor tissue expression of programmed death ligand-1 in regulating cardiac allograft rejection and vasculopathy. Circulation. 117, 660–669. Elgueta, R., Benson, M. J., de Vries, V. C., Wasiuk, A., Guo, Y., and Noelle, R. J. (2009) Molecular mechanism and function of CD40/ CD40L engagement in the immune system. Immunol. Rev. 229, 152–172. Yuan, X., Dong, V. M., Coito, A. J., Waaga, A.-M., Salama, A. D., Benjamin, C. D., et al.

54.

55.

56.

57.

58.

59.

60.

61.

62.

(2002) A novel CD154 monoclonal antibody in acute and chronic rat vascularized cardiac allograft rejection. Transplantation. 73, 1736–1742. Guillonneau, C., Hill, M., Hubert, F.-X., Chiffoleau, E., Hervé, C., Li, X.-L., et al. (2007) CD40Ig treatment results in allograft acceptance mediated by CD8CD45RC T cells, IFN-gamma, and indoleamine 2,3-dioxygenase. J. Clin. Invest. 117, 1096–1106. Kanmaz, T., Fechner, J. J. H., Torrealba, J., Kim, H. T., Dong, Y., Oberley, T. D., et al. (2004) Monotherapy with the novel human anti-CD154 monoclonal antibody ABI793 in rhesus monkey renal transplantation model. Transplantation. 77, 914–920. Kirk, A. D., Harlan, D. M., Armstrong, N. N., Davis, T. A., Dong, Y., Gray, G. S., et al. (1997) CTLA4-Ig and anti-CD40 ligand prevent renal allograft rejection in primates. Proc. Natl. Acad. Sci. USA. 94, 8789–8794. Ramirez, C. B., and Marino, I. R. (2007) The role of basiliximab induction therapy in organ transplantation. Expert. Opin. Biol. Ther. 7, 137–148. Lúdvíksson, B. R., Gray, B., Strober, W., and Ehrhardt, R.O. (1997) Dysregulated intrathymic development in the IL-2-deficient mouse leads to colitis-inducing thymocytes. J. Immunol. 158, 104–111. Fontenot, J. D., Rasmussen, J. P., Gavin, M. A., and Rudensky, A. Y. (2005) A function for interleukin 2 in Foxp3-expressing regulatory T cells. Nat. Immunol. 6, 1142–1151. Dai, Z., Konieczny, B. T., Baddoura, F. K., and Lakkis, F. G. (1998) Impaired alloantigen-mediated T cell apoptosis and failure to induce long-term allograft survival in IL-2deficient mice. J. Immunol. 161, 1659–1663. Kuttler, B., Kauert, C., Wanka, H., Diamantstein, T., and Hahn, H. J. (1996) Temporary anti-CD25/CsA therapy induces a CD4+ T-cell-mediated tolerance in BB/OK rats. J. Autoimmun. 9, 321–329. Bielekova, B., Catalfamo, M., ReichertScrivner S., Paker, A., Cerna, M., Waldmann, T. A., et al. (2006) Regulatory CD56(bright) natural killer cells mediate immunomodulatory effects of IL-2Ralpha-targeted therapy (daclizumab) in multiple sclerosis. Proc. Natl. Acad. Sci. USA. 103, 5941–5946.

Chapter 27 Immunosuppressive Mechanisms During Viral Infectious Diseases Ghanashyam Sarikonda and Matthias G. von Herrath Abstract For a virus to establish persistence in the host, it has to exploit the host immune system such that the active T-cell responses against the virus are curbed. On the other hand, the goal of the immune system is to clear the virus, following which the immune responses need to be downregulated, by a process known as immunoregulation. There are multiple known immunoregulatory mechanisms that appear to play a role in persistent viral infections. In the recent past, IL-10 and PD-1 have been identified to be playing a significant role in the regulation of antiviral immune responses. The evidence that viruses can escape immunologic attack by taking advantage of the host’s immune system is found in LCMV infection of mice and in humans persistently infected with HIV and HCV. The recent observation that the functionally inactive T-cells during chronic viral infections can be made to regain their cytokine secretion and cytolytic abilities is very encouraging. Thus, it would be likely that neutralization negative immune regulation during persistent viral infection would result in the preservation of effector T-cell responses against the virus, thereby resulting in the elimination of the persistent infection. Key words: Immunoregulation, Persistent infection, IL-10, PD-1, LCMV

1. Virus and Disease Most viruses that infect humans are usually cleared by the host immune system in a rapid (acute) manner, either (a) by the release of proinflammatory cytokines or (b) by killing the infected cells to limit the replication of the virus [1]. Such acute responses involve a first step innate immune response (macrophages, dendritic cells [DCs], and natural killer [NK] cells) [2], followed by the generation of adaptive immunity (T and B cells). Macrophages and DCs act as phagocytes thus eliminating the infected cells, and NK cells directly kill the infected cells. In adaptive immunity, which kicks

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_27, © Springer Science+Business Media, LLC 2011

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in 3–5 days after infection, presentation of viral antigen by macrophages and/or DCs or by infected cells that present viral peptides on host MHC molecules results in the activation of T-cells. These T-cells then mount a viral antigen-specific immune response and kill the infected cells either in a cell–cell contact-dependent manner (perforin/granzyme mediated) or through the release of inflammatory cytokines TNF-a or IFN-g. These cytokines can also act on the virus directly and prevent its replication. Some of the viral antigen-specific T-cells then survive for a very long time creating a memory pool. Most, if not all, viral infections trigger the production of interferon’s (IFNs) by the host immune system. IFNs then primarily limit viral replication to prevent damage to the infected cell. Type-I IFNs are essential part of innate resistance to viral infections as is evident in the case B6 mice infected with MHV-1; absence of type I IFN-mediated signaling in IFN-abR-KO mice results in progressive loss of body weight, culminating in the death of the mice by day 5 postinfection [3]. Intravenous infection of adult mice with lymphocytic choriomeningitis virus (LCMV)-Armstrong results in an acute infection, which is resolved by virus-specific CD8 and CD4 T cells within 8–10 days [4]. However, unlike in acute viral immune response where antiviral CTLs are present at all times examined; in persistent infections, whether it be the effector phase or memory phase, antiviral CD8 T-cells are rarely found in adult mice or in humans infected early in life [1]. A more detailed discussion of innate and adaptive immune responses in viral infections is beyond the scope of this chapter. Some virus, for instance HIV and hepatitis B virus (HBV)/ hepatitis C virus (HCV) in humans and LCMV in mice, escape killing by the host’s immune system, either by avoidance or manipulation and establish persistent infection in the host. To establish a persistent infection, the virus has to overcome several host-dependent factors. One unique strategy that viruses employ is that instead of killing its host cell, the virus causes little to no damage so that it can escape detection by the immune system. The virus then either continues to replicate inside the cell (HIV) or it establishes latency (EBV). Viruses can alter or interfere with the processing of viral peptides by professional antigen presenting cell (APC), a requirement for activation and expansion of the T cells that normally remove infected cells. Additionally, viruses can also inhibit the differentiation of antigen-presenting conventional DCs and can infect effector T and B cells directly [1]. Some viruses also encode miRNAs to regulate their replication or latency or to manipulate or evade host immune responses [5]. During latency, miRNAs keep protein levels of viral genes to a minimum, facilitating evasion of immune surveillance [6]. Finally, viruses also exploit the host’s immunoregulatory mechanisms such that

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the viral antigen-specific T-cells now become either anergic or are deleted and thus fail to clear the viral infection. This strategy is more detrimental to the host. Persistent viral infections, as would be expected, are a severe problem to humans, as is evident in the case infection with HIV; persistent infection with HIV leads to the exhaustion of CD4 T-cells thereby rendering the host susceptible to a variety of secondary infections, which ultimately lead to the demise of the host. However, such persistent infections, if are actively sought by the host immune system, due to the continued activation of T-cells can lead to severe immune pathology, wherein the host’s immune system destroys the host’s own tissue. There is a copious amount of data that are published, which provides evidence for the deleterious effects of an over-active immune system causing sever immune pathology in persistent viral infections. For instance, overexuberant immune response is implicated in fatal H5N1 influenza [7] and a dysregulated immune response contributes to clinical disease in patients with SARS [8, 9]. Similarly, in mice, excessive T-cell responses contribute to the tissue damage that occurs during the process of virus clearance in infections with LCMV, HSV, RSV [10–12], and MHV-1 [13]. 1.1. LCMV as a Model to Study Immune Regulation

In our laboratory, we have extensively used LCMV as a model for establishing persistent infection in mice [14–20]. It is easier to dissect the various immunoregulatory mechanisms involved in viral infections using LCMV due to the availability of two different strains; one that establishes acute infection (LCMVArmstrong) and is readily cleared from the host and the other that establishes persistent infection (LCMV-clone 13) for a very long time. Armstrong and clone 13 viruses share identical CD4 and CD8 T-cell epitopes but are phenotypically very distinct, replicate in different cell types, and have different clearance rates [21]. Further, since LCMV is not a cytolytic virus, it allows to distinctly examine the effects caused by the immune responsemediated damage to the host tissue [22]. Persistent viral infections ultimately result in immunosuppression in the host where the CTLs lose functionality by either functional inactivation and/or physical deletion [1]. Several different mechanisms, which act either individually or in combination, including clonal exhaustion, secretion of immunosuppressive cytokines such as IL-10 and TGF-b, and overexpression of programmed death 1 (PD-1) on activated T-cells, contribute to the failure of T-cell responses during persistent viral infections. In LCMV-clone 13 model of persistent infection, multiple immunoregulatory mechanisms are activated, including infection/impairment of DCs, and a global inactivation of the virus-specific T-cell response [23–27]. The antiviral CD4 and CD8 T-cells are either physically deleted or persist in an inactivated state and are

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nonresponsive toward viral antigens [23–27]. With respect to the significance of such inactivation in human patients, in HIV infection, CD4 T cells that can produce TNF-a, interleukin 2 (IL-2), and IFN-g are conspicuously absent. Consequently, in HIV+ patients, the presence of functionally active CD4 T-cells that can produce the cytokines TNF-a, IFN-g, and IL-2 directly correlates with low viremia [28]. Similarly, in mouse models of persistent infection, a significant three- and fourfold decrease in TNF-a and IL-2 producing cells, respectively, was observed in the spleens of clone 13 infected mice by day 100 postinfection [23]. However, unlike it was believed before, it is increasingly apparent that in persistent infections, the effector T-cells are not deleted but exist in a functionally inactive state. Thus, it was interesting to note that despite prolonged periods of inactivation, the remaining CD4 and CD8 T-cells still retain the ability to recover their functional activity [29]. This suggests that, in persistent viral infections, the host’s immune system can still be manipulated such that the host can reestablish immunity against the virus. Thus, functional preservation of CD4 and CD8 T-cells during viral persistence is the ultimate mode of achieving strong antiviral immunity and viral clearance.

2. Immune Regulation 2.1. General

As discussed previously, for a virus to establish persistence in the host, it has to exploit the host immune system such that the active T-cell responses against the virus are curbed. On the other hand, the goal of the immune system is to clear the virus, failing which the immune responses need to be downregulated so as to avoid unnecessary damage to the host tissue, by a process known as immunoregulation. The various known immunoregulatory mechanisms comprise (a) activation or induction of Tregs, (b) expression of inhibitory molecules such as programmed cell death-1 (PD-1) on activated T-cells, and (c) production of suppressor cytokines such as TGF-b or IL-10. In the recent past, among the major immunoregulatory molecules identified so far [15, 30–32], IL-10 [15, 32] and PD-1 [30] appear to be playing a significant role in the regulation of antiviral immune responses and are thus being actively investigated. The confounding aspect of all these different factors is that they not only regulate antiviral immune responses but they are also involved in the regulation of autoimmune responses. The evidence that viruses can escape immunologic attack by taking advantage of the host’s immune system is found in LCMV infection of mice [15, 30, 32] and in persistently infected with HIV and HCV [33–37]. Thus, it is important to understand the effectors involved in active immunoregulation, such that we can

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define how a virus exploits the host’s immunoregulatory mechanism and thereby design drugs that can circumvent this virally induced immunoregulation. 2.2. Immune Regulation by Innate Immunity

This aspect of the immune system has been recently reviewed, for details refer McGill et al. [2]. The innate immune system mainly comprises macrophages, DCs, and NK cells. Macrophages and DCs are primarily involved in antigen presentation to the T-cells, with DCs usually being the more efficient APC. NK cells are primarily involved in the direct killing of virally infected cells as part of the innate immune protection. However, as with adaptive immune system, hyperactive innate immune system could very well lead to tissue/organ damage. A direct evidence for innate immune-mediated pathology is evident in the case of fatal H5N1 influenza [7]. Here, we will focus only on those cell populations that exhibit immune regulatory function.

2.2.1. Macrophages

Although macrophages are not the most efficient of APCs, they are very efficient phagocytes. And thus, it is believed that macrophages may be a predominant cell population contributing to IAV-associated immunopathology [39]. The first evidence for the role of macrophages in immune regulation was evident in alveolar macrophages that exist in a relatively quiescent state during homeostasis but have a regulatory phenotype in the lungs [40]. In addition, Dillon et al. showed that stimulation of macrophages through the TLR2 receptor induces macrophages that promote immunological tolerance through the production of TGF-b1 [41]. And finally, macrophages have been shown to suppress the induction of innate and adaptive immunity [42–44].

2.2.2. Dendritic Cells

DCs are probably the most efficient APCs of the immune system, both in animal models and humans. Therefore, the initiation of effective Ag-specific immunity to pathogens is a hallmark of DC function. However, it was also demonstrated that DCs can induce and maintain self-antigen-specific tolerance in the periphery [45]. Further, DCs can actively induce T-cell anergy [46, 47], T-cell suppression [47, 48], and generation of Tregs [49]. As they did for macrophages, Dillon et al. also showed that stimulation through the TLR2 receptor on DCs induces regulatory DC [41]. Further, TLR2 activation in mice results in IL-10 upregulation and Treg survival [50, 51]. Interestingly, incubation of DCs in the presence of IL-10 generates DC with tolerogenic properties [52]. Thus, DCs, when activated through the TLR2 receptor, appear to acquire regulatory phenotype and indeed are able to suppress the activation antigen-specific T-cells. In support for the role of DC in immune regulation in viral infections, in in vitro experiments, it was found that the infection

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of DCs with high MOI results in truncated CD8 T-cell responses and increased IL-12p40 production [53]. Further, such high MOI infection also results in an increased production of the antiinflammatory cytokines IL-10 and TGF-b [54]. Finally, in IAV infections, rather than inducing T-cell activation, when influenza antigen-bearing immature DCs encounter naïve T cells, they induce tolerance to viral antigens [55, 56]. The immunoregulatory role played by DCs in other viral infections needs to be explored. Thus, under certain conditions, for instance activation through TLR2, both DCs and macrophages appear to acquire an immunoregulatory function. However, whether macrophages and DCs can exhibit similar activity in a majority of persistent viral infections needs to be determined. Be as it may, it would be interesting to speculate that since viral infections result in TLR2 activation, at least in infections with some viruses (chronic infections), such TLR2 activation induces tolerogenic DCs and/or macrophages, which prevent further activation of antiviral T-cells. 2.3. Immune Regulation by Adaptive Immunity 2.3.1. CD4+ Tregs

One of the most extensively studied cell population in immunology is CD4+CD25+ regulatory T-cells (Tregs), because of their demonstrated importance in the events leading to autoimmunity. Thus, doing a formal detailed review of Tregs takes a very extensive review of literature and is therefore beyond the scope of this review article. But, for those who are interested, for a detailed review of Tregs, refer to Sakaguchi et al. [57]. There exist mainly two types of Tregs, the natural Tregs (nTregs) that originate from the thymus and the induced Tregs (iTregs) that arise in the periphery. Tregs, either natural or induced, are characterized by the expression of the transcription factor Foxp3. Thus, it was found that the ectopic Foxp3 expression in normal T-cells enables these cells to exhibit suppressor function in vitro and in vivo [58]. Although the significance still needs to be determined, it was recently demonstrated that the expression of Foxp3 in Treg cells does not destine the T-cell to be a Treg for life, since a subset of Treg cells appear to downregulate Foxp3 expression (exFoxp3 cells) [59]. nTregs constitute 4–5% of peripheral CD4 T cells and are important in the maintenance of immunological self-tolerance in the periphery [60]. CD25+ T cells appear in the periphery after day 3 and rapidly increase to adult levels within 2 weeks (reviewed in [61, 62]). CD4+CD25+ Tregs are actively involved in the regulation of antiviral immunity in order to limit immune pathology in the host [63–67] but thereby they also inhibit viral clearance [68–70]. Tregs have been implicated to play a role in a number of different viral infections, in both mice and humans, including but not limited to Friends’ virus or HSV (in mice) and HCV and HIV (in humans) [12, 68–72]. To further demonstrate the importance of Tregs in viral immunity (or immune regulation), Tregs are

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relatively deficient in patients with severe dengue infection when compared to those with mild disease [73]. Tregs exert their suppressive function through a combination of various mechanisms including cell-contact (killing and/or functional modulation of APC and/or responder T cells (reviewed in [74–76])) and soluble factors (secretion of immunosuppressive cytokines, IL-10 and TGF-b [77, 78]). However, it appears that direct cell-contact-mediated suppression is critical for the action of Tregs. Tregs inhibit, via CTLA-4, the upregulation of CD80 and CD86 by immature DC upon antigenic stimulation and also downregulate the expression of CD80/CD86 by mature DC [79, 80], thus indirectly affecting antiviral CD4 and CD8 T-cell responses. Another plausible mechanism by which Tregs mediate their regulatory activity has been proposed; since Tregs cannot produce IL-2 and thus cannot expand in an autocrine fashion [81], but constitutively express high levels of IL-2R (CD25), they likely deprive effector T-cells of IL-2 in the vicinity and thus provide hindrance to the activation/expansion of responder T cells [82]. As such, the take home point is that natural CD4+CD25+ Tregs have only a minimal impact on viral elimination per se but they play an important part in limiting collateral tissue damage usually caused by strong antiviral T-cell responses [12, 66]. Thus, they are one of the most important players in mediating immune regulation, either in antiviral immunity or in autoimmunity. 2.3.2. PD-1/PD-L1

Based on few recent findings [30, 31, 35, 36, 38, 83–86], it is increasingly apparent that the PD-1/PDL-1 axis plays a major role in regulating immune responses during antiviral T-cell responses. PD-1 is an inhibitory receptor of the CD28 family and has two ligands PD-L1 and PD-L2, but most of the immunoregulatory activity of PD-1 is brought upon by its interaction with PD-L1 (reviewed in [87]). Accordingly, PD-L1−/− mice infected with LCMV clone 13 died owing to immunopathologic damage [30]. During chronic infection with LCMV-clone 13, PD-1 is upregulated at both RNA and protein level, in exhausted LCMVspecific CD8 T cells. Further, PD-1 expression continued to increase in exhausted CD8 T cells and the high level of expression was sustained resulting in the failure of these to clear infection [30, 31]. Blocking the activity of PD-1, using an antibody against PD-L1, enhanced virus-specific CD8 T cell proliferation, production of IFN-g, and their lytic ability [30]. And further, blocking PD-L1 results in enhanced epitope-specific responses to therapeutic vaccination [84]. In our laboratory, using LCMV-induced T1D model, the RIP-GP-LCMV mice, we found that the virus-induced upregulation of PD-L1 in prediabetic mice prevents the expansion of diabetogenic CD8+ T-cells, thus confirming the importance of PD-1 in regulating antiviral CD8 T-cell responses [85].

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Interestingly, such PD-L1 upregulation also resulted in synergistic enhancement in the capacity of virally enhanced CD4+CD25+ Tregs to mediate immunoregulation [83]. Finally, in humans infected with HIV who have increased PD-1, an association exists between in vivo T-cell unresponsiveness and restoration of T-cell function in vitro with antibody to PD-L1 [36, 85, 86]. Thus, the PD-1–PD-L1 axis is crucial for sustaining suppression of CD8 T cells during persistent infection. 2.3.3. CTLA-4

Costimulation through the CD28 molecule is essential for the generation of efficient antiviral T-cell responses. Consequently, the prevention of costimulation during the chronic phase of clone 13 infections diminishes antiviral CD8 T cell responses and prevents control of viral replication [88]. CTLA-4, an inhibitory receptor belonging to the B7-CD28 family of costimulatory molecules (reviewed in [89]), is constitutively expressed on murine Foxp3+ Treg, whether thymus derived or induced in the periphery [90–92] and can mediate the suppression of effector T-cell responses. CTLA-4 is also upregulated on activated T cells, and competes with CD28 for binding to the same ligands, B7-1 (CD80) and B7-2 (CD86). However, instead of inducing activation of T-cells, such as CD28, CTLA-4 antagonizes the production of IL-2 and inhibits T-cell activation. While the importance of CTLA-4-mediated regulation T-cell responses in tumor immunology is well known, its role in persistent viral infections is still debatable. In mice infected with LCMV, CTLA-4 expression or lack thereof by T cells has little or no effect on antiviral immunity, either in the effector or in memory generation during acute viral infection [30]. And, similar to acute infection, CTLA-4 was also not involved in T-cell exhaustion in mice chronically infected with LCMV clone 13 [30]. The strongest argument for the involvement of CTLA-4 in the regulation of antiviral immunity comes from the studies of HIV-infected humans (reviewed in [93]) or SIV-infected monkeys [94]. Arguing for a role of CTLA-4 in antiviral immunity, it was found that CTLA-4 expression was high in HIV-specific CD4 but not CD8 T cells in humans with acute HIV infection but low in people who were able to spontaneously control infection [95]. More specifically, consistent with the finding that CTLA-4 engagement downregulates IL-2 secretion [96], HIV-Gagspecific CD4 T cells that produced IL-2 in addition to IFN-g had less CTLA-4, while those CD4 T cells that produced only IFN-g had higher CTLA-4 expression [95]. And finally, it was shown that blockade of CTLA-4 with antibodies resulted in an augmentation of both SIV-specific CD4 and CD8 efffector T-cell responses [94].

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Thus, CTLA-4 plays a major role in immunoregulation by preventing costimulation through CD28 receptor leading to inefficient activation of CD8 T-cells, in some, if not all, persistent viral infections. Hence, therapeutic approaches aimed at preventing the action of CTLA-4 during some persistent infections, where CTLA-4 is known to play a role, will be beneficial to a subgroup of patients. 2.3.4. TGF-b

With the discovery that Tregs make an immunoregulatory cytokine TGF-b, quite a few laboratories performed extensive analysis of the role played by this cytokine in immune regulation (reviewed in [97]). It is now well known that TGF-b controls various aspects of the inflammatory response through the regulation of chemotaxis, activation, and survival of T cells, NK cells, and APCs and is crucial for the control of antiviral immunity (reviewed in [97]). TGF-b also exhibits highly potent inhibitory effects on cytolytic functions, cytokine production, and proliferation of T-cells (reviewed in [98]). Conversely, TGF-b also exhibits antiapoptotic effects [99–101] on T-cells best exemplified in the case of the effector T-cells. When cultured in the presence of TGF-b and IL-2 effector, CD4 T-cells exhibit enhanced accumulation (owing to increased proliferation and survival) and even better prolonged expansion [99]. And finally, TGF-b, produced by Tregs, can act in an autocrine fashion to induce further expansion of Tregs from which it is produced. Also, TGF-b can induce the conversion of naïve CD4+CD25− T cells into CD4+CD25+ T cells by the induction of Foxp3 [102]. Thus TGF-b is extensively used to induce the expansion of Treg in in vitro cultures. Data from our laboratory as well as others have now shown that such in vitro expanded Tregs are able to adoptively confer protection in virus-induced T1D model (reviewed in [103]). While the role of TGF-b in regulating immune responses is extensively known in a variety of autoimmune, infectious (bacterial) and other diseases evidence of TGF-b role in regulating antiviral immunity is scarce. Still, in chronic infections, such as influenza, HIV, HBV, HCV, and others, there is evidence of increased TGF-b production. In a virus-induced T1D mouse model, data from our laboratory showed that on one hand TGF-b suppresses naïve CD8+ T cell activation, while on the other hand it enhances the survival and function of antigenexperienced/memory CD8 T cells [104]. Most importantly, systemic administration of TGF-b was shown to protect from T1D [105]. Interestingly, several recent studies have convincingly demonstrated that TGF-b plays an essential role in the development of a newly discovered highly proinflammatory T-cells, the IL-17secreting T cells (Th17) [106]. Although, TGF-b cannot induce

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the conversion of naïve cells to Th17 by itself, together with IL-6, TGF-b facilitates the differentiation of T-cells into Th17 cells [107]. The most important aspect of TGF-b-induced effect is that TGF-b in conjunction with IL-2 induces the differentiation of naïve T-cells into Foxp3+ Tregs and this IL-2 inhibits the differentiation into Th17 cells [108]. Thus TGF-b is a very interesting cytokine, in that, in the presence of IL-2, it induces the generation of regulatory T-cells, CD4+ Foxp3+ Tregs, but the same TGF-b, in the presence of IL-6, induces the generation of proinflammatory Th17 T-cells. Hence, TGF-b apparently is very important in the regulation of antiviral immunity because while Tregs (as discussed above) are clearly involved in preventing tissue damage due to immune pathology, Th17 cells are highly proinflammatory and are known to be playing a significant role in autoimmune diseases. Thus, this dual functionality of TGF-b needs further elucidation. 2.3.5. IL-10

One of the most studied regulatory cytokine other than TGF-b is IL-10. IL-10 exerts its suppressive effect on macrophages [109] and T cells by blocking CD28 and ICOS signals in a rapid signal transduction cascade [110]. The best example for the role of IL-10, in regulating immune responses in viral infections, was provided in the last few years by a couple of laboratories, including data from our own laboratory. It is known that by 9 days after infection with LCMV clone 13, which establishes persistent infection in adult mice, both virusspecific CD4+ and CD8+ T-cells become unresponsive. Data from our laboratory showed that the use of an antibody to the IL10 receptor results in very efficient control of viral persistence, even leading to the elimination of virus. By removing IL-10 inhibitory effect, in this case on T-cell anergy, we were able to induce host resistance such that the host could now resolve persistent LCMV infection [15]. This was followed by further proof that such IL-10 acts on effector T-cells [32]. When IL-10 was blocked with antibody to IL-10 receptors, T-cell numbers increase sufficiently to clear virus from blood and tissues; these T-cells that were anergized because of the persistent virus infection now regained their ability to produced proinflammatory cytokines like TNF-a, IFN-g and thus were able to reacquire their normal effector T-cell functions [32]. There was a significant increase in the frequency of IL-10 producing cells in both spleen and liver during persistent clone 13 infection [23]. Accordingly, significantly more IL-10 RNA was found in the spleens of clone 13-infected mice than that of Armstrong-infected mice [32]. CD4+CD25+ Foxp3+ regulatory T-cells are thought to be the major producers of IL-10. However, in clone 13 persistent infection, although CD4 T-cells

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produced some IL-10 initially, they are not responsible for the majority of IL-10, since IL-10 was produced in significant quantities even after the functional inactivation of CD4 T-cells; further these CD4 T-cells stopped making any IL-10 after functional inactivation [32]. Interestingly, instead of the CD4 T-cells, it appears that DCs are the major producers of IL-10 in clone 13 infection [32]. Further, using IL-10−/− mice, the authors found that the lack of IL-10 resulted in the preservation of viral antigen (NP396-404)-specific CD8 T-cell responses that are normally lost during persistent infection [32]. And, as we had reported originally [15], antibody blockade of IL-10 prevented viral persistence [32], thus definitely showing that IL-10 plays a major role in immunoregulation during persistent viral infection with LCMV. Although it appears that IL-10 may be acting directly on T-cells, the involvement of Tregs in immunoregulation but the their failure to make IL-10 during persistent LCMV infection suggests that other than acting on T-cells, IL-10 may also be playing an immunoregulatory role by directly altering the antigen-presenting capabilities of the APCs. However, this hypothesis needs further testing since it was found that APC functionality is normal in terms of antigen presentation in clone 13 infection [23].

3. Concluding Remarks Persistent viral infections are a real threat to human health during a variety of viral infections with HIV, HBV, and HCV. The major role players responsible for the immunosuppression that happens during persistent infections have been identified as PD-1/PD-L1 and IL-10, with TGF-b and CTLA-4 also playing a role to some extent (Fig. 1). The recent observation that the functionally inactive T-cells during chronic viral infections can be made to regain their cytokine secretion and cytolytic abilities by neutralizing IL-10 is very encouraging [32]. More importantly, when IL-10 [111] or PD-L1 [84] was first neutralized, therapeutic vaccination against ongoing persistent LCMV infection was highly effective. Further, IL-10R blockade resulted in the ability of an otherwise ineffective DNA vaccination to be successful such that the DNA vaccination elicited a fourfold increase in the number of functional CD8 T cells [111]. Thus, it would be likely that neutralization of either IL-10 or PD-1/PD-L1 (depending on the virus) during the persistent viral infection would result in the preservation of effector T-cell responses against the virus thereby resulting in the elimination of the persistent infection.

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Fig. 1. Immune regulation during persistent viral infection. In an acute infection (left), when a virus infects a normal cell, it is phagocytosed by antigen presenting cell (APC) (dendritic cell [DC] or MF). The APCs efficiently present the viral antigens to CD4 and CD8 cells, which either release proinflammatory cytokines (TNF-a, IFN-g) or upregulate perforin, which kills the infected cells, ultimately leading to the elimination of the virus. However, in persistent infection (right), due to a number of reasons, including miRNAs and molecular mimicry, viruses evade immune recognition. Thus, the CD4 and CD8 cells are not sufficiently activated and the virus avoids immune-mediated elimination. On the other hand, APCs (MF and DCs), upon TLR2 activation, upregulate IL-10; IL-10 exhibits significant immunosuppressive effects leading to the inactivation of CD4 and CD8 T-cells. Further, activated T-cells also upregulate CTLA-4 that prevents costimulation, thereby leading to inactivation. And, finally, virus antigen-specific T-cells also upregulate PD-1 which, upon engagement with its ligand PD-L1, signals the death of that T-cell. These events, help to avoid immune response mediated host tissue damage but at the cost of establishing viral persistence.

References 1. Oldstone, M. B. (2009) Anatomy of viral persistence, PLoS Pathog 5, e1000523. 2. McGill, J., Heusel, J. W., and Legge, K. L. (2009) Innate immune control and regulation of influenza virus infections, J Leukoc Biol 86, 803–812. 3. Khanolkar, A., Hartwig, S. M., Haag, B. A., Meyerholz, D. K., Epping, L. L., Haring, J. S., Varga, S. M., and Harty, J. T. (2009) Protective and pathologic roles of the immune response to mouse hepatitis virus type 1: implications for severe acute respiratory syndrome, J Virol 83, 9258–9272. 4. Fung-Leung, W. P., Kundig, T. M., Zinkernagel, R. M., and Mak, T. W. (1991) Immune response against lymphocytic choriomeningitis virus infection in mice without CD8 expression, J Exp Med 174, 1425–1429. 5. Gottwein, E., and Cullen, B. R. (2008) Viral and cellular microRNAs as determinants of viral pathogenesis and immunity, Cell Host Microbe 3, 375–387.

6. Huang, J., Wang, F., Argyris, E., Chen, K., Liang, Z., Tian, H., Huang, W., Squires, K., Verlinghieri, G., and Zhang, H. (2007) Cellular microRNAs contribute to HIV-1 latency in resting primary CD4+ T lymphocytes, Nat Med 13, 1241–1247. 7. de Jong, M. D., Simmons, C. P., Thanh, T. T., Hien, V. M., Smith, G. J., Chau, T. N., Hoang, D. M., Chau, N. V., Khanh, T. H., Dong, V. C., Qui, P. T., Cam, B. V., Ha do, Q., Guan, Y., Peiris, J. S., Chinh, N. T., Hien, T. T., and Farrar, J. (2006) Fatal outcome of human influenza A (H5N1) is associated with high viral load and hypercytokinemia, Nat Med 12, 1203–1207. 8. Cameron, M. J., Ran, L., Xu, L., Danesh, A., Bermejo-Martin, J. F., Cameron, C. M., Muller, M. P., Gold, W. L., Richardson, S. E., Poutanen, S. M., Willey, B. M., DeVries, M. E., Fang, Y., Seneviratne, C., Bosinger, S. E., Persad, D., Wilkinson, P., Greller, L. D., Somogyi, R., Humar, A., Keshavjee, S., Louie, M., Loeb, M. B., Brunton, J., McGeer, A. J., and Kelvin, D. J. (2007) Interferon-mediated immunopathologi-

Immunosuppressive Mechanisms During Viral Infectious Diseases

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

cal events are associated with atypical innate and adaptive immune responses in patients with severe acute respiratory syndrome, J Virol 81, 8692–8706. Jones, B. M., Ma, E. S., Peiris, J. S., Wong, P. C., Ho, J. C., Lam, B., Lai, K. N., and Tsang, K. W. (2004) Prolonged disturbances of in vitro cytokine production in patients with severe acute respiratory syndrome (SARS) treated with ribavirin and steroids, Clin Exp Immunol 135, 467–473. Hussell, T., Pennycook, A., and Openshaw, P. J. (2001) Inhibition of tumor necrosis factor reduces the severity of virus-specific lung immunopathology, Eur J Immunol 31, 2566–2573. Oldstone, M. B. (2002) Biology and pathogenesis of lymphocytic choriomeningitis virus infection, Curr Top Microbiol Immunol 263, 83–117. Suvas, S., Azkur, A. K., Kim, B. S., Kumaraguru, U., and Rouse, B. T. (2004) CD4+CD25+ regulatory T cells control the severity of viral immunoinflammatory lesions, J Immunol 172, 4123–4132. De Albuquerque, N., Baig, E., Ma, X., Zhang, J., He, W., Rowe, A., Habal, M., Liu, M., Shalev, I., Downey, G. P., Gorczynski, R., Butany, J., Leibowitz, J., Weiss, S. R., McGilvray, I. D., Phillips, M. J., Fish, E. N., and Levy, G. A. (2006) Murine hepatitis virus strain 1 produces a clinically relevant model of severe acute respiratory syndrome in A/J mice, J Virol 80, 10382–10394. Bot, A., Rodrigo, E., Wolfe, T., Bot, S., and Von Herrath, M. G. (2003) Infectiontriggered regulatory mechanisms override the role of STAT 4 in control of the immune response to influenza virus antigens, J Virol 77, 5794–5800. Ejrnaes, M., Filippi, C. M., Martinic, M. M., Ling, E. M., Togher, L. M., Crotty, S., and von Herrath, M. G. (2006) Resolution of a chronic viral infection after interleukin-10 receptor blockade, J Exp Med 203, 2461–2472. Ejrnaes, M., von Herrath, M. G., and Christen, U. (2006) Cure of chronic viral infection and virusinduced type 1 diabetes by neutralizing antibodies, Clin Dev Immunol 13, 337–347. Filippi, C. M., and von Herrath, M. G. (2008) IL-10 and the resolution of infections, J Pathol 214, 224–230. Homann, D., Dummer, W., Wolfe, T., Rodrigo, E., Theofilopoulos, A. N., Oldstone, M. B., and von Herrath, M. G. (2006) Lack of intrinsic CTLA-4 expression has minimal effect on regulation of antiviral T-cell immunity, J Virol 80, 270–280. von Herrath, M. G., Berger, D. P., Homann, D., Tishon, T., Sette, A., and Oldstone, M. B.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

443

(2000) Vaccination to treat persistent viral infection, Virology 268, 411–419. Wolfe, T., Asseman, C., Hughes, A., Matsue, H., Takashima, A., and von Herrath, M. G. (2002) Reduction of antiviral CD8 lymphocytes in vivo with dendritic cells expressing Fas ligand-increased survival of viral (lymphocytic choriomeningitis virus) central nervous system infection, J Immunol 169, 4867–4872. Fuller, M. J., Khanolkar, A., Tebo, A. E., and Zajac, A. J. (2004) Maintenance, loss, and resurgence of T cell responses during acute, protracted, and chronic viral infections, J Immunol 172, 4204–4214. Oldstone, M. B. (2007) A suspenseful game of “hide and seek” between virus and host, Nat Immunol 8, 325–327. Brooks, D. G., Teyton, L., Oldstone, M. B., and McGavern, D. B. (2005) Intrinsic functional dysregulation of CD4 T cells occurs rapidly following persistent viral infection, J Virol 79, 10514–10527. Moskophidis, D., Lechner, F., Pircher, H., and Zinkernagel, R. M. (1993) Virus persistence in acutely infected immunocompetent mice by exhaustion of antiviral cytotoxic effector T cells, Nature 362, 758–761. Zajac, A. J., Blattman, J. N., Murali-Krishna, K., Sourdive, D. J., Suresh, M., Altman, J. D., and Ahmed, R. (1998) Viral immune evasion due to persistence of activated T cells without effector function, J Exp Med 188, 2205–2213. Ou, R., Zhou, S., Huang, L., and Moskophidis, D. (2001) Critical role for alpha/beta and gamma interferons in persistence of lymphocytic choriomeningitis virus by clonal exhaustion of cytotoxic T cells, J Virol 75, 8407–8423. Wherry, E. J., Blattman, J. N., Murali-Krishna, K., van der Most, R., and Ahmed, R. (2003) Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment, J Virol 77, 4911–4927. Kannanganat, S., Kapogiannis, B. G., Ibegbu, C., Chennareddi, L., Goepfert, P., Robinson, H. L., Lennox, J., and Amara, R. R. (2007) Human immunodeficiency virus type 1 controllers but not noncontrollers maintain CD4 T cells coexpressing three cytokines, J Virol 81, 12071–12076. Brooks, D. G., McGavern, D. B., and Oldstone, M. B. (2006) Reprogramming of antiviral T cells prevents inactivation and restores T cell activity during persistent viral infection, J Clin Invest 116, 1675–1685. Barber, D. L., Wherry, E. J., Masopust, D., Zhu, B., Allison, J. P., Sharpe, A. H., Freeman, G. J., and Ahmed, R. (2006) Restoring function in

444

31.

32.

33.

34.

35.

36.

37.

38.

Sarikonda and von Herrath exhausted CD8 T cells during chronic viral infection, Nature 439, 682–687. Blackburn, S. D., Shin, H., Haining, W. N., Zou, T., Workman, C. J., Polley, A., Betts, M. R., Freeman, G. J., Vignali, D. A., and Wherry, E. J. (2009) Coregulation of CD8+ T cell exhaustion by multiple inhibitory receptors during chronic viral infection, Nat Immunol 10, 29–37. Brooks, D. G., Trifilo, M. J., Edelmann, K. H., Teyton, L., McGavern, D. B., and Oldstone, M. B. (2006) Interleukin-10 determines viral clearance or persistence in vivo, Nat Med 12, 1301–1309. Brockman, M. A., Kwon, D. S., Tighe, D. P., Pavlik, D. F., Rosato, P. C., Sela, J., Porichis, F., Le Gall, S., Waring, M. T., Moss, K., Jessen, H., Pereyra, F., Kavanagh, D. G., Walker, B. D., and Kaufmann, D. E. (2009) IL-10 is up-regulated in multiple cell types during viremic HIV infection and reversibly inhibits virus-specific T cells, Blood 114, 346–356. Barrett, L., Gallant, M., Howley, C., Bowmer, M. I., Hirsch, G., Peltekian, K., and Grant, M. (2008) Enhanced IL-10 production in response to hepatitis C virus proteins by peripheral blood mononuclear cells from human immunodeficiency virus-monoinfected individuals, BMC Immunol 9, 28. Freeman, G. J., Wherry, E. J., Ahmed, R., and Sharpe, A. H. (2006) Reinvigorating exhausted HIV-specific T cells via PD-1-PD-1 ligand blockade, J Exp Med 203, 2223–2227. Day, C. L., Kaufmann, D. E., Kiepiela, P., Brown, J. A., Moodley, E. S., Reddy, S., Mackey, E. W., Miller, J. D., Leslie, A. J., DePierres, C., Mncube, Z., Duraiswamy, J., Zhu, B., Eichbaum, Q., Altfeld, M., Wherry, E. J., Coovadia, H. M., Goulder, P. J., Klenerman, P., Ahmed, R., Freeman, G. J., and Walker, B. D. (2006) PD-1 expression on HIV-specific T cells is associated with T-cell exhaustion and disease progression, Nature 443, 350–354. Landay, A. L., Clerici, M., Hashemi, F., Kessler, H., Berzofsky, J. A., and Shearer, G. M. (1996) In vitro restoration of T cell immune function in human immunodeficiency virus-positive persons: effects of interleukin (IL)-12 and anti-IL-10, J Infect Dis 173, 1085–1091. Velu, V., Titanji, K., Zhu, B., Husain, S., Pladevega, A., Lai, L., Vanderford, T. H., Chennareddi, L., Silvestri, G., Freeman, G. J., Ahmed, R., and Amara, R. R. (2009) Enhancing SIV-specific immunity in vivo by PD-1 blockade, Nature 458, 206–210.

39. Thepen, T., Van Rooijen, N., and Kraal, G. (1989) Alveolar macrophage elimination in vivo is associated with an increase in pulmonary immune response in mice, J Exp Med 170, 499–509. 40. Holt, P. G. (1978) Inhibitory activity of unstimulated alveolar macrophages on T-lymphocyte blastogenic response, Am Rev Respir Dis 118, 791–793. 41. Dillon, S., Agrawal, S., Banerjee, K., Letterio, J., Denning, T. L., Oswald-Richter, K., Kasprowicz, D. J., Kellar, K., Pare, J., van Dyke, T., Ziegler, S., Unutmaz, D., and Pulendran, B. (2006) Yeast zymosan, a stimulus for TLR2 and dectin-1, induces regulatory antigen-presenting cells and immunological tolerance, J Clin Invest 116, 916–928. 42. Holt, P. G., Oliver, J., Bilyk, N., McMenamin, C., McMenamin, P. G., Kraal, G., and Thepen, T. (1993) Downregulation of the antigen presenting cell function(s) of pulmonary dendritic cells in vivo by resident alveolar macrophages, J Exp Med 177, 397–407. 43. Bilyk, N., and Holt, P. G. (1995) Cytokine modulation of the immunosuppressive phenotype of pulmonary alveolar macrophage populations, Immunology 86, 231–237. 44. Strickland, D. H., Thepen, T., Kees, U. R., Kraal, G., and Holt, P. G. (1993) Regulation of T-cell function in lung tissue by pulmonary alveolar macrophages, Immunology 80, 266–272. 45. Steinman, R. M., Hawiger, D., and Nussenzweig, M. C. (2003) Tolerogenic dendritic cells, Annu Rev Immunol 21, 685–711. 46. Steinbrink, K., Wolfl, M., Jonuleit, H., Knop, J., and Enk, A. H. (1997) Induction of tolerance by IL-10-treated dendritic cells, J Immunol 159, 4772–4780. 47. Steinbrink, K., Graulich, E., Kubsch, S., Knop, J., and Enk, A. H. (2002) CD4(+) and CD8(+) anergic T cells induced by interleukin-10-treated human dendritic cells display antigen-specific suppressor activity, Blood 99, 2468–2476. 48. Kubsch, S., Graulich, E., Knop, J., and Steinbrink, K. (2003) Suppressor activity of anergic T cells induced by IL-10-treated human dendritic cells: association with IL-2and CTLA-4-dependent G1 arrest of the cell cycle regulated by p27Kip1, Eur J Immunol 33, 1988–1997. 49. Penna, G., Roncari, A., Amuchastegui, S., Daniel, K. C., Berti, E., Colonna, M., and Adorini, L. (2005) Expression of the inhibitory receptor ILT3 on dendritic cells is dispensable for induction of CD4+Foxp3+ regulatory T cells by 1,25-dihydroxyvitamin D3, Blood 106, 3490–3497.

Immunosuppressive Mechanisms During Viral Infectious Diseases 50. Netea, M. G., Sutmuller, R., Hermann, C., Van der Graaf, C. A., Van der Meer, J. W., van Krieken, J. H., Hartung, T., Adema, G., and Kullberg, B. J. (2004) Toll-like receptor 2 suppresses immunity against Candida albicans through induction of IL-10 and regulatory T cells, J Immunol 172, 3712–3718. 51. Sutmuller, R. P., den Brok, M. H., Kramer, M., Bennink, E. J., Toonen, L. W., Kullberg, B. J., Joosten, L. A., Akira, S., Netea, M. G., and Adema, G. J. (2006) Toll-like receptor 2 controls expansion and function of regulatory T cells, J Clin Invest 116, 485–494. 52. Penna, G., and Adorini, L. (2000) 1 Alpha,25dihydroxyvitamin D3 inhibits differentiation, maturation, activation, and survival of dendritic cells leading to impaired alloreactive T cell activation, J Immunol 164, 2405–2411. 53. Oh, S., and Eichelberger, M. C. (1999) Influenza virus neuraminidase alters allogeneic T cell proliferation, Virology 264, 427–435. 54. Oh, S., McCaffery, J. M., and Eichelberger, M. C. (2000) Dose-dependent changes in influenza virus-infected dendritic cells result in increased allogeneic T-cell proliferation at low, but not high, doses of virus, J Virol 74, 5460–5469. 55. Dhodapkar, M. V., and Steinman, R. M. (2002) Antigen-bearing immature dendritic cells induce peptide-specific CD8(+) regulatory T cells in vivo in humans, Blood 100, 174–177. 56. Dhodapkar, M. V., Steinman, R. M., Krasovsky, J., Munz, C., and Bhardwaj, N. (2001) Antigen-specific inhibition of effector T cell function in humans after injection of immature dendritic cells, J Exp Med 193, 233–238. 57. Sakaguchi, S., Wing, K., and Yamaguchi, T. (2009) Dynamics of peripheral tolerance and immune regulation mediated by Treg, Eur J Immunol 39, 2331–2336. 58. Hori, S., Nomura, T., and Sakaguchi, S. (2003) Control of regulatory T cell development by the transcription factor Foxp3, Science 299, 1057–1061. 59. Zhou, X., Bailey-Bucktrout, S. L., Jeker, L. T., Penaranda, C., Martinez-Llordella, M., Ashby, M., Nakayama, M., Rosenthal, W., and Bluestone, J. A. (2009) Instability of the transcription factor Foxp3 leads to the generation of pathogenic memory T cells in vivo, Nat Immunol 10, 1000–1007. 60. Toda, A., and Piccirillo, C. A. (2006) Development and function of naturally occurring CD4+CD25+ regulatory T cells, J Leukoc Biol 80, 458–470. 61. Ozdemir, C., Akdis, M., and Akdis, C. A. (2009) T regulatory cells and their counterparts: masters of immune regulation, Clin Exp Allergy 39, 626–639.

445

62. Horai, R., Asano, M., Sudo, K., Kanuka, H., Suzuki, M., Nishihara, M., Takahashi, M., and Iwakura, Y. (1998) Production of mice deficient in genes for interleukin (IL)-1alpha, IL-1beta, IL-1alpha/beta, and IL-1 receptor antagonist shows that IL-1beta is crucial in turpentineinduced fever development and glucocorticoid secretion, J Exp Med 187, 1463–1475. 63. Aandahl, E. M., Michaelsson, J., Moretto, W. J., Hecht, F. M., and Nixon, D. F. (2004) Human CD4+ CD25+ regulatory T cells control T-cell responses to human immunodeficiency virus and cytomegalovirus antigens, J Virol 78, 2454–2459. 64. Boettler, T., Spangenberg, H. C., NeumannHaefelin, C., Panther, E., Urbani, S., Ferrari, C., Blum, H. E., von Weizsacker, F., and Thimme, R. (2005) T cells with a CD4+CD25+ regulatory phenotype suppress in vitro proliferation of virus-specific CD8+ T cells during chronic hepatitis C virus infection, J Virol 79, 7860–7867. 65. Robertson, S. J., Messer, R. J., Carmody, A. B., and Hasenkrug, K. J. (2006) In vitro suppression of CD8+ T cell function by Friend virus-induced regulatory T cells, J Immunol 176, 3342–3349. 66. Suvas, S., Kumaraguru, U., Pack, C. D., Lee, S., and Rouse, B. T. (2003) CD4+CD25+ T cells regulate virus-specific primary and memory CD8+ T cell responses, J Exp Med 198, 889–901. 67. Truong, P., and McGavern, D. B. (2008) A novel virus carrier state to evaluate immunotherapeutic regimens: regulatory T cells modulate the pathogenicity of antiviral memory cells, J Immunol 181, 1161–1169. 68. Belkaid, Y. (2007) Regulatory T cells and infection: a dangerous necessity, Nat Rev Immunol 7, 875–888. 69. Belkaid, Y., and Rouse, B. T. (2005) Natural regulatory T cells in infectious disease, Nat Immunol 6, 353–360. 70. Robertson, S. J., and Hasenkrug, K. J. (2006) The role of virus-induced regulatory T cells in immunopathology,SpringerSeminImmunopathol 28, 51–62. 71. Iwashiro, M., Messer, R. J., Peterson, K. E., Stromnes, I. M., Sugie, T., and Hasenkrug, K. J. (2001) Immunosuppression by CD4+ regulatory T cells induced by chronic retroviral infection, Proc Natl Acad Sci U S A 98, 9226–9230. 72. MacDonald, A. J., Duffy, M., Brady, M. T., McKiernan, S., Hall, W., Hegarty, J., Curry, M., and Mills, K. H. (2002) CD4 T helper type 1 and regulatory T cells induced against the same epitopes on the core protein in hepatitis C virusinfected persons, J Infect Dis 185, 720–727.

446

Sarikonda and von Herrath

73. Luhn, K., Simmons, C. P., Moran, E., Dung, N. T., Chau, T. N., Quyen, N. T., Thao le, T. T., Van Ngoc, T., Dung, N. M., Wills, B., Farrar, J., McMichael, A. J., Dong, T., and Rowland-Jones, S. (2007) Increased frequencies of CD4+ CD25(high) regulatory T cells in acute dengue infection, J Exp Med 204, 979–985. 74. Shevach, E. M. (2009) Mechanisms of foxp3+ T regulatory cell-mediated suppression, Immunity 30, 636–645. 75. Tang, Q., and Bluestone, J. A. (2008) The Foxp3+ regulatory T cell: a jack of all trades, master of regulation, Nat Immunol 9, 239–244. 76. Vignali, D. A., Collison, L. W., and Workman, C. J. (2008) How regulatory T cells work, Nat Rev Immunol 8, 523–532. 77. Miyara, M., and Sakaguchi, S. (2007) Natural regulatory T cells: mechanisms of suppression, Trends Mol Med 13, 108–116. 78. Thornton, A. M., and Shevach, E. M. (1998) CD4+CD25+ immunoregulatory T cells suppress polyclonal T cell activation in vitro by inhibiting interleukin 2 production, J Exp Med 188, 287–296. 79. Wing, K., Onishi, Y., Prieto-Martin, P., Yamaguchi, T., Miyara, M., Fehervari, Z., Nomura, T., and Sakaguchi, S. (2008) CTLA-4 control over Foxp3+ regulatory T cell function, Science 322, 271–275. 80. Onishi, Y., Fehervari, Z., Yamaguchi, T., and Sakaguchi, S. (2008) Foxp3+ natural regulatory T cells preferentially form aggregates on dendritic cells in vitro and actively inhibit their maturation, Proc Natl Acad Sci U S A 105, 10113–10118. 81. de la Rosa, M., Rutz, S., Dorninger, H., and Scheffold, A. (2004) Interleukin-2 is essential for CD4+CD25+ regulatory T cell function, Eur J Immunol 34, 2480–2488. 82. Pandiyan, P., Zheng, L., Ishihara, S., Reed, J., and Lenardo, M. J. (2007) CD4+CD25+Foxp3+ regulatory T cells induce cytokine deprivationmediated apoptosis of effector CD4+ T cells, Nat Immunol 8, 1353–1362. 83. Filippi, C. M., Estes, E. A., Oldham, J. E., and von Herrath, M. G. (2009) Immunoregulatory mechanisms triggered by viral infections protect from type 1 diabetes in mice, J Clin Invest 119, 1515–1523. 84. Ha, S. J., Mueller, S. N., Wherry, E. J., Barber, D. L., Aubert, R. D., Sharpe, A. H., Freeman, G. J., and Ahmed, R. (2008) Enhancing therapeutic vaccination by blocking PD-1mediated inhibitory signals during chronic infection, J Exp Med 205, 543–555.

85. Petrovas, C., Casazza, J. P., Brenchley, J. M., Price, D. A., Gostick, E., Adams, W. C., Precopio, M. L., Schacker, T., Roederer, M., Douek, D. C., and Koup, R. A. (2006) PD-1 is a regulator of virus-specific CD8+ T cell survival in HIV infection, J Exp Med 203, 2281–2292. 86. Trautmann, L., Janbazian, L., Chomont, N., Said, E. A., Gimmig, S., Bessette, B., Boulassel, M. R., Delwart, E., Sepulveda, H., Balderas, R. S., Routy, J. P., Haddad, E. K., and Sekaly, R. P. (2006) Upregulation of PD-1 expression on HIV-specific CD8+ T cells leads to reversible immune dysfunction, Nat Med 12, 1198–1202. 87. Fife, B. T., and Bluestone, J. A. (2008) Control of peripheral T-cell tolerance and autoimmunity via the CTLA-4 and PD-1 pathways, Immunol Rev 224, 166–182. 88. Williams, M. A., Onami, T. M., Adams, A. B., Durham, M. M., Pearson, T. C., Ahmed, R., and Larsen, C. P. (2002) Cutting edge: persistent viral infection prevents tolerance induction and escapes immune control following CD28/CD40 blockade-based regimen, J Immunol 169, 5387–5391. 89. Rudd, C. E., Taylor, A., and Schneider, H. (2009) CD28 and CTLA-4 coreceptor expression and signal transduction, Immunol Rev 229, 12–26. 90. Read, S., Malmstrom, V., and Powrie, F. (2000) Cytotoxic T lymphocyte-associated antigen 4 plays an essential role in the function of CD25(+) CD4(+) regulatory cells that control intestinal inflammation, J Exp Med 192, 295–302. 91. Takahashi, T., Tagami, T., Yamazaki, S., Uede, T., Shimizu, J., Sakaguchi, N., Mak, T. W., and Sakaguchi, S. (2000) Immunologic selftolerance maintained by CD25(+)CD4(+) regulatory T cells constitutively expressing cytotoxic T lymphocyte-associated antigen 4, J Exp Med 192, 303–310. 92. Salomon, B., Lenschow, D. J., Rhee, L., Ashourian, N., Singh, B., Sharpe, A., and Bluestone, J. A. (2000) B7/CD28 costimulation is essential for the homeostasis of the CD4+ CD25+ immunoregulatory T cells that control autoimmune diabetes, Immunity 12, 431–440. 93. Simone, R., Piatti, G., and Saverino, D. (2009) The inhibitory co-receptors: a way to save from anergy the HIV-specific T cells, Curr HIV Res 7, 266–272. 94. Hryniewicz, A., Boasso, A., Edghill-Smith, Y., Vaccari, M., Fuchs, D., Venzon, D., Nacsa, J., Betts, M. R., Tsai, W. P., Heraud, J. M., Beer, B., Blanset, D., Chougnet, C., Lowy, I., Shearer, G. M., and Franchini, G. (2006) CTLA-4 blockade decreases TGF-beta, IDO, and viral RNA expression in tissues of SIVmac251infected macaques, Blood 108, 3834–3842.

Immunosuppressive Mechanisms During Viral Infectious Diseases 95. Kaufmann, D. E., Kavanagh, D. G., Pereyra, F., Zaunders, J. J., Mackey, E. W., Miura, T., Palmer, S., Brockman, M., Rathod, A., Piechocka-Trocha, A., Baker, B., Zhu, B., Le Gall, S., Waring, M. T., Ahern, R., Moss, K., Kelleher, A. D., Coffin, J. M., Freeman, G. J., Rosenberg, E. S., and Walker, B. D. (2007) Upregulation of CTLA-4 by HIVspecific CD4+ T cells correlates with disease progression and defines a reversible immune dysfunction, Nat Immunol 8, 1246–1254. 96. Krummel, M. F., and Allison, J. P. (1996) CTLA-4 engagement inhibits IL-2 accumulation and cell cycle progression upon activation of resting T cells, J Exp Med 183, 2533–2540. 97. Li, M. O., Wan, Y. Y., Sanjabi, S., Robertson, A. K., and Flavell, R. A. (2006) Transforming growth factor-beta regulation of immune responses, Annu Rev Immunol 24, 99–146. 98. Gorelik, L., and Flavell, R. A. (2002) Transforming growth factor-beta in T-cell biology, Nat Rev Immunol 2, 46–53. 99. Zhang, X., Giangreco, L., Broome, H. E., Dargan, C. M., and Swain, S. L. (1995) Control of CD4 effector fate: transforming growth factor beta 1 and interleukin 2 synergize to prevent apoptosis and promote effector expansion, J Exp Med 182, 699–709. 100. Genestier, L., Kasibhatla, S., Brunner, T., and Green, D. R. (1999) Transforming growth factor beta1 inhibits Fas ligand expression and subsequent activationinduced cell death in T cells via downregulation of c-Myc, J Exp Med 189, 231–239. 101. Cerwenka, A., Kovar, H., Majdic, O., and Holter, W. (1996) Fas- and activationinduced apoptosis are reduced in human T cells preactivated in the presence of TGFbeta 1, J Immunol 156, 459–464. 102. Chen, W., Jin, W., Hardegen, N., Lei, K. J., Li, L., Marinos, N., McGrady, G., and Wahl, S. M. (2003) Conversion of peripheral CD4+CD25-naive T cells to CD4+CD25+ regulatory T cells by TGF-beta induction of transcription factor Foxp3, J Exp Med 198, 1875–1886. 103. Filippi, C., Bresson, D., and von Herrath, M. (2005) Antigen-specific induction of regulatory

104.

105.

106.

107.

108.

109.

110.

111.

447

T cells for type 1 diabetes therapy, Int Rev Immunol 24, 341–360. Filippi, C. M., Juedes, A. E., Oldham, J. E., Ling, E., Togher, L., Peng, Y., Flavell, R. A., and von Herrath, M. G. (2008) TGF-{beta} suppresses the activation of CD8+ T cells when naive but promotes their survival and function once antigen-experienced: a two-faced impact on autoimmunity, Diabetes 57, 2684–2692. Piccirillo, C. A., Chang, Y., and Prud’homme, G. J. (1998) TGF-beta1 somatic gene therapy prevents autoimmune disease in nonobese diabetic mice, J Immunol 161, 3950–3956. Bettelli, E., Carrier, Y., Gao, W., Korn, T., Strom, T. B., Oukka, M., Weiner, H. L., and Kuchroo, V. K. (2006) Reciprocal developmental pathways for the generation of pathogenic effector TH17 and regulatory T cells, Nature 441, 235–238. Stockinger, B., and Veldhoen, M. (2007) Differentiation and function of Th17 T cells, Curr Opin Immunol 19, 281–286. Laurence, A., Tato, C. M., Davidson, T. S., Kanno, Y., Chen, Z., Yao, Z., Blank, R. B., Meylan, F., Siegel, R., Hennighausen, L., Shevach, E. M., and O’Shea, J. J. (2007) Interleukin-2 signaling via STAT5 constrains T helper 17 cell generation, Immunity 26, 371–381. de Waal Malefyt, R., Abrams, J., Bennett, B., Figdor, C. G., and de Vries, J. E. (1991) Interleukin 10(IL-10) inhibits cytokine synthesis by human monocytes: an autoregulatory role of IL-10 produced by monocytes, J Exp Med 174, 1209–1220. Taylor, A., Akdis, M., Joss, A., Akkoc, T., Wenig, R., Colonna, M., Daigle, I., Flory, E., Blaser, K., and Akdis, C. A. (2007) IL-10 inhibits CD28 and ICOS costimulations of T cells via src homology 2 domain-containing protein tyrosine phosphatase 1, J Allergy Clin Immunol 120, 76–83. Brooks, D. G., Lee, A. M., Elsaesser, H., McGavern, D. B., and Oldstone, M. B. (2008) IL-10 blockade facilitates DNA vaccine-induced T cell responses and enhances clearance of persistent virus infection, J Exp Med 205, 533–541.

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Chapter 28 Ocular Immune Privilege Sites Sharmila Masli and Jose L. Vega Abstract The eye is one of the immune privilege sites of the body that is consequently protected from the detrimental and potentially blinding influences of immunologic inflammation. Within the eye, the anterior chamber has been recognized for its immune privilege property for many years now; however, a similar property detectable in the subretinal space has only recently been appreciated. These ocular sites are not only equipped with specialized mechanisms that barricade local inflammatory responses, but also induce systemic regulatory immune response. Numerous studies have characterized molecular and cellular mechanisms involved in conferring both these sites with an immune privilege status. Pigmented epithelial cells lining the anterior chamber in the iris and ciliary body area as well as those in the retina are endowed with immunomodulatory properties that contribute to ocular immune privilege. These cells, via expression of either soluble factors or membrane molecules, inhibit inflammatory T cell activation and promote the generation of regulatory T cells. In the anterior chamber resident antigen-presenting cells, influenced by the various immunosuppressive factors present in the aqueous humor, capture ocular antigens and present them in the spleen to T cells in association with NKT cells and marginal zone B cells. Immunomodulatory microenvironment created by these cells helps generate regulatory T cells, capable of interrupting the induction as well as expression of inflammatory responses. Furthermore, neural regulation of both intraocular and systemic regulatory mechanisms also contributes to ocular immune privilege. Key words: Immune privilege, Anterior chamber, Antigen-presenting cells, Thrombospondin, Regulatory T cells, Ocular sympathetic innervations

1. Introduction The discovery of immune privilege came in 1948, when Sir Peter Medawar observed that, while allogeneic tissue grafts implanted under the skin resulted in their rapid elimination by the immune system, the implantation of similar grafts into either the eye or the brain resulted in their prolonged survival [1]. Later on, through the injection of tumor cells into various ocular compartments, it was shown that the anterior chamber, the subretinal space, and the vitreous compartment are all immune-privileged sites [2, 3]. Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_28, © Springer Science+Business Media, LLC 2011

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This chapter will address the main properties and mechanisms of ocular immune-privileged sites and tissues as depicted in Fig. 1, and will make an emphasis on the role of three key aspects of this important biological phenomenon: regulatory T cells (Tregs), antigen-presenting cells (APCs) , and the local nerve supply.

Fig. 1. Ocular immune privileged sites and mechanisms of immune regulation. (a) Intraocular immune regulation is achieved by pigmented epithelial (PE) cells of the iris, ciliary body and retina during their interactions with immune cells. While iris and ciliary body PE cells utilize molecules such as CD86, membrane bound TGF-b and TSP-1, retinal PE cells express FasL, somatostatin, TSP-1, and PEDF to induce Treg. Peripheral immune regulation is attributed to local APCs exposed to TGF-b2, among many other immunosuppressive factors in the ocular environment. These APCs upon capturing ocular antigen migrate via blood to the spleen where they present antigen to CD4+ and CD8+ T cells. By producing the chemokine MIP-2, TGF-b-exposed APCs recruit NKT cells and together with marginal zone B cells create an immunomodulatory microenvironment comprised of TGF-b and IL-10. Antigen presentation in this microenvironment results in the induction of regulatory T cells. Neural pathways also are involved in immune regulation at ocular immune privilege sites. Sensory (trigeminal ganglion) and sympathetic (superior cervical ganglion) nerves of the cornea, iris, and ciliary body regulate the microenvironment in the anterior chamber. (b) TGF-b-exposed ocular APCs utilize unique molecular mechanisms that promote the induction of Tregs. These include increased expression of molecules like TNF-a, IFN-b, TNF-R2, IkBa, and TSP-1. While most of these molecules contribute to the inhibition of IL-12, TSP-1 also facilitates the activation of latent TGF-b produced by APCs and tethers it to their surface providing TGF-b-rich microenvironment during antigen presentation.

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Fig. 1 (Continued)

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The anterior chamber (AC) of the eye is filled with aqueous humor, a clear fluid, which is saturated with multiple immunosuppressive substances that include TGF-b2, arguably the most important mediator of immune privilege, as well as neuropeptides such as vasointestinal peptide (VIP), alpha-melanocyte stimulating hormone (a-MSH), and somatostatin, among others [4]. The immunoactive substances found in aqueous humor mirror the very molecules that are synthesized and released, either actively or passively, by the tissue cells that line the anterior chamber, including the corneal endothelium, the ciliary body, the iris pigment epithelium, and the trabecular meshwork. Aqueous humor from normal eyes exhibits a wide range of immunosuppressive abilities [5], which include the suppression of T cell proliferation and the production of proinflammatory cytokines, and in some cases, the conversion of activated T cells into Tregs. These also include the attenuation of macrophage activation and the release of IFN-g and other proinflammatory mediators, and the conversion of resident F4/80+ APCs into highly effective tolerogenic APCs. The posterior aspect of the anterior chamber is lined by the ciliary body pigment epithelium (CBPE) and the iris pigment epithelium (IPE) which express membrane-bound TGF-b and other molecules, and allow them to carry out the regulation of inflammatory responses in a contact-dependent fashion [6]. The posterior eye pole is lined in its entirety by the retinal pigment epithelium (RPE), a single layer of cells with very potent immunosuppressive abilities that support immune privilege in the subretinal space.

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2. Intraocular Immune Regulation

3. Peripheral Immune Regulation

Ocular parenchymal cells play a significant role in inducing regulatory effectors that can avoid local inflammation thereby protecting the delicate anatomy within the eye. The intraocular pigment epithelial cells from the iris, ciliary body, and retina are equipped with mechanisms to convert naïve T cells into Tregs. Early work by Yoshida and coworkers showed that cultured mouse I/CBPE cells suppressed T-cell proliferation, IFN-g, IL-2, IL-4, and IL-10 secretion, through a contact-dependent mechanism [7], and induced a population of TGF-b-producing suppressor T cells [8]. Subsequent experiments by Sugita and colleagues demonstrated that IPE cells express high levels of surface TGF-b2, and B7-2, which synergize to induce the upregulation of both TGF-b1 and B7-2 on activated CD8+ T cells. These CD8+ T cells can in turn go on to suppress the activation of other, freshly isolated naïve T cells in a TGF-b-dependent manner [6]. Like IPE cells, cultured RPE cells also demonstrate a constitutive expression of TGF-b2 [9], and other molecules such as program cell death ligand 1 (PD-L1), CTLA-2a, and FasL through which they regulate T cell-mediated immune responses [10–12]. More recent work has demonstrated that CBPE and RPE cells also induce Foxp3+ Tregs in a TGF-b-dependent fashion [6, 11]. While cultured RPE cells have been shown to induce regulatory function through soluble TGF-b, more recently we found that membrane-bound TGF-b on RPE cells also works together with T cell-derived TGF-b1 to mediate the conversion of naïve CD4+ T cells into Foxp3-expressing Tregs. In fact, RPE-derived soluble TGF-b does not participate in mediating this effect. This finding is physiologically relevant, as it is known that the untargeted activation of soluble TGF-b in the eye can fuel fibrosis in diseases such as proliferative vitreoretinopathy, diabetic retinopathy, and open angle glaucoma (reviewed in ref. 13). Therefore, it is likely that all pigmented epithelial cells induce Treg function through membrane-bound TGF-b, which decreases the possibility of undesirable intraocular fibrosis.

Aside from suppressing inflammatory responses through a multiplicity of intraocular mechanisms, the eye is also capable of orchestrating systemic regulatory immune responses against intraocular antigens. For example, animals injected with allogeneic tumor cells into the AC fail to develop delayed type hypersensitivity

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(DTH) responses against tumor cell alloantigens [2]. Although these animals do develop antigen-specific immune responses, which consist of noncomplement fixing antibodies and CD8+ T cells [14, 15], their proinflammatory effect is thwarted by Tregs. The systemic induction of a tolerogenic response against eyederived antigens is characterized by the induction of two kinds of Tregs. The first set has been coined “afferent,” is made up of CD4+ cells, and is known to prevent the activation and differentiation of antigen-specific Th1 effector cells [16, 17]. The second or “efferent” set of Tregs is made up of CD8+ T cells, and has been shown to block already developed efferent DTH responses mediated by Th1 cells. Recently, it was also demonstrated that antigens introduced in the AC of an eye result in an expansion of the CD4+CD25+Foxp3+ Treg population in the recipient spleen [18]. These results were further supported by the findings of Zhang and coworkers who reported a similar increase in CD4+CD25+Foxp3+ Treg population in recipients when antigen was introduced via i.v. infusion of antigen-pulsed TGF-b-treated APCs that resemble eye-derived APCs [19]. Thus, Foxp3+ Tregs are implicated in the suppression of the DTH response against ocular antigens. These regulatory mechanisms begin with the influence of an intraocular immunosuppressive microenvironment that changes the phenotype and homing abilities of local F4/80+ APCs, which upon capture of intracameral antigens, and under the influence of locally produced TGF-b, migrate preferentially to the marginal zone of the spleen where the secretion of the chemokine MIP-2 results in the recruitment of NKT cells [20, 21]. In conjunction with marginal zone B cells, these ocular APCs activate regulatory effectors. Importantly, an intact spleen represents an absolute requirement for the development of this regulatory response as animals that are splenectomized before, and up to 5 days after intracameral antigen injections, fail to suppress subsequent DTH responses against eye-derived antigens [22]. Although eye-derived APCs carrying ocular antigens are known to reach the spleen via the blood, which mimics i.v. injection of high-dose antigens, it is important to emphasize that the systemic immune response generated against eye-derived antigens is radically different from the long-lasting antigen specific unresponsiveness induced by the i.v. injections [23]. The latter has been shown to work through a combination of apoptosis of antigen-specific thymic and peripheral T cell clones, and through T cell receptor desensitization [24]. Furthermore, tolerance induced by i.v. antigen injections cannot suppress immune responses against antigens to which the immune system has been previously primed [16]. Thus, the DTH response in mice previously injected with subcutaneous ovalbumin (OVA) can be nearly completely

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suppressed by subsequent AC, but not i.v, antigen injections, indicating that the mechanisms of immune tolerance against eyederived antigens are uniquely potent [16]. 3.1. Immune Regulation by Eye-Derived APCs

Bone marrow-derived cells of the iris and ciliary body exposed to factors in the aqueous humor exhibit immunoregulatory properties in vivo as well as in vitro [25]. These cells express predominantly F4/80 and CD11b/Mac-1 markers associated with macrophages and dendritic cells are putative eye-derived APCs that present ocular antigens and give rise to peripheral regulatory immune response (immunologic tolerance). This ability of ocular APCs to induce tolerance is attributed to their exposure to ocular factors, predominantly TGF-b [26]. In fact, conventional APCs from nonocular sites, when exposed to eye-derived factors either in vivo by AC injection or in vitro by culturing with supernatants derived from I/CB PE cultures or TGF-b, exhibit functional properties similar to ocular APCs. Conventional APCs, upon their exposure to TGF-b, are known to increase their expression of the immunomodulatory cytokines, TGF-b and IL-10 [27, 28]. These cytokines allow APCs to present antigens to T cells in a unique cytokine milieu, which results in the induction of Tregs. Furthermore, TGF-b-exposed APCs fail to express molecules known to support inflammatory Th1 effectors such as IL-12 and CD40 [29]. Effectors activated by these APCs, either in vivo or in vitro, produce TGF-b while their ability to secrete IFN-g and IL-2 is diminished [30, 31]. This phenotype of effectors resembles that recently described for CD4+CD25+Foxp3+ Tregs. Therefore, it is quite likely that ocular APCs activate T cells to generate what are now described as inducible Tregs (iTregs). More recently, analysis of differentially expressed genes in TGF-b-exposed APCs as compared to untreated APCs revealed new mechanisms utilized by these APCs to promote immunoregulatory responses (depicted in Fig. 1b). In response to TGF-b exposure, APCs increase the expression of molecules like IFN-b, TNF-a, TNF-R2, Thrombospondin-1 (TSP-1), and IkBa which were found to be critical for their ability to induce regulatory effectors [32–35]. Increased expression of a typically proinflammatory cytokine such as TNF-a by APCs that support a regulatory response, although paradoxical, was found to be essential as TNF-a-deficient APCs failed to acquire the functional phenotype of ocular APCs upon their exposure to TGF-b. While the proinflammatory effects of TNF-a are associated with signaling primarily through TNF-R1, TGF-b exposed APCs negate this effect by increasing their expression of TNF-R2, thereby altering signaling in a way that supports the anti-inflammatory functional phenotype of ocular APCs [34]. Moreover, an increased expression of IFN-b, TNF-a, TSP-1, and IkBa contributes to impaired expression of IL-12 in TGF-b-exposed APCs. Increased synthesis

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of IkBa by TGF-b-exposed APCs results in increased nuclear levels of this inhibitor of the proinflammatory nuclear factor NFkB and the subsequent dissociation of NFkB, which effectively diminishing the transcription of IL-12. Thrombospondin-1, a matricellular protein capable of binding various cell surface receptors such as CD36, CD47, and integrins, emerged as a critical molecule not only for the immunoregulatory function of ocular APCs, but also for the maintenance of immune privilege in the eye [33, 36]. A multifunctional protein like TSP-1, known to deliver varied biological effects on target cells depending on the receptors ligated, contributes to IL-12 inhibition via CD47 on TGF-b-exposed APCs and facilitates the activation of latent TGF-b by binding CD36. Thrombospondin tethers latent TGF-b to the cell surface allowing the release of active TGF-b in a microenvironment in which T cells are activated. The influence of TGF-b on the development of Tregs is well appreciated and thus TSP-1 happens to be a key player in orchestrating the induction of Tregs by TGF-b-exposed APCs. Ocular APCs, with their surface bound TSP-1, create a satellite microenvironment in the spleen during antigen presentation that mimics the TGF-b-rich ocular environment. Furthermore, a potential CD36-TSP-CD47 trimolecular interaction is likely to maintain an effective APC-T cell contact increasing the efficiency of signals delivered during antigen presentation. Some reports suggest that signals mediated by CD47 ligation on T cells interrupt TCR-mediated signals [37]. In human T cells, CD47 ligation has been reported to result in the induction and expansion of Foxp3+ Tregs [38, 39]. In fact, CD47 ligation during Th1 polarization was reported to downregulate the secretion of the inflammatory cytokines IFN-g and IL-2 [40]. These reports clearly suggest that the successful induction of an immunoregulatory response and the prevention of an inflammatory response against ocular antigens is due to unique molecular mechanisms employed by eye-derived APCs involving TSP-1 interactions. 3.2. Neural Regulation of Ocular Immune Privilege Sites

Neural regulation of ocular immune privilege has also become evident from several investigations that addressed the involvement of sensory, sympathetic as well as parasympathetic pathways of the nervous system. An important early clinical observation suggested a role for corneal sensory fibers. When Streilein and colleagues noticed that eyes that received a corneal graft failed to suppress the DTH against ocular antigens for the first 8 weeks after the graft, but regained the ability to do so after 12 weeks. Because the return of the DTH suppression coincided with the return of sensory innervation to the newly grafted cornea, Streilein and colleagues hypothesized that sensory nerves supplying the cornea might play an important role in the eye-derived regulatory immune response. Subsequent experiments demonstrated that

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circumferential partial-thickness incisions that transected the sensory innervation of the cornea, slightly medial to the limbus, recreated the loss of DTH suppression previously seen in grafted eyes [41], making a strong argument that the sensory innervation of the cornea plays an important role in establishing the immunosuppressive environment of the anterior chamber. More recently, we investigated the role of intraocular sympathetic nerves in ocular immune privilege [42]. We were motivated to do this study by the multiple case reports found in the medical literature of patients with chronic anterior uveitis (Fuch’s uveitis) who concurrently are found to suffer from Horner’s syndrome (i.e., a defect in the sympathetic innervation of head and neck structures). We found that while surgical denervation of ocular sympathetic fibers does not cause inflammation by itself, it does disrupt ocular immune privilege, thus setting up a microenvironment that allows for inflammation to occur. Importantly, this breakdown of ocular immune privilege is temporally associated with a decrease in the production of TGF-b in the aqueous humor, suggesting that the sympathetic system might partially regulate the production of this cytokine by structures in the anterior chamber. Thus, it is possible, that the association between Fuch’s uveitis and Horner’s syndrome is explained by a breakdown in ocular immune privilege. Further research in humans will be needed to confirm or negate this hypothesis. Another major component of the nervous system present inside the orbit is the parasympathetic innervation of the iris and ciliary body, which is provided by the oculomotor nerve. To our knowledge, no studies have been performed to elucidate the role of this nerve in ocular immune privilege. Nonetheless, various neuropeptides can be found inside the various types of nerve terminals in the orbit and in the aqueous humor such as VIP, a-MSH, and somatostatin. These neuropeptides can suppress lymphocytic proliferation and IFN-g production in vitro [43, 44], and appear to participate in the generation of regulatory T cells [45]. In addition, calcitonin gene-related peptide (CGRP), which is also found in the aqueous humor, has been shown to suppress nitric oxide production by activated macrophages [46]. Thus, in the aggregate, strong evidence exists that sensory, sympathetic as well as parasympathetic pathways in the eye play an important role in maintaining immune privilege. However, the specific mechanisms by which the associated nerves accomplish their important effects are not known. Also, role of intraocular neurotransmitters in the direct regulation of T cells and APCs inside the eye, and in the production of TGF-b by the tissues that make up the anterior chamber still remains unknown. Elucidation of these potential effects will certainly advance our understanding of the neuroimmune connection in the eye.

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References 1. Medawar, P. B. (1948) Immunity to homologous grafted skin; the fate of skin homografts transplanted to the brain, to subcutaneous tissue, and to the anterior chamber of the eye. Br J Exp Pathol 29, 58–69 2. Niederkorn, J., Streilein, J. W., and Shadduck, J. A. (1981) Deviant immune responses to allogeneic tumors injected intracamerally and subcutaneously in mice. Invest Ophthalmol Vis Sci 20, 355–363 3. Jiang, L. Q., Jorquera, M., and Streilein, J. W. (1993) Subretinal space and vitreous cavity as immunologically privileged sites for retinal allografts. Invest Ophthalmol Vis Sci 34, 3347–3354 4. Streilein, J. W., Masli, S., Takeuchi, M., and Kezuka, T. (2002) The eye’s view of antigen presentation. Hum Immunol 63, 435–443 5. Taylor, A. W. (2007) Ocular immunosuppressive microenvironment. Chem Immunol Allergy 92, 71–85 6. Sugita, S., Futagami, Y., Smith, S. B., Naggar, H., and Mochizuki, M. (2006) Retinal and ciliary body pigment epithelium suppress activation of T lymphocytes via transforming growth factor beta. Exp Eye Res 83, 1459–1471 7. Yoshida, M., Takeuchi, M., and Streilein, J. W. (2000) Participation of pigment epithelium of iris and ciliary body in ocular immune privilege. 1. Inhibition of T-cell activation in vitro by direct cell-to-cell contact. Invest Ophthalmol Vis Sci 41, 811–821 8. Yoshida, M., Kezuka, T., and Streilein, J. W. (2000) Participation of pigment epithelium of iris and ciliary body in ocular immune privilege. 2. Generation of TGF-beta-producing regulatory T cells. Invest Ophthalmol Vis Sci 41, 3862–3870 9. Pfeffer, B. A., Flanders, K. C., Guerin, C. J., Danielpour, D., and Anderson, D. H. (1994) Transforming growth factor beta 2 is the predominant isoform in the neural retina, retinal pigment epithelium-choroid and vitreous of the monkey eye. Exp Eye Res 59, 323–333 10. Usui, Y., Okunuki, Y., Hattori, T., Kezuka, T., Keino, H., Ebihara, N., Sugita, S., Usui, M., Goto, H., and Takeuchi, M. (2008) Functional expression of B7H1 on retinal pigment epithelial cells. Exp Eye Res 86, 52–59 11. Sugita, S., Horie, S., Nakamura, O., Futagami, Y., Takase, H., Keino, H., Aburatani, H., Katunuma, N., Ishidoh, K., Yamamoto, Y., and Mochizuki, M. (2008) Retinal pigment epithelium-derived CTLA-2alpha induces TGFbeta-producing T regulatory cells. J Immunol 181, 7525–7536

12. Wenkel, H., and Streilein, J. W. (2000) Evidence that retinal pigment epithelium functions as an immune-privileged tissue. Invest Ophthalmol Vis Sci 41, 3467–3473 13. Saika, S. (2006) TGFbeta pathobiology in the eye. Lab Invest 86, 106–115 14. Wilbanks, G. A., and Streilein, J. W. (1990) Distinctive humoral immune responses following anterior chamber and intravenous administration of soluble antigen. Evidence for active suppression of IgG2-secreting B lymphocytes. Immunology 71, 566–572 15. Ksander, B. R., and Streilein, J. W. (1989) Analysis of cytotoxic T cell responses to intracameral allogeneic tumors. Invest Ophthalmol Vis Sci 30, 323–329 16. Kosiewicz, M. M., Okamoto, S., Miki, S., Ksander, B. R., Shimizu, T., and Streilein, J. W. (1994) Imposing deviant immunity on the presensitized state. J Immunol 153, 2962–2973 17. Wilbanks, G. A., and Streilein, J. W. (1990) Characterization of suppressor cells in anterior chamber-associated immune deviation (ACAID) induced by soluble antigen. Evidence of two functionally and phenotypically distinct T-suppressor cell populations. Immunology 71, 383–389 18. Saban, D. R., Cornelius, J., Masli, S., Schwartzkopff, J., Doyle, M., Chauhan, S. K., Peck, A. B., and Grant, M. B. (2008) The role of ACAID and CD4+CD25+ FOXP3+ regulatory T cells on CTL function against MHC alloantigens. Mol Vis 14, 2435–2442 19. Zhang, H., Yang, P., Zhou, H., Meng, Q., and Huang, X. (2008) Involvement of Foxp3expressing CD4+ CD25+ regulatory T cells in the development of tolerance induced by transforming growth factor-beta2-treated antigenpresenting cells. Immunology 124, 304–314 20. Wilbanks, G. A., and Streilein, J. W. (1992) Macrophages capable of inducing anterior chamber associated immune deviation demonstrate spleen-seeking migratory properties. Reg Immunol 4, 130–137 21. Faunce, D. E., Sonoda, K. H., and SteinStreilein, J. (2001) MIP-2 recruits NKT cells to the spleen during tolerance induction. J Immunol 166, 313–321 22. Kaplan, H. J., and Streilein, J. W. (1974) Do immunologically privileged sites require a functioning spleen? Nature 251, 553–554 23. Weigle, W. O. (1973) Immunological unresponsiveness. Adv Immunol 16, 61–122 24. Liblau, R. S., Tisch, R., Shokat, K., Yang, X., Dumont, N., Goodnow, C. C., and McDevitt, H. O. (1996) Intravenous injection of soluble

458

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

Masli and Vega antigen induces thymic and peripheral T-cells apoptosis. Proc Natl Acad Sci USA 93, 3031–3036 Williamson, J. S., Bradley, D., and Streilein, J. W. (1989) Immunoregulatory properties of bone marrow-derived cells in the iris and ciliary body. Immunology 67, 96–102 Wilbanks, G. A., Mammolenti, M., and Streilein, J. W. (1992) Studies on the induction of anterior chamber-associated immune deviation (ACAID). III. Induction of ACAID depends upon intraocular transforming growth factor-beta. Eur J Immunol 22, 165–173 Takeuchi, M., Kosiewicz, M. M., Alard, P., and Streilein, J. W. (1997) On the mechanisms by which transforming growth factorbeta 2 alters antigen-presenting abilities of macrophages on T cell activation. Eur J Immunol 27, 1648–1656 D’Orazio, T. J., and Niederkorn, J. Y. (1998) A novel role for TGF-beta and IL-10 in the induction of immune privilege. J Immunol 160, 2089–2098 Takeuchi, M., Alard, P., and Streilein, J. W. (1998) TGF-beta promotes immune deviation by altering accessory signals of antigenpresenting cells. J Immunol 160, 1589–1597 Kosiewicz, M. M., Alard, P., and Streilein, J. W. (1998) Alterations in cytokine production following intraocular injection of soluble protein antigen: impairment in IFN-gamma and induction of TGF-beta and IL-4 production. J Immunol 161, 5382–5390 Kezuka, T., and Streilein, J. W. (2000) Analysis of in vivo regulatory properties of T cells activated in vitro by TGFbeta2-treated antigen presenting cells. Invest Ophthalmol Vis Sci 41, 1410–1421 Hecker, K. H., Niizeki, H., and Streilein, J. W. (1999) Distinct roles for transforming growth factor-beta2 and tumour necrosis factor-alpha in immune deviation elicited by haptenderivatized antigen-presenting cells. Immunology 96, 372–380 Masli, S., Turpie, B., and Streilein, J. W. (2006) Thrombospondin orchestrates the tolerance-promoting properties of TGFbetatreated antigen-presenting cells. Int Immunol 18, 689–699 Masli, S., and Turpie, B. (2009) Antiinflammatory effects of tumour necrosis factor (TNF)-alpha are mediated via TNF-R2 (p75) in tolerogenic transforming growth factor-beta-treated antigen-presenting cells. Immunology 127, 62–72 Ghafoori, P., Yoshimura, T., Turpie, B., and Masli, S. (2009) Increased IkappaB alpha expression is essential for the tolerogenic

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

property of TGF-beta-exposed APCs. FASEB J 23, 2226–2234 Zamiri, P., Masli, S., Kitaichi, N., Taylor, A. W., and Streilein, J. W. (2005) Thrombospondin plays a vital role in the immune privilege of the eye. Invest Ophthalmol Vis Sci 46, 908–919 Li, Z., He, L., Wilson, K., and Roberts, D. (2001) Thrombospondin-1 inhibits TCRmediated T lymphocyte early activation. J Immunol 166, 2427–2436 Van, V. Q., Darwiche, J., Raymond, M., Lesage, S., Bouguermouh, S., Rubio, M., and Sarfati, M. (2008) Cutting edge: CD47 controls the in vivo proliferation and homeostasis of peripheral CD4+ CD25+ Foxp3+ regulatory T cells that express CD103. J Immunol 181, 5204–5208 Grimbert, P., Bouguermouh, S., Baba, N., Nakajima, T., Allakhverdi, Z., Braun, D., Saito, H., Rubio, M., Delespesse, G., and Sarfati, M. (2006) Thrombospondin/CD47 interaction: a pathway to generate regulatory T cells from human CD4+ CD25- T cells in response to inflammation. J Immunol 177, 3534–3541 Avice, M. N., Rubio, M., Sergerie, M., Delespesse, G., and Sarfati, M. (2000) CD47 ligation selectively inhibits the development of human naive T cells into Th1 effectors. J Immunol 165, 4624–4631 Streilein, J. W., Bradley, D., Sano, Y., and Sonoda, Y. (1996) Immunosuppressive properties of tissues obtained from eyes with experimentally manipulated corneas. Invest Ophthalmol Vis Sci 37, 413–424 Vega, J. L., Keino, H., and Masli, S. (2009) Surgical denervation of ocular sympathetic afferents decreases local transforming growth factor-beta and abolishes immune privilege. Am J Pathol 175, 1218–1225 Taylor, A. W., and Yee, D. G. (2003) Somatostatin is an immunosuppressive factor in aqueous humor. Invest Ophthalmol Vis Sci 44, 2644–2649 Taylor, A. W., Streilein, J. W., and Cousins, S. W. (1994) Immunoreactive vasoactive intestinal peptide contributes to the immunosuppressive activity of normal aqueous humor. J Immunol 153, 1080–1086 Taylor, A., and Namba, K. (2001) In vitro induction of CD25+ CD4+ regulatory T cells by the neuropeptide alpha-melanocyte stimulating hormone (alpha-MSH). Immunol Cell Biol 79, 358–367 Taylor, A. W., Yee, D. G., and Streilein, J. W. (1998) Suppression of nitric oxide generated by inflammatory macrophages by calcitonin gene-related peptide in aqueous humor. Invest Ophthalmol Vis Sci 39, 1372–1378

Chapter 29 Immunoprivileged Sites: The Testis Monika Fijak, Sudhanshu Bhushan, and Andreas Meinhardt Abstract The testis is an immunological privileged tissue as evidenced by its ability to support grafts with minimal rejection. Immune privilege is essential for the tolerance of neo-antigens from developing germ cells that appear after the constitution of self-tolerance, but imposes the paradoxical task of also providing efficient protection against pathogens and tumor cells. It is becoming increasingly clear that immune privilege cannot be attributed to a single factor such as the sequestration of neo-antigens from the immune system behind the blood–testis barrier, but is based on a complex multifaceted interplay between cells and factors that are essential for the reproductive function of the testis and the testicular immune system. This review summarizes the evidence that has accumulated regarding the role of Sertoli cells, androgens, and selected population of leukocytes in the maintenance of immune privilege and its perturbation in testicular inflammatory sub- and infertility. Key words: Sertoli cells, Testosterone, Regulatory T lymphocytes, Dendritic cells, Cytokines

1. Testicular Cytoarchitecture and Blood–Testis Barrier

The adult mammalian testis is a complex organ that consists of two structurally distinct but functionally connected compartments, i.e., the tubular seminiferous epithelium and the interstitial compartment interspersed between the tubules. In the seminiferous epithelium, germ cells divide and differentiate to form spermatozoa in a process called spermatogenesis. Columnar Sertoli cells encompass and nourish the germ cells forming the scaffolding structure of the seminiferous epithelium. The contractile myoid peritubular cells form single (rodents) or multiple (human) flat layers surrounding the seminiferous tubules. The interstitial compartment contains predominantly androgenproducing Leydig cells, a considerable number of leukocytes, fibrocytes, and the vasculature including lymph vessels draining

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0_29, © Springer Science+Business Media, LLC 2011

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to renal and paraaortal lymph nodes (1–3). Testosterone secreted by the Leydig cells is found in high concentrations in the interstitial fluid. Androgens act in a paracrine fashion directly on peritubular and Sertoli cells, but not on germ cells, providing the most striking functional link between both compartments. This stimulation of Sertoli and peritubular cells is essential to complete spermatogenesis (4, 5). The seminiferous epithelium is not vascularized and intercellular transport into the seminiferous epithelium and is restricted by a specialized junctional complex, the blood–testis barrier (BTB). The BTB is physically formed by the borders of adjacent Sertoli cells and separates the seminiferous epithelium into two (sub)compartments. Spermatogonia and early primary spermatocytes are found in the basal section and are segregated from the more advanced meiotic and all post-meiotic germ cells located in the adluminal compartment. The BTB is a dynamic structure which while allowing the passage of developing germ cells, continuously maintains its barrier function. This process requires the disassembly of junctional complexes above translocating preleptotene spermatocytes with the parallel reassembly underneath these cells. Besides limiting the intercellular diffusion of molecules, the BTB also serves as an immunologic barrier which blocks the entry of leukocytes into the seminiferous epithelium. Why the BTB is so efficient in this function is unclear as most proteins constituting this barrier (e.g., the junctional adhesion molecules (JAM) and nectins) are also found in other epithelial boundaries such as the endothelial barrier which can easily be passed by migrating leukocytes (for review, see ref. 6). Immune privilege defines sites in the body which tolerate experimentally or naturally introduced antigens by prompting an inflammatory response. Such sites include the placenta, the anterior chamber of the eye, the brain, and the male gonad. It has long been assumed that the BTB constitutes the principle cause of immune-privileged status of the testis. This is due to its ability to sequester various neo-autoantigens expressed by the meiotic and haploid germ cells that appear for a long time after the establishment of self-tolerance, namely during puberty. However, the protection of autoantigenic cells by the BTB from the immune system has now been shown to represent only one of many contributors that establish the special immune environment of the testis (7, 8). Convincing evidence for the presence of alternative sources to the BTB has been found from mouse allografts which when transplanted into the interstitial area of the testis, a region where no physical separation from the immune system and efficient lymphatic drainage exists, survived considerably longer than grafts transplanted into non-immune-privileged sites (1, 9). Similarly, germ cells that leaked into the interstitium during trauma experiments were not attacked by the testicular immune

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system (10), although very recent data suggest that injury-induced testicular autoimmunity is dependent on the mouse strain (11). Moreover, with the onset of puberty, neo-autoantigens have been found in the basal compartment of the seminiferous epithelium, on early primary spermatocytes before the cells traverse the BTB (12, 13). Paradoxically, the breakdown of the BTB with its subsequent leukocytic infiltration of the tubules can be found in human biopsy specimens from cases of idiopathic infertility and in models of testicular inflammation (14–16). Interestingly, gene array studies found that inflammation-related genes such as cytokines also increased in human spermatogenic failure comparable to the signature known from inflammation-related pathologies (17). Besides immune cell activation and migration, cytokines also regulate junctional protein expression. Mechanistically, elevated levels of IL-1a, TNFa, NO, and transforming growth factor (TGF)-b, which are found in systemic and local testicular inflammation (82, 83), have been shown to perturb the assembly of the tight junctions in cultured Sertoli cells by downregulating the expression of barrier protein components, altering proteinase/ antiprotease activity adjacent to the junctions, and modulating the cytoskeleton (84, 85). Studies have also shown that the MAP kinases Jnk, p38, and Erk1/2 are critical intermediates in the regulation and maintenance of the BTB (86). On balance, the data emphasize the view that a delicate cytokine balance is important for BTB function.

2. Sentinel Role of the Sertoli Cells Besides their nutritional role in spermatogenesis and in BTB formation, there is now increasing evidence that suggests an important immunoregulatory function for Sertoli cells. Characterized by their ability to suppress immune function and thus protect the seminiferous epithelium from harmful immune reactions, Sertoli cells also produce a number of immunoregulatory mediators among which, TGF-b is thought to play a major role in immunosuppression. This is best demonstrated in co-transplantation studies where TGF-b from Sertoli cells favors Th2 over Th1 responses and be doing so protects pancreatic islets from allo- and autoimmune graft destruction in rodents (18). In vitro lymphocytes cultured with Sertoli cell secretions strongly reduce IL-2 release and the Sertoli cell product serpina3n effectively inhibits granzyme B-mediated apoptosis (19, 20). On the other hand, lymphocyte apoptosis can be induced by rat Sertoli cells which express FasL and TGF-b1, indicating a potential mechanism by which lymphocyte access to the seminiferous epithelium can be prevented (21). The Fas/FasL system is established as an inducer of extrinsic cell

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death in cell-mediated cytotoxicity, peripheral immune regulation, immune privilege, and “counterattack” of malignant tumor cells against the host immune system. Fas (APO-1/CD45) is a type I membrane bound protein, while its ligand FasL is a type II transmembrane protein that can be converted into soluble form and subsequently released by metalloproteinases (22). In the adult testis, Fas is mainly expressed in germ cells and in some Sertoli cells (23), and has been considered to be one of the most important mechanisms in immunological homeostasis of the testis. Supporting evidence for this proposition can be found in the increased expression of Fas/FasL in inflamed and degenerated testis. Similar expression can be found in testes with Sertoli cellonly syndrome and maturation arrest (24, 25), linking Fas/FasL with apoptotic elimination of germ cells (26). It has also emerged that communication between germ cells and Sertoli cells involves mechanisms that overlap with the inflammatory processes which are linked to recognition of infection. Unquestionably, these networks play a critical role in spermatogenic disruption caused by inflammation and infection (27).

3. Immune Privilege: More Than Immaturity of Dendritic Cells

Dendritic cells (DC) are a bone marrow-derived highly specialized heterogeneous population of antigen-presenting cells (APC) that initiate and regulate immune responses. DC not only initiate immunity by the activation of naïve B and T cells, but also tolerize T cells to antigens, thereby minimizing autoaggressive immune responses (28). DC function is dependent on their maturation stage. Within tissues, immature DC differentiate and become active in the uptake and processing of antigens, but show a very low T-cell stimulatory capacity. Upon appropriate stimulation, DC undergo maturation which leads to an upregulation of major histocompatibility complex (MHC) class II, co-stimulatory molecules CD80/CD86, the production of IL-12, TNF-a, and alterations in migratory behavior (29, 30). Testicular DC have received little attention despite the possibility that they may play an important role in testicular immune privilege and autoimmune-based male infertility. Although the presence of DC in the normal testis of mice (31, 32), rats (33), and humans (34, 35) has been previously reported, it is only recently with the use of newly available antibodies against DC-specific markers (Ox62 and CD11c) that there has been an unequivocal demonstration of the existence of testicular DC (36). DC numbers were found to be roughly one tenth of those of macrophages in rat testis and orchitis (37). Expression patterns of

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the co-stimulatory molecules CD80 and CD86 suggest that DC-triggered activation of naïve T lymphocytes may eventually occur in the normal rodent testis (38). A supposition is supported by the finding that, compared with DC from untreated animals, DC from orchitis (EAO) animals enhanced naϊve T-cell proliferation in vitro. Although the role of DC in the testis is far from being understood, these results suggest that DC in unaffected testis are functionally tolerogenic and not in an immature state. When faced with a challenge as in experimental autoimmune orchitis (EAO), however, they develop a mature immunogenic state, ready to transfer to draining lymph nodes and amplify immune responses against testicular antigens (38). The mechanism of how DC participate in the activation of autoimmune response in the testis and the subsequent damage of testicular tissue could be explained by the “danger model” (39). This model proposes that stressed and necrotic cells release “danger” signals, mostly in the form of heat shock proteins (Hsp), that enhance the maturation of DC which in turn can trigger autoimmunity (40). With the identification of Hsp 60 and Hsp70 as testicular autoantigens in EAO (41), we hypothesize that immature DC, normally involved in maintaining immune privilege, mature under inflammatory pathological conditions and overcome immune privilege/tolerance by the local activation and expansion of autoreactive T cells. In EAO, DC maturation may be enhanced by the increased levels of TNF-a as this cytokine is known to serve as an important factor influencing the maturation and migration of DC in inflamed tissue (42, 43).

4. Special Role of the Testicular Macrophages

Representing the largest population of leukocytes in the testis, macrophages are found exclusively in the interstitium of the normal mammalian testis and have been shown to play a central role in the establishment and maintenance of the immune privilege of the testis. This pivotal role of macrophages was first indicated in studies where isolated testicular macrophages were found to exhibit immunosuppressive characteristics and an inhibited capacity to produce IL-1b, IL-6, and TNF-a (44–46). Testicular macrophages are a heterogeneous mix, the largest group ED2+ positive macrophages have a substantially reduced ability to mount immune responses. Being mainly trophic in function, their immune regulatory tasks preserve immune privilege (47). In contrast, the small subset of ED1+ “inflammatory” macrophages/monocytes is able to respond to LPS challenge with increased expression of immune mediators such as MCP-1 and iNOS (48, 49). The importance of

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the ED1+ subgroup is illustrated by the observation that these cells are able to migrate into the acute and chronically inflamed testis thus shifting the macrophage balance, and hence the spectrum of produced cytokines, in favor of an inflammatory response with the potential to overcome the immune privilege (37, 42, 48, 49). Recent studies suggest that the presence of a third macrophage population in mouse testis requires IL-13 stimulation for its immunosuppressive function (50).

5. T Lymphocytes In contrast to other immune-privileged organs such as the anterior chamber of the eye, the testis is well connected to afferent lymph nodes, which excludes the possibility that restricted access of T cells accounts for the presence of immune privilege. Approximately 15% of the immune cells in the normal adult rat testis have been shown to be T lymphocytes, predominantly CD8+ T cells, whereas B cells are not present (51–53). T-cell numbers are directly and indirectly (via macrophages) regulated by Leydig cells, a process that appears to involve testosterone and lymphocyte receptor molecule VCAM-1 (51, 53–55). There is clear evidence that lymphocyte numbers increase in the testes of men with infertility and sperm autoimmunity (56, 57) as in rodent models of EAO (58). In EAO, the number of CD4+ and CD8+ effector T lymphocytes dramatically increase at the onset of disease. However, during disease progression, CD4+ effector cells decline in number suggesting an involvement in maintenance of the chronic phase of EAO, while the CD8+ subset remains unchanged (59). Interestingly, both subsets contain regulatory T cells (Tregs) which can induce apoptosis of APC such as DC, macrophages, and B cells, and inhibit their activation and function. In this way they can regulate any subsequent innate and/or adaptive immune response (60). The critical role of T cells is further emphasized by the involvement of surveying CD8+ memory T cells in immunosuppressive mechanisms which are present at immune-privileged sites (61). Taken as a whole, these observations suggest that testicular immune privilege and its disturbance may be at least, partly attributable, to a localized phenomenon which affects T-cell activation and maturation events. However, the situation appears to be further complicated by the finding that increased numbers of Treg fail to effectively suppress inflammation in EAO (59). Potential explanations for the manner in which various mechanisms of immune regulation by Treg may have been lost are inadequate numbers of Treg cells, a Treg cell intrinsic functional defect, or the presence of pathogenic T effector cells

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resistant to Tregs control (62). Moreover, Treg function can be impaired by high levels of TNF-a, IL-6, and IL-12 that are present in EAO rats (38, 63).

6. The Immunosuppressive Function of Androgens

Testosterone, the main androgen produced by Leydig cells, has long been acknowledged as the crucial hormone for the initiation and maintenance of spermatogenesis. Recent findings, accumulated from both clinical and experimental studies, also strongly indicate an immunosuppressive role that appears to act on multiple levels. Testosterone treatment of macrophages causes a decrease in Toll-like receptor 4 (TLR4) expression with concomitant loss of sensitivity to the TLR4 ligand LPS. An observation that is supported by the finding that castration leads to enhanced synthesis of TLR4 on macrophages and elevated susceptibility to septic shock (64). Although classical androgen receptors do not appear to be expressed by macrophages, studies by Benten at al. suggest the existence of non-genomic cell surface receptors for testosterone (65). This could explain how testosterone downregulates the stimulated activation of pro-inflammatory transcriptional regulators, cytokines (IL-1, IL-6, and TNF-a), and adhesion molecules in isolated macrophages and non-immune cells (66–70). The purported immunosuppressive role of androgens is further substantiated by the finding that men receiving acyline, a gonadotropin-releasing hormone antagonist, show a significantly reduced percentage of CD4+CD25+ Treg cells, decreased mitogen-induced CD8+ T-cell IFN-g expression, and increased percentage of NK cells in their peripheral blood (71). Suggesting that testosterone may help to maintain the physiological balance between autoimmunity and tolerance by controlling the number and activity of the Treg population. Direct evidence for a link between testosterone levels and testicular immune privilege was found when rats pretreated with estrogen to suppress Leydig cell testosterone production rapidly rejected intratesticular allografts, whereas they were tolerated for much longer period in untreated controls (33). These studies indicate that the uniquely high local testosterone concentration in the testis, tenfold above the serum levels, must also be considered a pertinent contributor to the maintenance of testicular immune privilege. This is further underlined by own unpublished results which show that testosterone replacement during the development of EAO in rats significantly reduces the condition’s characteristic pathological symptoms such as leukocytic infiltrations and elevated pro-inflammatory mediators (TNF-a, IL-6, and MCP-1).

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7. Challenging Immune Privilege by Pathological Implications

Despite its status as an immune-privileged organ, testicular function can be impaired by local and systemic infection and inflammation. Disruption of the prevailing immune status affects both steroidogenesis and spermatogenesis and can lead to temporary or sometimes permanent infertility. Orchitis alone is a rare entity and mainly seen as a consequence of systemic viral infection of paramyxovirus (mumps orchitis) and human immunodeficiency virus (HIV). The most common complication of the mumps virus is testicular atrophy (72). Bacterial infection of the testis is usually observed as a combined epididymo-orchitis caused by sexually transmitted diseases or ascending urinary tract infections. The most common sexually transmitted causes of epididymo-orchitis being Neisseria gonorrhoeae, Chlamydia trachomatis, and Ureaplasma urealyticum, while Escherichia coli, Pseudomonas aeruginosa, and Enterobacter aerogenes are the most frequently encountered urinary tract pathogens. The innate immune response against invading microorganism depends on the recognition of conserved molecules by the Tolllike receptors (TLR). TLR sense characteristic microbial patterns such as double-stranded viral RNA (TLR3) or bacterial molecules like lipopolysaccharide (TLR4) and peptidoglycan (TLR6). This leads to the activation of signaling pathways which ultimately triggers the secretion of pro-inflammatory cytokines or the expression of anti-viral response genes depending on the type of stimulated TLR (73). Recently, TLR expression was found in all testicular cells, with testicular macrophages, Sertoli cells, and DC showing the broadest range of TLR expression (74–76). This finding strengthens the view that not only immune cells, but also Sertoli cells have an eminent role in protecting the testis from bacterial and viral infections. Further corroboration was found when mouse Sertoli cells were challenged with TLR2–TLR5 ligands in vitro resulted in strong anti-inflammatory and anti-viral responses in the form of the upregulation of IL-1a, IL-6, INF-a, and INF-b, while germ cells responded weakly (77–80). There is no compelling evidence to suggest that the immune system of the testis and its reproductive function are separate entities. On the contrary, a dual function of cells and factors in both systems is becoming increasingly apparent. The extent of this overlap is exemplified by the Sertoli cells which were originally known only for their association with the reproductive function of the testis, but are now also recognized as important contributors to the maintenance of immune privilege and host defense. Conversely, a classical immune cell, like the macrophage, in the testis is capable of influencing essential reproductive functions such as testosterone secretion in Leydig cells (81). The insights of the synergistic importance in the testis of the immune system, spermatogenesis,

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and steroidogenesis present a peek into what promises to be a fascinating new and emerging field of investigation.

Acknowledgments The grant support of the Deutsche Forschungsgemeinschaft (DFG Clinical Research Unit KFO 181, Me 1323/4-4) is gratefully acknowledged. The authors are grateful to Dr. Con Mallidis (Münster, Germany) for linguistic help. References 1. Head, J.R., Neaves, W.B., Billingham, R.E., 1983. Reconsideration of the lymphatic drainage of the rat testis. Transplantation 35, 91–95. 2. de Kretser, D.M., Kerr, J.B. (1994). The cytology of the testis. In: Physiology of Reproduction, eds. E. Knobil and J. Neill, New York: Raven, 1177–1300. 3. de Kretser, D.M., Loveland, K.L., Meinhardt, A., Simorangkir, D., Wreford, N., 1998. Spermatogenesis. Hum Reprod 13 Suppl 1, 1–8. 4. De Gendt, K., Swinnen, J.V., Saunders, P.T., Schoonjans, L., Dewerchin, M., Devos, A., Tan, K., Atanassova, N., Claessens, F., Lecureuil, C., Heyns, W., Carmeliet, P., Guillou, F., Sharpe, R.M., Verhoeven, G., 2004. A Sertoli cell-selective knockout of the androgen receptor causes spermatogenic arrest in meiosis. Proc Natl Acad Sci USA 101, 1327–1332. 5. Welsh, M., Saunders, P.T., Atanassova, N., Sharpe, R.M., Smith, L.B., 2009. Androgen action via testicular peritubular myoid cells is essential for male fertility. FASEB J 23(12):4218–4230. 6. Wang, C.Q., Cheng, C.Y., 2007. A seamless trespass: germ cell migration across the seminiferous epithelium during spermatogenesis. J Cell Biol 178, 549–556. 7. Filippini, A., Riccioli, A., Padula, F., Lauretti, P., D’Alessio, A., De Cesaris, P., Gandini, L., Lenzi, A., Ziparo, E., 2001. Control and impairment of immune privilege in the testis and in semen. Hum Reprod Update 7, 444–449. 8. Fijak, M., Meinhardt, A., 2006. The testis in immune privilege. Immunol Rev 213, 66–81. 9. Head, J.R., Neaves, W.B., Billingham, R.E., 1983. Immune privilege in the testis. I. Basic parameters of allograft survival. Transplantation 36, 423–431.

10. Itoh, M., Xie, Q., Miyamoto, K., Takeuchi, Y., 1999. Major differences between the testis and epididymis in the induction of granulomas in response to extravasated germ cells. I. A light microscopical study in mice. Int J Androl 22, 316–323. 11. Naito, M., Sakamoto, Y., Terayama, H., Hirai, S., Ning, Q., Aota, Y., Itoh, M., 2009. Effect of unilateral testicular rupture on histopathology and germ cell delayed-type hypersensitivity in C3H/He and A/J mice. J Reprod Immunol 81, 55–61. 12. Mahi-Brown, C.A., Yule, T.D., Tung, K.S., 1988. Evidence for active immunological regulation in prevention of testicular autoimmune disease independent of the blood-testis barrier. Am J Reprod Immunol Microbiol 16, 165–170. 13. Yule, T.D., Montoya, G.D., Russell, L.D., Williams, T.M., Tung, K.S., 1988. Autoantigenic germ cells exist outside the blood testis barrier. J Immunol 141, 1161–1167. 14. Johnson, M.H., 1970. Changes in the bloodtestis barrier of the guinea-pig in relation to histological damage following iso-immunization with testis. J Reprod Fertil 22, 119–127. 15. Nistal, M., Riestra, M.L., Paniagua, R., 2002. Focal orchitis in undescended testes: discussion of pathogenetic mechanisms of tubular atrophy. Arch Pathol Lab Med 126, 64–69. 16. Naito, M., Itoh, M., 2008. Patterns of infiltration of lymphocytes into the testis under normal and pathological conditions in mice. Am J Reprod Immunol 59, 55–61. 17. Spiess, A.N., Feig, C., Schulze, W., Chalmel, F., Cappallo-Obermann, H., Primig, M., Kirchhoff, C., 2007. Cross-platform gene expression signature of human spermatogenic failure reveals inflammatory-like response. Hum Reprod 22, 2936–2946.

468

Fijak, Bhushan, and Meinhardt

18. Suarez-Pinzon, W., Korbutt, G.S., Power, R., Hooton, J., Rajotte, R.V., Rabinovitch, A., 2000. Testicular sertoli cells protect islet beta-cells from autoimmune destruction in NOD mice by a transforming growth factor-beta1-dependent mechanism. Diabetes 49, 1810–1818. 19. De Cesaris, P., Filippini, A., Cervelli, C., Riccioli, A., Muci, S., Starace, G., Stefanini, M., Ziparo, E., 1992. Immunosuppressive molecules produced by Sertoli cells cultured in vitro: biological effects on lymphocytes. Biochem Biophys Res Commun 186, 1639–1646. 20. Sipione, S., Simmen, K.C., Lord, S.J., Motyka, B., Ewen, C., Shostak, I., Rayat, G.R., Dufour, J.M., Korbutt, G.S., Rajotte, R.V., Bleackley, R.C., 2006. Identification of a novel human granzyme B inhibitor secreted by cultured sertoli cells. J Immunol 177, 5051–5058. 21. Yin, Z.Z., Xie, L., Zeng, M.H., Li, R., Huang, Y.B., Zhu, M., Chen, G., Chen, S., 2006. Sertoli cells induce xenolymphocyte apoptosis in vitro. Transplant Proc 38, 3309–3311. 22. Strasser, A., Jost, P.J., Nagata, S., 2009. The many roles of FAS receptor signaling in the immune system. Immunity 30, 180–192. 23. Francavilla, S., D’Abrizio, P., Rucci, N., Silvano, G., Properzi, G., Straface, E., Cordeschi, G., Necozione, S., Gnessi, L., Arizzi, M., Ulisse, S., 2000. Fas and Fas ligand expression in fetal and adult human testis with normal or deranged spermatogenesis. J Clin Endocrinol Metab 85, 2692–2700. 24. Theas, S., Rival, C., Lustig, L., 2003. Germ cell apoptosis in autoimmune orchitis: involvement of the Fas-FasL system. Am J Reprod Immunol 50, 166–176. 25. Kajihara, T., Okagaki, R., Ishihara, O., 2006. LPS-induced transient testicular dysfunction accompanied by apoptosis of testicular germ cells in mice. Med Mol Morphol 39, 203–208. 26. Kim, S.K., Yoon, Y.D., Park, Y.S., Seo, J.T., Kim, J.H., 2007. Involvement of the Fas-Fas ligand system and active caspase-3 in abnormal apoptosis in human testes with maturation arrest and Sertoli cell-only syndrome. Fertil Steril 87, 547–553. 27. O’Bryan, M.K., Hedger, M.P., 2009. Inflammatory networks in the control of spermatogenesis: chronic inflammation in an immunologically privileged tissue? Adv Exp Med Biol 636, 92–114. 28. Banchereau, J., Steinman, R.M., 1998. Dendritic cells and the control of immunity. Nature 392, 245–252. 29. Banchereau, J., Briere, F., Caux, C., Davoust, J., Lebecque, S., Liu, Y.J., Pulendran, B., Palucka, K., 2000. Immunobiology of dendritic cells. Annu Rev Immunol 18, 767–811.

30. Hackstein, H., Thomson, A.W., 2004. Dendritic cells: emerging pharmacological targets of immunosuppressive drugs. Nat Rev Immunol 4, 24–34. 31. Itoh, M., De Rooij, D.G., Jansen, A., Drexhage, H.A., 1995. Phenotypical heterogeneity of testicular macrophages/dendritic cells in normal adult mice: an immunohistochemical study. J Reprod Immunol 28, 217–232. 32. Hoek, A., Allaerts, W., Leenen, P.J., Schoemaker, J., Drexhage, H.A., 1997. Dendritic cells and macrophages in the pituitary and the gonads. Evidence for their role in the fine regulation of the reproductive endocrine response. Eur J Endocrinol 136, 8–24. 33. Head, J.R., Billingham, R.E., 1985. Immune privilege in the testis. II. Evaluation of potential local factors. Transplantation 40, 269–275. 34. Haas, G.G., Jr., D’Cruz, O.J., De Bault, L.E., 1988. Distribution of human leukocyte antigen-ABC and -D/DR antigens in the unfixed human testis. Am J Reprod Immunol Microbiol 18, 47–51. 35. Derrick, E.K., Barker, J.N., Khan, A., Price, M.L., Macdonald, D.M., 1993. The tissue distribution of factor XIIIa positive cells. Histopathology 22, 157–162. 36. Rival, C., Lustig, L., Iosub, R., Guazzone, V.A., Schneider, E., Meinhardt, A., Fijak, M., 2006. Identification of a dendritic cell population in normal testis and in chronically inflamed testis of rats with autoimmune orchitis. Cell Tissue Res 324(2):311–318. 37. Rival, C., Theas, M.S., Suescun, M.O., Jacobo, P., Guazzone, V., van Rooijen, N., Lustig, L., 2008. Functional and phenotypic characteristics of testicular macrophages in experimental autoimmune orchitis. J Pathol 215, 108–117. 38. Rival, C., Guazzone, V.A., von Wulffen, W., Hackstein, H., Schneider, E., Lustig, L., Meinhardt, A., Fijak, M., 2007. Expression of co-stimulatory molecules, chemokine receptors and proinflammatory cytokines in dendritic cells from normal and chronically inflamed rat testis. Mol Hum Reprod 13, 853–861. 39. Matzinger, P., 1994. Tolerance, danger, and the extended family. Annu Rev Immunol 12, 991–1045. 40. Millar, D.G., Garza, K.M., Odermatt, B., Elford, A.R., Ono, N., Li, Z., Ohashi, P.S., 2003. Hsp70 promotes antigen-presenting cell function and converts T-cell tolerance to autoimmunity in vivo. Nat Med 9, 1469–1476. 41. Fijak, M., Iosub, R., Schneider, E., Linder, M., Respondek, K., Klug, J., Meinhardt, A., 2005. Identification of immunodominant autoantigens in rat autoimmune orchitis. J Pathol 207, 127–138.

Immunoprivileged Sites: The Testis 42. Suescun, M.O., Rival, C., Theas, M.S., Calandra, R.S., Lustig, L., 2003. Involvement of tumor necrosis factor-alpha in the pathogenesis of autoimmune orchitis in rats. Biol Reprod 68, 2114–2121. 43. Iizasa, H., Yoneyama, H., Mukaida, N., Katakoka, Y., Naito, M., Yoshida, N., Nakashima, E., Matsushima, K., 2005. Exacerbation of granuloma formation in IL-1 receptor antagonist-deficient mice with impaired dendritic cell maturation associated with Th2 cytokine production. J Immunol 174, 3273–3280. 44. Kern, S., Maddocks, S., 1995. Indomethacin blocks the immunosuppressive activity of rat testicular macrophages cultured in vitro. J Reprod Immunol 28, 189–201. 45. Kern, S., Robertson, S.A., Mau, V.J., Maddocks, S., 1995. Cytokine secretion by macrophages in the rat testis. Biol Reprod 53, 1407–1416. 46. Bryniarski, K., Szczepanik, M., Maresz, K., Ptak, M., Ptak, W., 2004. Subpopulations of mouse testicular macrophages and their immunoregulatory function. Am J Reprod Immunol 52, 27–35. 47. Hedger, M.P., 2002. Macrophages and the immune responsiveness of the testis. J Reprod Immunol 57, 19–34. 48. Gerdprasert, O., O’Bryan, M.K., Muir, J.A., Caldwell, A.M., Schlatt, S., de Kretser, D.M., Hedger, M.P., 2002. The response of testicular leukocytes to lipopolysaccharide-induced inflammation: further evidence for heterogeneity of the testicular macrophage population. Cell Tissue Res 308, 277–285. 49. Gerdprasert, O., O’Bryan, M.K., NikolicPaterson, D.J., Sebire, K., de Kretser, D.M., Hedger, M.P., 2002. Expression of monocyte chemoattractant protein-1 and macrophage colony-stimulating factor in normal and inflamed rat testis. Mol Hum Reprod 8, 518–524. 50. Maresz, K., Ponomarev, E.D., Barteneva, N., Tan, Y., Mann, M.K., Dittel, B.N., 2008. IL-13 induces the expression of the alternative activation marker Ym1 in a subset of testicular macrophages. J Reprod Immunol 78, 140–148. 51. Wang, J., Wreford, N.G., Lan, H.Y., Atkins, R., Hedger, M.P., 1994. Leukocyte populations of the adult rat testis following removal of the Leydig cells by treatment with ethane dimethane sulfonate and subcutaneous testosterone implants. Biol Reprod 51, 551–561. 52. Hedger, M.P., 1997. Testicular leukocytes: what are they doing? Rev Reprod 2, 38–47. 53. Hedger, M.P., Meinhardt, A., 2000. Local regulation of T cell numbers and lymphocyteinhibiting activity in the interstitial tissue of the adult rat testis. J Reprod Immunol 48, 69–80.

469

54. Sainio-Pollanen, S., Sundstrom, J., Erkkila, S., Hanninen, A., Vainiopaa, M., Martikainen, M., Salminen, E., Verajankorva, E., Antola, H., Nikula, H., Simell, O., Pollanen, P., 1997. CD106 (VCAM-1) in testicular immunoregulation. J Reprod Immunol 33, 221–238. 55. Meinhardt, A., Bacher, M., Metz, C., Bucala, R., Wreford, N., Lan, H., Atkins, R., Hedger, M., 1998. Local regulation of macrophage subsets in the adult rat testis: examination of the roles of the seminiferous tubules, testosterone, and macrophage-migration inhibitory factor. Biol Reprod 59, 371–378. 56. el-Demiry, M.I., Hargreave, T.B., Busuttil, A., James, K., Ritchie, A.W., Chisholm, G.D., 1985. Lymphocyte sub-populations in the male genital tract. Br J Urol 57, 769–774. 57. el-Demiry, M.I., Hargreave, T.B., Busuttil, A., Elton, R., James, K., Chisholm, G.D., 1987. Immunocompetent cells in human testis in health and disease. Fertil Steril 48, 470–479. 58. Lustig, L., Lourtau, L., Perez, R., Doncel, G.F., 1993. Phenotypic characterization of lymphocytic cell infiltrates into the testes of rats undergoing autoimmune orchitis. Int J Androl 16, 279–284. 59. Jacobo, P., Guazzone, V.A., Jarazo-Dietrich, S., Theas, M.S., Lustig, L., 2009. Differential changes in CD4(+) and CD8(+) effector and regulatory T lymphocyte subsets in the testis of rats undergoing autoimmune orchitis. J Reprod Immunol 81(1):44–54. 60. Andre, S., Tough, D.F., Lacroix-Desmazes, S., Kaveri, S.V., Bayry, J., 2009. Surveillance of antigen-presenting cells by CD4+ CD25+ regulatory T cells in autoimmunity: immunopathogenesis and therapeutic implications. Am J Pathol 174, 1575–1587. 61. Dai, Z., Nasr, I.W., Reel, M., Deng, S., Diggs, L., Larsen, C.P., Rothstein, D.M., Lakkis, F.G., 2005. Impaired recall of CD8 memory T cells in immunologically privileged tissue. J Immunol 174, 1165–1170. 62. Schneider, A., Rieck, M., Sanda, S., Pihoker, C., Greenbaum, C., Buckner, J.H., 2008. The effector T cells of diabetic subjects are resistant to regulation via CD4+ FOXP3+ regulatory T cells. J Immunol 181, 7350–7355. 63. Theas, M.S., Rival, C., Jarazo-Dietrich, S., Jacobo, P., Guazzone, V.A., Lustig, L., 2008. Tumour necrosis factor-alpha released by testicular macrophages induces apoptosis of germ cells in autoimmune orchitis. Hum Reprod 23, 1865–1872. 64. Rettew, J.A., Huet-Hudson, Y.M., Marriott, I., 2008. Testosterone reduces macrophage expression in the mouse of toll-like receptor 4,

470

65.

66.

67.

68.

69.

70.

71.

72.

73.

74.

75.

Fijak, Bhushan, and Meinhardt a trigger for inflammation and innate immunity. Biol Reprod 78, 432–437. Benten, W.P., Lieberherr, M., Stamm, O., Wrehlke, C., Guo, Z., Wunderlich, F., 1999. Testosterone signaling through internalizable surface receptors in androgen receptor-free macrophages. Mol Biol Cell 10, 3113–3123. Li, Z.G., Danis, V.A., Brooks, P.M., 1993. Effect of gonadal steroids on the production of IL-1 and IL-6 by blood mononuclear cells in vitro. Clin Exp Rheumatol 11, 157–162. Gornstein, R.A., Lapp, C.A., Bustos-Valdes, S.M., Zamorano, P., 1999. Androgens modulate interleukin-6 production by gingival fibroblasts in vitro. J Periodontol 70, 604–609. Liva, S.M., Voskuhl, R.R., 2001. Testosterone acts directly on CD4+ T lymphocytes to increase IL-10 production. J Immunol 167, 2060–2067. Hatakeyama, H., Nishizawa, M., Nakagawa, A., Nakano, S., Kigoshi, T., Uchida, K., 2002. Testosterone inhibits tumor necrosis factoralpha-induced vascular cell adhesion molecule-1 expression in human aortic endothelial cells. FEBS Lett 530, 129–132. Benten, W.P., Guo, Z., Krucken, J., Wunderlich, F., 2004. Rapid effects of androgens in macrophages. Steroids 69, 585–590. Page, S.T., Plymate, S.R., Bremner, W.J., Matsumoto, A.M., Hess, D.L., Lin, D.W., Amory, J.K., Nelson, P.S., Wu, J.D., 2006. Effect of medical castration on CD4+ CD25+ T cells, CD8+ T cell IFN-gamma expression, and NK cells: a physiological role for testosterone and/or its metabolites. Am J Physiol Endocrinol Metab 290, E856–E863. Bartak, V., 1973. Sperm count, morphology and motility after unilateral mumps orchitis. J Reprod Fertil 32, 491–494. Akira, S., Uematsu, S., Takeuchi, O., 2006. Pathogen recognition and innate immunity. Cell 124, 783–801. Girling, J.E., Hedger, M.P., 2007. Toll-like receptors in the gonads and reproductive tract: emerging roles in reproductive physiology and pathology. Immunol Cell Biol 85, 481–489. Palladino, M.A., Johnson, T.A., Gupta, R., Chapman, J.L., Ojha, P., 2007. Members of the Toll-like receptor family of innate immunity pattern-recognition receptors are abundant in the male rat reproductive tract. Biol Reprod 76, 958–964.

76. Bhushan, S., Tchatalbachev, S., Klug, J., Fijak, M., Pineau, C., Chakraborty, T., Meinhardt, A., 2008. Uropathogenic Escherichia coli block MyD88-dependent and activate MyD88independent signaling pathways in rat testicular cells. J Immunol 180, 5537–5547. 77. Dejucq, N., Dugast, I., Ruffault, A., van der Meide, P.H., Jegou, B., 1995. Interferonalpha and -gamma expression in the rat testis. Endocrinology 136, 4925–4931. 78. Dejucq, N., Lienard, M.O., Guillaume, E., Dorval, I., Jegou, B., 1998. Expression of interferons-alpha and -gamma in testicular interstitial tissue and spermatogonia of the rat. Endocrinology 139, 3081–3087. 79. Melaine, N., Lienard, M.O., Guillaume, E., Ruffault, A., Dejucq-Rainsford, N., Jegou, B., 2003. Production of the antiviral proteins 2¢5¢oligoadenylate synthetase, PKR and Mx in interstitial cells and spermatogonia. J Reprod Immunol 59, 53–60. 80. Wu, H., Wang, H., Xiong, W., Chen, S., Tang, H., Han, D., 2008. Expression patterns and functions of toll-like receptors in mouse sertoli cells. Endocrinology 149, 4402–4412. 81. Nes, W.D., Lukyanenko, Y.O., Jia, Z.H., Quideau, S., Howald, W.N., Pratum, T.K., West, R.R., Hutson, J.C., 2000. Identification of the lipophilic factor produced by macrophages that stimulates steroidogenesis. Endocrinology 141, 953–958. 82. Hales, D.B., Diemer, T., Hales, K.H., 1999. Role of cytokines in testicular function. Endocrine 10:201–217. 83 Huleihel, M., Lunenfeld, E., 2004. Regulation of spermatogenesis by paracrine/ autocrine testicular factors. Asian J Androl 6: 259–268. 84. Siu, M.K., Lee, W.M., Cheng, C.Y., 2003. The interplay of collagen IV, tumor necrosis factor-alpha, gelatinase B (matrix metalloprotease-9), and tissue inhabitor-of metalloproteases-1 in the basal lamina regulates Sertoli cell-tight junction dynamics in the rat testis. Endocrinology 144:371–387. 85. Sarkar, O., Mathur, P.P., Cheng, C.Y., Mruk DD., 2008. Interleukin 1 alpha (IL1A) is a novel regulator of the blood-testis barrier in the rat. Biol Reprod 78:445–454. 86. Wong, C.H., Cheng, C.Y., 2005. Mitogenactivated protein kinases, adherenes junction dynamics, and spermatogenesis: a review of recent data. Dev Biol 286:1–15.

INDEX

A Activation/expansion ..........................................28, 85, 437 a-Galactosylceramide (a-GalCer) .........194, 195, 199–205 a-GalCer. See a-Galactosylceramide Allergy ......................................................66, 254, 274, 316 Amino acid starvation ....................................270, 274, 275 Andrographolide ............................340–342, 344, 347–351 Anterior chamber (AC) .......... 449–452, 454, 456, 460, 464 Antigen presentation ........................... 69, 74–76, 128, 129, 144, 225, 398, 401, 435, 441, 450, 455 presenting cells .............................. 6, 27, 32, 54–56, 86, 91, 165–167, 272, 367–368, 398–399, 420, 450, 462 Apoptosis................................7, 77, 86, 157, 170, 222, 254, 274, 281, 296, 313, 314, 317, 325, 360–365, 381, 384, 399, 410, 453, 464 Arginase ................................. 170, 376, 377, 380, 381, 385 Autoimmunity ................................... 7, 8, 10, 15, 100, 149, 150, 162, 272, 273, 277, 282, 307, 309, 310, 319–321, 340–343, 385, 403, 427, 437, 461, 463–465

B B cells .......................................22, 51, 70, 73, 99–110, 194, 197, 204, 208, 241, 285, 314, 319, 365, 384, 401, 425, 426, 432, 453, 464 BMCMCs. See Bone marrow-derived cultured mast cell BMMCs ........................................................................ 207 Bone marrow-derived cultured mast cell (BMCMCs)......................................... 208–219

C Cancer ..............................66, 149, 170, 276, 314, 322–326, 356, 357, 360, 361, 365 Carbon monoxide .................................................. 376, 406 5-,6-Carboxyfluorescein diacetate succinimidyl ester (CFSE) .................................... 5, 19, 26, 34, 42, 67, 69, 75, 76, 78, 79, 88, 93–95, 183, 187–189, 196, 204 Cellular therapy ........................................16, 149–157, 181

CFSE. See 5-,6-Carboxyfluorescein diacetate succinimidyl ester Colitis ..........................7, 15, 33, 47–59, 273, 313, 324, 379 Culture .................................5, 17, 34, 49, 67, 88, 100, 115, 129, 151, 162, 171, 183, 196, 210, 222, 234, 271, 283, 343, 409 Cytokines ...................................6, 17, 31, 47, 88, 129, 150, 177, 194, 208, 222, 253, 256, 293, 306, 340, 359, 377, 398, 420, 431, 451, 461

D Dendritic cells (DCs) bone marrow-derived.............................88, 91–92, 132, 140, 149–157, 161–167, 285, 293, 346, 367, 462 plasmacytoid dendritic cells (PDC) ...............66, 73–78, 80, 113, 127–145, 204, 272 tolerogenic ................32, 33, 35, 37, 149, 151, 153–156, 160–162, 164, 166, 315, 339–352, 367, 376, 436 Dexamethasone .........32, 150, 151, 154–157, 184, 191, 310

E EAE. See Encephalomyelitis Encephalomyelitis (EAE) ......................... 33, 66, 254, 263, 275, 315, 321, 339, 341, 343, 344, 348–350, 425 Engraftment ............201, 209, 210, 215, 216, 218, 227, 282

F Fcg receptors ...........................................199, 341–343, 345 Foxp3 ............................ 3–11, 15, 31, 47, 66, 114, 129, 209, 258, 274, 283, 307, 359, 376, 404, 436

G Galectins.........................................................355–369, 398 Glycosylation .................................. 355–358, 364, 369, 402

H Heme oxygenase-1 (HO-1)................... 167, 170, 173, 177, 247–264, 398, 399, 405, 407–411 Human ...............................4, 15, 31, 48, 86, 115, 129, 132, 150, 170, 181, 193, 209, 222, 235, 253, 270, 283, 305, 345, 378, 398, 422, 431, 455, 461

Maria Cristina Cuturi and Ignacio Anegon (eds.), Suppression and Regulation of Immune Responses, Methods in Molecular Biology, vol. 677, DOI 10.1007/978-1-60761-869-0, © Springer Science+Business Media, LLC 2011

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SUPPRESSION AND REGULATION OF IMMUNE RESPONSES 472 Index I IDO. See Indoleamine 2,3-dioxygenase IFN-g26 ...........................1, 39–42, 44, 116, 177, 185, 190, 191, 194, 202, 203, 205, 272, 273, 277, 281–298, 306, 307, 312–315, 318, 322, 323, 325, 361, 362, 364–366, 368, 377–384, 403, 408, 425, 432, 434, 437, 438, 442, 451, 452, 454–456, 465 IL-10. See Interleukin-10 Immune privilege.................... 323, 398, 449–456, 459–467 Immunoregulation ..................................435, 436, 439, 441 Immunosuppression ......................... 66, 209, 276, 361, 441 Immunotherapy ........................................33, 169, 170, 314 Indoleamine 2,3-dioxygenase (IDO)......128, 269–277, 282 Infection ...........................15, 128, 170, 253, 269, 296, 315, 375, 407, 431, 462 Inflammation 10 ...............48, 129, 170, 184, 208, 222, 234, 253, 275, 306, 351, 356, 375, 403, 452, 461 Inflammatory bowel disease .................................... 33, 48, 58, 66, 321 conditions .........................................114, 128, 170, 194 Interleukin-10 (IL-10) .......................... 6, 31, 99, 128, 150, 190, 208, 222, 253, 276, 291, 307, 340, 364, 376, 402, 433, 450 Isolation ............................16–22, 28, 35, 49, 51, 67, 86, 87, 89, 91, 93, 103, 106, 108, 109, 151, 152, 154, 173, 185, 188, 194–195, 197–199, 201, 222–224, 226, 228, 240, 258, 343–345, 347

K

Neural progenitor .................................................. 233–242 Neural stem cell ..................................................... 233–242 NF-kB ........................................... 272, 274, 277, 339–352, 420, 424 Nitric oxide (NO) .......................... 170, 294, 318, 375–386, 407, 410, 456, 460 Nitric oxide synthase endothelial (eNOS) ..................376–378, 382–385, 410 inducible (iNOS) .......257, 258, 264, 376–385, 410, 463 neuronal (nNOS)...................... 376–378, 382, 383, 385

O Ocular sympathetic innervations ................................... 449 Organ injury .................................................................. 221

P PD-1. See Programmed cell death receptor 1 Persistent infection .................................432–434, 438–442 Pig ......................................................................... 100, 240 Potency assay ......................................................... 221–229 Pregnancy .......................................................272, 396–411 Programmed cell death receptor 1 (PD-1) ............365, 401, 424–426, 433, 434, 437, 438, 441, 442

R Retinoic acid (RA) ....................... 10, 47, 49, 52–54, 56, 58 Rosiglitazone .................................................340, 342, 344, 347–349, 351

S Kynurenines............................................270, 274–275, 376

L LCMV. See Lymphocytic choriomeningitis virus Lipopolysaccharide (LPS) ..................... 37, 38, 88, 92, 100, 103, 105, 106, 109, 170–172, 174–178, 184, 190, 223, 224, 226, 253, 258, 340, 344, 346, 352, 362, 366, 377, 463, 465, 466 Lymphocytic choriomeningitis virus (LCMV) ............ 321, 432–434, 437, 438, 440, 441

M Macrophage ..............................51, 156, 181, 253, 276, 321, 362, 376, 380, 398, 431, 451 Mast cell-deficient mice .................................209–213, 215 Mesenchymal stem cell (MSC) ..............221–229, 257, 264 Mixed leukocyte reaction (MLR) ..................42–44, 66, 67, 69, 74–78, 164–167, 256, 257 Myeloid (-derived) suppressor macrophage (MDSC) ...............................169–179, 378–381

N Natural killer cells (NKT) .............. 193–205, 399, 450, 453

Sertoli cells .....................................................459–462, 466 Steady state conditions ...................................113–125, 128 Suppressive function .................... 42–44, 63, 66, 67, 69, 75, 76, 78, 79, 173, 254, 262, 437, 465 Systemic lupus erythematosus (SLE) ................... 319–320, 342, 345, 349–352

T T cells CD5, 7, 91 CD4+ effector.............................................66, 295, 464 CD8 + regulatory......................................... 63–80, 263 double negative ............................................ 85–98, 201 induced regulatory ....................................................... 3 natural regulatory..................................................... 376 regulatory (Treg) .....................3, 15, 31, 47, 63, 85, 114, 128, 150, 162, 262, 272, 281, 310, 340, 359, 376, 398, 420, 436, 450, 464 suppressor ............................................. 4, 7, 63, 66, 452 therapy ....................................................................... 33 type 1 regulatory T (Tr1) .............. 31–44, 291, 367, 404 Testosterone ...................................................460, 464, 465 TGFb48 ................................. 52–54, 56, 58, 132, 133, 140

SUPPRESSION AND REGULATION OF IMMUNE RESPONSES 473 Index Thrombospondin............................................304, 454, 455 Tolerance ............................4, 16, 31, 63, 85, 113, 127, 149, 161, 170, 181, 193, 207, 255, 269, 303, 339, 361, 397, 424, 436, 453, 460 Transplantation, tolerance ......................129, 422, 423, 427 Transwell analysis ............................................................ 63

Tryptophan catabolism .................................................. 270 Tumor immunity ..................................................323, 355–365 microenvironment ................... 315, 323–325, 355–358, 360–362, 367 Type 1 regulatory T (Tr1) cells .................................. 31–44